Hydroxycinnamic Acid Esters Enzymatic Synthesis and

Research Collection

Doctoral Thesis

Enzymatic Synthesis and Hydrolysis of Linear Alkyl and SterylHydroxycinnamic Acid Esters

Author(s): Schär, Aline Lea

Publication Date: 2016

Permanent Link: https://doi.org/10.3929/ethz-a-010670414

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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

i

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

ii

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.

iii

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

iv

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.

v

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

vi

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

Introduction

1

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

Introduction

2

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.

PART A - Review of Literature

3

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.

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 1: Examples of the hydroxybenzoic acid derivatives (p-hydroxybenzoic acid, left) and

the hydroxycinnamic acid derivatives (p-coumaric acid, right).

Figure 2: Major hydroxycinnamic acids: ferulic acid, p-coumaric acid, caffeic acid, and sinapic

acid (from left to right).

Hydroxycinnamic acids

4

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


5

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


6

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


7

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.

Plant materials Steryl hydroxy-

cinnamates

Individual

content (μg/g)

Total content

(μg/g) Reference

Brown rice Steryl ferulates 262-627b (Miller &

Engel, 2006)

Rice bran Steryl ferulates

Steryl caffeates n.r. n.r.

(Fang et al.,

2003)

Cargo rice, wild rice

Steryl ferulates

Steryl p-coumarates

Steryl sinapate

n.r. n.r.

(Zhu &

Nyström,

2015)

Wild rice kernels

Steryl ferulates

Steryl caffeates

Steryl cinnamates

670-1029

79-182

73-141

850 -1352 (Aladedunye

et al., 2013)

Rye kernels,

wheat kernels,

spelt kernels,

corn kernels

Steryl ferulates

Steryl p-coumarates

0.8a,c

2a,c

<LOQ,c

3.7a,c

92a

142a

92a

158a

(Esche et

al., 2012)

Corn bran and

related fractions

Steryl p-coumarates

Steryl ferulates 3-11a,c 70-540a

(Norton,

1995)

Corn, whole grain Steryl p-coumarates

Steryl ferulates

1.5-6

31-70

(Seitz, 1989)

Wheat, whole grain

Rye, whole grain

Triticale, whole grain

Steryl ferulates 62-123

29

52

(Seitz, 1989)

Canary seeds Steryl caffeates n.r. n.r. (Takagi &

Iida, 1980) a: based on dry matter, b: based on fresh weight, c: content of steryl p-coumarates

corresponding to the plant material, n.r.: not reported, LOQ: limit of quantification


8

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


9

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


10

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


11

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.

.

General enzymatic reactions

12

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:

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.

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

Scheme 1: Michaelis-Menten model.

𝑣0 =𝑣𝑚𝑎𝑥∙[𝑆]

𝐾𝑚+[𝑆] (1)

Figure 3: Initial reaction rate as a function of the substrate

concentration based on the Michaelis-Menten equation.


13

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:

In this case also the definition of the kinetic constants changes, where Ks represents the

substrate binding constant:

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:

In the case of k3 >> k2 where the formation of the acyl-enzyme intermediate is limiting it leads

to:

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

Scheme 2: Adapted Michaelis-Menten model including an enzyme-acyl complex, adapted

from (Zerner & Bender, 1964).

𝑘𝑐𝑎𝑡 =𝑘2𝑘3

𝑘2+𝑘3 (2) 𝐾𝑚 = 𝐾𝑆

𝑘3

𝑘2+𝑘3 (3)

𝑘𝑐𝑎𝑡 = 𝑘3 (4) 𝐾𝑚 = 𝐾𝑆𝑘3

𝑘2 (5)

𝑘𝑐𝑎𝑡 = 𝑘2 (6) 𝐾𝑚 = 𝐾𝑆 (7)


14

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


15

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


16

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).

Enzymatic lipophilization of hydroxycinnamic acids

17

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.


18

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

alcohol

Novozym 435 Solvent-free (Guyot et al., 1997)

Ferulic acid, ethanol, octanol/

ethyl ferulate, octanol, monoolein triolein Novozym 435

t-Butanol, toluene,

solvent-free (triolein)

(Compton et al.,

2000)

Hydroxycinnamic acid and benzoic acid

derivatives, octanol

Novozym 435,

Lipozyme RM IM

C. rugosa lipase

Solvent-free (Stamatis et al.,

1999, 2001)

Ferulic acid, ethanol and

p-methoxycinnamic acid, 2-ethyl hexanol Novozym 435 Isooctane (Lee et al., 2006)

Ferulic acid, pentanol, hexanol and

heptanol

Immobilized lipase

from C. antarctica

Chirazyme L-2 C2

Solvent-free,

continuous system

(Yoshida et al.,

2006)


methanol, ethanol, propanol, butanol,

hexanol, octanol, geraniol

Novozym 435,

Lipozyme RM IM

Ionic liquid or

hexane and acetone

(Katsoura et al.,

2009)

Hydroxycinnamic acid and benzoic acid

derivatives, octanol

Lipases from

R. miehei and

C. antarctica in

modified cellulose

organogels

Solvent-free (Zoumpanioti et al.,

2010)

Ferulic acid, ethanol Steapsin

immobilized on celite DMSO

(Kumar & Kanwar,

2011)

Dihydrocaffeic acid, ferulic acid, caffeic

acid, butanol, hexanol, octanol, decanol,

dodecanol, octadecanol

Novozym 435

Hexane/butanone

mixtures or ionic

liquids

(Yang et al., 2012b;

Yang et al., 2012c)

Methyl p-coumarate, methyl ferulate,

octanol Novozym 435

Deep eutectic

solvent–water binary

mixtures

(Durand et al.,

2013)

Ferulic acid, ethanol R. oryzae lipase on

Fe3O4-chitosan Isooctane or hexane (Kumar et al., 2013)

Methyl caffeate, propanol Novozym 435 Ionic liquid (Pang et al., 2013)

p-Coumaric aid, methanol, ethanol,

propanol, butanol

B. licheniformis

SCD11501 lipase on

celite

Solvent-free (Sharma et al.,

2014)

Caffeic acid, 2-pentanol, 2-heptanol,

2-octanol Novozym 435 Isooctane (Xiao et al., 2014)

Ferulic acid, ethanol, dodecanol Novozym 435 Diisopropyl ether (Sandoval et al.,

2015)

Caffeic acid, methanol Novozym 435

Ionic liquid,

ultrasound

irradiation

(Wang et al.,

2015a)

Esterification of long chain alcohols

Dihydrocaffeic acid, ferulic acid, linolenyl

alcohol Novozym 435

Hexane/2-butanone

75:25 or 65:35 (v/v)

(Sabally et al.,

2005)


methyl or ethyl esters thereof, oleyl

alcohol

Lipozyme RM IM,

Novozym 435 Solvent-free, 80 kPa

(Vosmann et al.,

2006) (Weitkamp et

al., 2006)


19

Table 2 continued:

Substrates Enzymes Solvents / conditions

References

Ferulic acid, oleyl alcohol Novozym 435

Ionic liquid/

isooctane binary

system

(Chen et al., 2011a)

Caffeic acid esterification with phenyl alcohols

Cinnamic acid and hydroxy and methoxy

derivatives, phenylethanol, 4-methoxy

phenylethanol, tyrosol

Novozym 435 t-Butanol (Stevenson et al.,

2007)

Caffeic acid, 2-phenylethanol Novozym 435 Isooctane (Widjaja et al.,

2008)

Methyl caffeate, 2-cyclohexylethanol,

3-cyclohexyl-1-propanol, 4-phenylbutanol,

5-phenylpentanol

Novozym 435 Ionic liquid, 845 hPa (Kurata et al., 2010)

Caffeic acid, 2-phenylethanol, octanol Novozym 435 Isooctane (Chen et al., 2010a;

Chen et al., 2010b)

Caffeic acid, 2-phenylethanol Novozym 435

Continuous

ultrasound-assisted

packed-bed reactor,

in isooctane/

t-butanol 9:1

(Chen et al., 2011b)

Caffeic acid, phenethyl alcohol Novozym 435 Ionic liquid (Ha et al., 2013)

Methyl caffeate, propanol, 2-phenylethanol Novozym 435

Ionic liquid,

continuous flow

microreactor

(Wang et al., 2014;

Wang et al., 2013)

Caffeic acid, 2-phenylethanol Novozym 435 2% DMSO in ionic

liquid (Gu et al., 2014)

Esterification with acylglycerols

Ethyl ferulate, soybean oil Novozym 435 Solvent free

(Laszlo & Compton,

2006; Laszlo et al.,

2003)

p-Hydroxyphenyl acetic acid, p-coumaric

acid, sinapic acid, ferulic acid and

3,4-dihydroxybenzoic acid, triolein

Novozym 435 Hexane/2-butanone

85:15 (v/v) (Safari et al., 2006)

Ethyl ferulate, tributyrine Novozym 435 Toluene (Zheng et al., 2008)

Hydroxycinnamic acid derivatives, ethyl

ferulate, flaxseed oil Novozym 435

Hexane/2-butanone

85:15 or solvent-

free, surfactants

(Karboune et al.,

2008; Sorour et al.,

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)

Ferulic acid, flaxseed oil Novozym 435 Supercritical CO2 (Ciftci & Saldana,

2012)

Ethyl ferulate, glycerol, fish oil Novozym 435 Solvent-free (Yang et al., 2012a)

Ethyl ferulate, distearin, monostearin Novozym 435 Solvent-free,

10 mm Hg

(Sun et al., 2012;

Sun & Zhou, 2014)

Ethyl ferulate, phosphatidylcholine Novozym 435 Toluene/chloroform

9:1 (v/v) (Yang et al., 2013)

p-Coumaric acid, triolein, seal blubber oil,

menhaden oil Novozym 435

Hexane/2-butanone

3:1 (v/v)

(Wang & Shahidi,

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.


20

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.


21

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.


22

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


23

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-

ammoniumbromide microemulsion

Feruloyl esterase

from A. niger (Giuliani et al., 2001)

Cinnamic, p-coumaric, ferulic,

p-hydroxyphenyl propionic acid, 1-octanol,

solvent free

F. oxysporum

esterase, F. solani

cutinase

(Stamatis et al., 2001)

p-Hydroxyphenylacetic acid,

p-hydroxyphenylpropionic, cinnamic acid,

p-coumaric acid, ferulic acid, 1-propanol in

n-hexane/1-propanol/water

F. oxysporum

feruloyl esterase (Topakas et al., 2003)

Methyl ferulate, methyl p-coumarate,

methyl caffeate, methyl sinapate,

1-butanol, L-arabinose in

n-hexane/butanol/water

S. thermophile

feruloyl esterase (Topakas et al., 2005)

Methyl ferulate, methyl p-coumarate,

methyl caffeate, methyl sinapate,

1-butanol, 2-butanol in


CLEAs of A. niger

type A feruloyl

esterase

(Vafiadi et al., 2008a)

Methyl ferulate, 1-butanol in


CLEAs of Ultraflo L,

Depol 670L, Depol

740L

(Vafiadi et al., 2008b)

Methyl ferulate, 1-butanol and 7.5% buffer Depol 740L on

mesoporous silica (Thorn et al., 2011)

Ferulic acid, sinapic acid, caffeic acid,

p-coumaric acid, ethanol, methanol,

1-propanol in buffer

Rice koji enzyme (Hashizume et al.,

2013a)

CLEA: cross-linked enzyme aggregate


24

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.

Esterification of phytosterols

25

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.


26

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,

C14:0, C16:0, C18:0 in hexane

Immobilized C. antarctica

lipase B, Novozym 435 (He et al., 2010)

β-Sitosterol, fish oil in hexane Immobilized T. lanuginosus

lipase, Lipozyme TL IM

(Sengupta & Ghosh,

2011)

Phytosterols, oleic acid in isooctane Candida sp. 99–125

immobilized on textile (Pan et al., 2012)

Phytosterols, lauric acid in hexane

with trehalose addition

C. rugosa lipase on

macroporous resin (Jiang et al., 2013)

β-Sitosterol, fatty acids C2:0-C18:0

in hexane


lipase A (Panpipat et al., 2013)

β-Sitosterol, conjugated linoleic acid

in hexane

Chirazyme L-2 c.-f. C2

(from C. antartica) (Li et al., 2010)

Phytosterols, triglycerides and free

fatty acids from sunflower, rapeseed,

corn, tea seed, linseed and rice

bran; free fatty acids (C16:0, C18:1,

C18:2, C18:3, conjugate linoleic

acid) in isooctane or hexane

C. rugosa lipase

immobilized on various

carriers (functionalized

silica particles or polymer

particles)

(Zheng et al., 2012a;

Zheng et al., 2014;

Zheng et al., 2012b;

Zheng et al., 2013;

Zheng et al., 2012c;

Zheng et al., 2015)

Non-immobilized lipase in solvent system

Canola phytosterols, oleic acid,

methyl oleate in hexane C. rugosa lipase (Villeneuve et al., 2005)

Physotsterols, oleic acid in hexane C. rugosa lipase (Kim & Akoh, 2007)

Phytosterols, caprylic acids in

hexane C. rugosa lipase (Tan et al., 2012)

Phytosterols, docosahexaenoic acid

in hexane Lipoprotein lipase 311 (Tan & Shahidi, 2012b)

Non-immobilized lipase in solvent-free system

Phytosterols, sunflower oil Lipase QLM (Alcaligenes

sp.) (Negishi et al., 2003)

Soybean oil deodorizer distillate,

olive oil deodorizer distillates, refined

olive oil, oleic acid

C. rugosa lipase

(Teixeira et al., 2011,

2012, 2014; Torres et

al., 2007)


27

Table 4 continued :

Substrates, conditions Enzymes References

Solvent-free system under reduced pressure

Cholesterol, sitostanol,

stigmasterol, 5α-cholestan-3β-ol,

methyl oleate, oleic acid, triolein,

methyl fatty acid esters, rapeseed

oil, soybean oil, 20-40 mbar

Immobilized lipases from: T.

lanuginosus, R. miehei, C.

antarctica; non-immobilized

lipases from: C. rugosa lipase,

Carica papaya lipase

(Weber et al., 2001a,

2001b, 2002, 2003)

Wood sterols, sunflower fatty acid

methyl esters, 2 mbar Lipase from P. stutzeri PL-836

(Martinez et al.,

2004)

Phytosterols, fatty acids from butter

oil, 100 mbar C. rugosa lipase on octylsilica (Torrelo et al., 2009)

Phytosterols, tributyrine, ethyl

butyrate, fatty acid ethyl esters

from butter fat, 100-350 mbar

C. rugosa lipase, P. stutzeri

lipase (Torrelo et al., 2012)

Phytosterols, fatty acid from pine

nut, 80 kPa

C. rugosa lipase on Lewatit VP

OC 1600 (No et al., 2013)

Non-conventional reaction medias

Cholesterol, cholestanol, and

sitosterol, fatty acids C22:6, C20:5,

C18:3, C18:2, and 30% water

Pseudomonas sp. lipase (Shimada et al.,

1999)

Sitostanol, C8:0, C10:0, C12:0,

C16:0, C18:0 in supercritical CO2

Lipase from Burkholderia

cepacia, Chirazyme L-1 (King et al., 2001)

β-Sitosterol, C6:0, C8:0, C10:0,

C12:0, conjugated linoleic acid, and

0.3 mL/gsterol water or hexane

C. rugosa lipase (Vu et al., 2004)

Phytosterols, fatty acids C12:0,

C14:0, C16:0, C18:0, C18:1 in

water-in-ionic liquid microemulsion

C. rugosa lipase (Zeng et al., 2015)

Phytosterols, oleic acid in

isooctane, under microwave

irradiation

C. rugosa lipase immobilized on

ZnO nanowires/macroporous

SiO2

(Shang et al., 2015)

Phytosterols, soybean oil in

supercritical CO2


lipase, Novozym 435 (Hu et al., 2015)

Sterol esterases

Dihydrocholesterol, cholesterol,

β-sitosterol, sitosterol, stigmasterol,

ergosterol, butyric acid, oleic acid,

in buffer containing bile salts

Homogenate from hog

pancreas (Swell et al., 1954)

Phytosterols, caprylic acid

sunflower oil, solvent-free Sterol esterase from A. oryzae (Hellner et al., 2010)

Cholesterol, stigmasterol, stearic

acid, in buffer or biphasic hexane-

water system

Cholesterol esterase from

Trichoderma sp. AS59

(Morinaga et al.,

2011)


28

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


29

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


30

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


31

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


32

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


33

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.


34

Table

5:

Overv

iew

of

stu

die

s c

on

ducting c

hem

oen

zym

atic s

tery

l phe

nola

tes s

yn

the

sis

.

Refe

ren

ces

(Chig

ori

mb

o-

Mure

fu e

t al.,

20

09

)

(Tan &

Shahid

i,

20

11

, 2

01

2a

,

20

13

)

(Wang e

t a

l.,

20

15

b)

Substr

ate

ratio r

efe

rs to m

ola

r sub

str

ate

ratio o

f vin

yl phe

nola

te t

o s

tero

l.

Iso

late

d

yie

ld

56

%

44

%

90

%

50

%

80

%

88

%

17

.31

%

23

.64

%

38

.04

%

31

.95

%

21

.56

%

01

.79

%

72

.11

%

27

.47

%

45

.41

%

69

.49

%

Ste

rols

Dih

ydro

cho

leste

rol,

5α

-and

rosta

ne

-

3β

,17

β-d

iol

β-S

itoste

rol (7

6%

pure

with o

ther

ste

rols

)

β-S

itoste

rol (9

0%

with 1

0%

oth

er

ste

rols

)

Vin

yl p

hen

ola

tes

Vin

yl fe

rula

te

Vin

yl fe

rula

te

Vin

yl caff

eate

Vin

yl sin

ap

ate

Vin

yl van

illate

Vin

yl 4

-hydro

xybe

nzoa

te

Vin

yl van

illate

Vin

yl 4

-chlo

roph

enyla

ceta

te

Vin

yl hyd

rocin

nam

ate

Vin

yl 4

-phe

nylb

uty

rate

Vin

yl 5

-phe

nylv

ale

rate

Vin

yl cin

na

mate

Vin

yl m

-coum

ara

te

Vin

yl fe

rula

te

Vin

yl 3,4

-dim

eth

oxycin

nam

ate

Co

nd

itio

ns

10

.1 m

g/m

L a

nd

7.6

m

g/m

L s

tero

id

(26

mM

), s

ubstr

ate

ratio

8.7

:1,

tert

-buty

l-m

eth

yl

eth

er,

45

°C

20

mg/m

L p

hyto

ste

rols

,

substr

ate

ratio 1

:2, he

xane

an

d 2

-bu

tan

on

e (

9:1

, v/v

),

45°C

, 10

da

ys

13

.8 m

g/m

L β

-sitoste

rol,

substr

ate

ratio

1:1

, he

xane

an

d 2

-bu

tan

on

e (

8:2

, v/v

),

55°C

, 24

h

En

zym

e t

yp

e a

nd

am

ou

nt

C. ru

go

sa

lip

ase,

100

mg/m

L

C. ru

go

sa

type

VII,

8%

of

the

tota

l

substr

ate

s w

eig

ht,

6 m

g/m

L

C. ru

go

sa lip

ase,

10

0 U

/mL,

0.0

85

mg/m

L

Feruloyl esterases

35

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.

Feruloyl esterases

36

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

Feruloyl esterases

37

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).

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

Table 6: Classification of microbial feruloyl esterases as proposed by Crepin et al., 2004.

Type A Type B Type C Type D

Example A. niger FaeA M. thermophila

FaeB

T. stipitatus

FaeC

P. fluorescens

XYLD

Hydrolyze

methyl ester of

Ferulic acid,

sinapic acid,

p-coumaric acid

Ferulic acid,

caffeic acid,

p-coumaric acid

Ferulic acid,

caffeic acid,

p-coumaric acid,

sinapic acid

Ferulic acid,

caffeic acid,

p-coumaric acid,

sinapic acid

Release of

diferulic acid Yes (5-5’) No No Yes (5-5’)

Sequence

similarity to Lipase

Acetyl xylan

esterase

Chlorogenate

esterase,

tannase

Xylanase

Content adapted from (Crepin et al., 2004).

Feruloyl esterases

38

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). Overall, these results do not show a

clear trend concerning the lipophilicity, which is probably due to the fact, that the substrates

are too similar and more lipophilic substrates could be explored further.

Finally, in recent studies the activities of two feruloyl esterase from L. plantarum were

characterized (Esteban-Torres et al., 2015; Esteban-Torres et al., 2013). Two esterases with

feruloyl esterase activity were identified and recombinantly produced. Both were

characterized on various substrates, including a series of p-nitrophenyl esters. For both a

maximum activity towards the C4 ester was determined. Quite low activity was measured for

the C12 and C14 and slightly higher again for C16 ester. However, for example the activities

towards trilaurin and ethyl oleate were very small (Esteban-Torres et al., 2015; Esteban-

Torres et al., 2013). Nevertheless, no experiments with long-chain ferulates were conducted

in these studies, neither.

References

39

References

Adlercreutz, P. (2013). Immobilisation and application of lipases in organic media. Chemical Society Reviews, 42(15), 6406-6436.

Aladedunye, F., Przybylski, R., Rudzinska, M., & Klensporf-Pawlik, D. (2013). -Oryzanols of North American Wild Rice (Zizania palustris). Journal of the American Oil Chemists Society, 90(8), 1101-1109.

Aliwan, F. O., Kroon, P. A., Faulds, C. B., Pickersgill, R., & Williamson, G. (1999). Ferulic acid esterase-III from Aspergillus niger does not exhibit lipase activity. Journal of the Science of Food and Agriculture, 79(3), 457-459.

Andreasen, M. F., Kroon, P. A., Williamson, G., & Garcia-Conesa, M. T. (2001a). Esterase activity able to hydrolyze dietary antioxidant hydroxycinnamates is distributed along the intestine of mammals. Journal of Agricultural and Food Chemistry, 49(11), 5679-5684.

Andreasen, M. F., Kroon, P. A., Williamson, G., & Garcia-Conesa, M. T. (2001b). Intestinal release and uptake of phenolic antioxidant diferulic acids. Free Radical Biology and Medicine, 31(3), 304-314.

Belitz, H. D., Grosch, W., & Schieberle, P. (2009). Food chemistry (4th rev. and extended ed., pp. 93-155): Springer.

Benoit, I., Danchin, E. G. J., Bleichrodt, R. J., & de Vries, R. P. (2008). Biotechnological applications and potential of fungal feruloyl esterases based on prevalence, classification and biochemical diversity. Biotechnology Letters, 30(3), 387-396.

Berger, A., Rein, D., Schafer, A., Monnard, I., Gremaud, G., Lambelet, P., & Bertoli, C. (2005). Similar

cholesterol-lowering properties of rice bran oil, with varied -oryzanol, in mildly hypercholesterolemic men. European Journal of Nutrition, 44(3), 163-173.

Bernards, M. A. (2002). Demystifying suberin. Canadian Journal of Botany, 80(3), 227-240.

Bugg, T. D. H. (2012). Methods for Studying Enzymatic Reactions Introduction to Enzyme and Coenzyme Chemistry (pp. 50-76): John Wiley & Sons, Ltd.

Buisman, G. J. H., van Helteren, C. T. W., Kramer, G. F. H., Veldsink, J. W., Derksen, J. T. P., & Cuperus, F. P. (1998). Enzymatic esterifications of functionalized phenols for the synthesis of lipophilic antioxidants. Biotechnology Letters, 20(2), 131-136.

Chaiyasit, W., Elias, R. J., McClements, D. J., & Decker, E. A. (2007). Role of physical structures in bulk oils on lipid oxidation. Critical Reviews in Food Science and Nutrition, 47(3), 299-317.

Chen, B. L., Liu, H. Z., Guo, Z., Huang, J. A., Wang, M. Z., Xu, X. B., & Zheng, L. F. (2011a). Lipase-Catalyzed Esterification of Ferulic Acid with Oleyl Alcohol in Ionic Liquid/Isooctane Binary Systems. Journal of Agricultural and Food Chemistry, 59(4), 1256-1263.

Chen, H.-C., Ju, H. Y., Twu, Y. K., Chen, J. H., Chang, C. M., Liu, Y. C., Chang, C., & Shieh, C. J. (2010a). Optimized enzymatic synthesis of caffeic acid phenethyl ester by RSM. New biotechnology, 27(1), 89-93.

Chen, H.-C., Kuo, C.-H., Twu, Y.-K., Chen, J.-H., Chang, C.-M. J., Liu, Y.-C., & Shieh, C.-J. (2011b). A continuous ultrasound-assisted packed-bed bioreactor for the lipase-catalyzed synthesis of caffeic acid phenethyl ester. Journal of Chemical Technology and Biotechnology, 86(10), 1289-1294.

References

40

Chen, H.-C., Twu, Y.-K., Chang, C.-m. J., Liu, Y.-C., & Shieh, C.-J. (2010b). Optimized synthesis of lipase-catalyzed octyl caffeate by Novozym

® 435. Industrial Crops and Products, 32(3), 522-

526.

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.

Choo, W. S., Birch, E. J., & Stewart, I. (2009). Radical Scavenging Activity of Lipophilized Products from Transesterification of Flaxseed Oil with Cinnamic Acid or Ferulic Acid. Lipids, 44(9), 807-815.

Ciftci, D., & Saldana, M. D. A. (2012). Enzymatic synthesis of phenolic lipids using flaxseed oil and ferulic acid in supercritical carbon dioxide media. Journal of Supercritical Fluids, 72, 255-262.

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). Improved method for the synthesis

of trans-feruloyl--sitostanol. Journal of Agricultural and Food Chemistry, 49(10), 4961-4964.

Couteau, D., McCartney, A. L., Gibson, G. R., Williamson, G., & Faulds, C. B. (2001). Isolation and characterization of human colonic bacteria able to hydrolyse chlorogenic acid. Journal of Applied Microbiology, 90(6), 873-881.

Couto, J., Karboune, S., & Mathew, R. (2010). Regioselective synthesis of feruloylated glycosides using the feruloyl esterases expressed in selected commercial multi-enzymatic preparations as biocatalysts. Biocatalysis and Biotransformation, 28(4), 235-244.

Couto, J., St-Louis, R., & Karboune, S. (2011). Optimization of feruloyl esterase-catalyzed synthesis of feruloylated oligosaccharides by response surface methodology. Journal of Molecular Catalysis B: Enzymatic, 73(1-4), 53-62.

Crepin, V. F., Faulds, C. B., & Connerton, I. F. (2003). A non-modular type B feruloyl esterase from Neurospora crassa exhibits concentration-dependent substrate inhibition. Biochemical Journal, 370, 417-427.

Crepin, V. F., Faulds, C. B., & Connerton, I. F. (2004). Functional classification of the microbial feruloyl esterases. Applied Microbiology and Biotechnology, 63(6), 647-652.

D'Ambrosio, M. (2013). Performances of CN-columns for the analysis of -oryzanol and its p-coumarate and caffeate derivatives by normal phase HPLC and a validated method of quantitation. Food Chemistry, 138(4), 2079-2088.

Decker, E. A. (1998). Strategies for manipulating the prooxidative/antioxidative balance of foods to maximize oxidative stability. Trends in Food Science & Technology, 9(6), 241-248.

Dominguez de Maria, P., Sanchez-Montero, J. M., Sinisterra, J. V., & Alcantara, A. R. (2006). Understanding Candida rugosa lipases: an overview. Biotechnology Advances, 24(2), 180-196.

Dordick, J. S. (1989). Enzymatic catalysis in monophasic organic solvents. Enzyme and Microbial Technology, 11(4), 194-211.

Durand, E., Lecomte, J., Barea, B., Dubreucq, E., Lortie, R., & Villeneuve, P. (2013). Evaluation of deep eutectic solvent-water binary mixtures for lipase-catalyzed lipophilization of phenolic acids. Green Chemistry, 15(8), 2275-2282.

References

41

El-Seedi, H. R., El-Said, A. M. A., Khalifa, S. A. M., Göransson, U., Bohlin, L., Borg-Karlson, A.-K., & Verpoorte, R. (2012). Biosynthesis, Natural Sources, Dietary Intake, Pharmacokinetic Properties, and Biological Activities of Hydroxycinnamic Acids. Journal of Agricultural and Food Chemistry, 60(44), 10877-10895.

Elenkov, I. J., Todorova, D. I., Bankova, V. S., & Milkova, T. S. (1995). Synthesis of Steryl Esters of Phenolic-Acids by a Heterogeneous Wittig Reaction. Journal of Natural Products, 58(2), 280-283.

Esche, R., Barnsteiner, A., Scholz, B., & Engel, K. H. (2012). Simultaneous Analysis of Free Phytosterols/Phytostanols and Intact Phytosteryl/Phytostanyl Fatty Acid and Phenolic Acid Esters in Cereals. Journal of Agricultural and Food Chemistry, 60(21), 5330-5339.

Esteban-Torres, M., Landete, J. M., Reveron, I., Santamaria, L., de las Rivas, B., & Munoz, R. (2015). A Lactobacillus plantarum esterase active on a broad range of phenolic esters. Applied and Environment Microbiology, 81(9), 3235-3242.

Esteban-Torres, M., Reveron, I., Mancheno, J. M., de Las Rivas, B., & Munoz, R. (2013). Characterization of a feruloyl esterase from Lactobacillus plantarum. Applied and Environment Microbiology, 79(17), 5130-5136.

Facchini, P. J., Hagel, J., & Zulak, K. G. (2002). Hydroxycinnamic acid amide metabolism: physiology and biochemistry. Canadian Journal of Botany, 80(6), 577-589.

Fang, J., Reichelt, M., Kai, M., & Schneider, B. (2012). Metabolic profiling of lignans and other secondary metabolites from rapeseed ( Brassica napus L.). Journal of Agricultural and Food Chemistry, 60(42), 10523-10529.

Fang, N. B., Yu, S. G., & Badger, T. M. (2003). Characterization of triterpene alcohol and sterol ferulates in rice bran using LC-MS/MS. Journal of Agricultural and Food Chemistry, 51(11), 3260-3267.

Faulds, C. B. (2010). What can feruloyl esterases do for us? Phytochemistry Reviews, 9(1), 121-132.

Faulds, C. B., Molina, R., Gonzalez, R., Husband, F., Juge, N., Sanz-Aparicio, J., & Hermoso, J. A. (2005). Probing the determinants of substrate specificity of a feruloyl esterase, AnFaeA, from Aspergillus niger. FEBS Journal, 272(17), 4362-4371.

Faulds, C. B., & Williamson, G. (1994). Purification and characterization of a ferulic acid esterase (FAE-III) from Aspergillus niger: specificity for the phenolic moiety and binding to microcrystalline cellulose. Microbiology, 140, 779-787.

Figueroa-Espinoza, M. C., Laguerre, M., Villeneuve, P., & Lecomte, J. (2013). From phenolics to phenolipids: Optimizing antioxidants in lipid dispersions. Lipid Technology, 25(6), 131-134.

Figueroa-Espinoza, M. C., & Villeneuve, P. (2005). Phenolic acids enzymatic lipophilization. Journal of Agricultural and Food Chemistry, 53(8), 2779-2787.

Fu, Y. Q., Zhang, Y., Hu, H. Y., Chen, Y., Wang, R., Li, D., & Liu, S. B. (2014). Design and straightforward synthesis of novel galloyl phytosterols with excellent antioxidant activity. Food Chemistry, 163, 171-177.

Giuliani, S., Piana, C., Setti, L., Hochkoeppler, A., Pifferi, P. G., Williamson, G., & Faulds, C. B. (2001). Synthesis of pentylferulate by a feruloyl esterase from Aspergillus niger using water-in-oil microemulsions. Biotechnology Letters, 23(4), 325-330.

Gopalan, N., Rodriguez-Duran, L. V., Saucedo-Castaneda, G., & Nampoothiri, K. M. (2015). Review on technological and scientific aspects of feruloyl esterases: A versatile enzyme for biorefining of biomass. Bioresource Technology, 193, 534-544.

References

42

Graça, J. (2010). Hydroxycinnamates in suberin formation. Phytochemistry Reviews, 9(1), 85-91.

Graça, J., & Pereira, H. (2000). Suberin structure in potato periderm: Glycerol, long-chain monomers, and glyceryl and feruloyl dimers. Journal of Agricultural and Food Chemistry, 48(11), 5476-5483.

Gu, S. S., Wang, J., Wei, X. B., Cui, H. S., Wu, X. Y., & Wu, F. (2014). Enhancement of Lipase-catalyzed Synthesis of Caffeic Acid Phenethyl Ester in Ionic Liquid with DMSO Co-solvent. Chinese Journal of Chemical Engineering, 22(11-12), 1314-1321.

Guyot, B., Bosquette, B., Pina, M., & Graille, J. (1997). Esterification of phenolic acids from green coffee with an immobilized lipase from Candida antarctica in solvent-free medium. Biotechnology Letters, 19(6), 529-532.

Ha, S. H., Anh, T. V., & Koo, Y.-M. (2013). Optimization of lipase-catalyzed synthesis of caffeic acid phenethyl ester in ionic liquids by response surface methodology. Bioprocess and Biosystems Engineering, 36(6), 799-807.

Hashizume, K., Ito, T., Ishizuka, T., & Takeda, N. (2013a). Formation of ethyl ferulate by rice koji enzyme in sake and mirin mash conditions. Journal of Bioscience and Bioengineering, 116(2), 209-213.

Hashizume, K., Ito, T., Shimohashi, M., Ishizuka, T., & Okuda, M. (2013b). Ferulic Acid and Ethyl Ferulate in Sake: Comparison of Levels between Sake and Mirin and Analysis of Their Sensory Properties. Food Science and Technology Research, 19(4), 705-709.

He, D., Simoneit, B. R. T., Jara, B., & Jaffé, R. (2015). Gas chromatography mass spectrometry based profiling of alkyl coumarates and ferulates in two species of cattail (Typha domingensis P., and Typha latifolia L.). Phytochemistry Letters, 13, 91-98.

He, W., Jia, C., Ma, Y., Yang, Y., Zhang, X., Feng, B., & Yue, L. (2010). Lipase-catalyzed synthesis of phytostanyl esters in non-aqueous media. Journal of Molecular Catalysis B: Enzymatic, 67(1-2), 60-65.

Hellner, G., Toke, E. R., Nagy, V., Szakacs, G., & Poppe, L. (2010). Integrated enzymatic production of specific structured lipid and phytosterol ester compositions. Process Biochemistry, 45(8), 1245-1250.

Hermoso, J. A., Sanz-Aparicio, J., Molina, R., Juge, N., Gonzalez, R., & Faulds, C. B. (2004). The crystal structure of feruloyl esterase A from Aspergillus niger suggests evolutive functional convergence in feruloyl esterase family. Journal of Molecular Biology, 338(3), 495-506.

Hixson, J. L., Sleep, N. R., Capone, D. L., Elsey, G. M., Curtin, C. D., Sefton, M. A., & Taylor, D. K. (2012). Hydroxycinnamic Acid Ethyl Esters as Precursors to Ethylphenols in Wine. Journal of Agricultural and Food Chemistry, 60(9), 2293-2298.

Hu, L., Llibin, S., Li, J., Qi, L., Zhang, X., Yu, D., Walid, E., & Jiang, L. (2015). Lipase-catalyzed transesterification of soybean oil and phytosterol in supercritical CO2. Bioprocess and Biosystems Engineering, 38(12), 2343-2347.

Humberstone, F. J., & Briggs, D. E. (2002). Partial purification of ferulic acid esterase from malted barley. Journal of the Institute of Brewing, 108(4), 439-443.

Jayaprakasam, B., Vanisree, M., Zhang, Y. J., Dewitt, D. L., & Nair, M. G. (2006). Impact of alkyl esters of caffeic and ferulic acids on tumor cell proliferation, cyclooxygenase enzyme, and lipid peroxidation. Journal of Agricultural and Food Chemistry, 54(15), 5375-5381.

Jiang, Z. H., Yu, M., Ren, L. W., Zhou, H., & Wei, P. (2013). Synthesis of phytosterol esters catalyzed by immobilized lipase in organic media. Chinese Journal of Catalysis, 34(12), 2255-2262.

References

43

Karboune, S., St-Louis, R., & Kermasha, S. (2008). Enzymatic synthesis of structured phenolic lipids by acidolysis of flaxseed oil with selected phenolic acids. Journal of Molecular Catalysis B: Enzymatic, 52-53, 96-105.

Katsoura, M. H., Polydera, A. C., Tsironis, L. D., Petraki, M. P., Rajacic, S. K., Tselepis, A. D., & Stamatis, H. (2009). Efficient enzymatic preparation of hydroxycinnamates in ionic liquids enhances their antioxidant effect on lipoproteins oxidative modification. New biotechnology, 26(1-2), 83-91.

Kern, S. M., Bennett, R. N., Needs, P. W., Mellon, F. A., Kroon, P. A., & Garcia-Conesa, M. T. (2003). Characterization of metabolites of hydroxycinnamates in the in vitro model of human small intestinal epithelium caco-2 cells. Journal of Agricultural and Food Chemistry, 51(27), 7884-7891.

Kim, B. H., & Akoh, C. C. (2007). Modeling and optimization of lipase-catalyzed synthesis of phytosteryl esters of oleic acid by response surface methodology. Food Chemistry, 102(1), 336-342.

Kim, I. A., Kim, B. G., Kim, M., & Ahn, J. H. (2012). Characterization of hydroxycinnamoyltransferase from rice and its application for biological synthesis of hydroxycinnamoyl glycerols. Phytochemistry, 76, 25-31.

King, J. W., Snyder, J. M., Frykman, H., & Neese, A. (2001). Sterol ester production using lipase-catalyzed reactions in supercritical carbon dioxide. European Food Research and Technology, 212(5), 566-569.

Kondo, H., Sugiyama, H., Katayama, S., & Nakamura, S. (2014). Enhanced antiamyloidal activity of hydroxy cinnamic acids by enzymatic esterification with alkyl alcohols. Biotechnology and Applied Biochemistry, 61(4), 401-407.

Kondo, Y., Nakajima, K., Nozoe, S., & Suzuki, S. (1988). Isolation of Ovulatory-Active Substances from Crops of Job's Tears (Coix lacryma-jobi L. var. ma-yuen STAPF.). Chemical & Pharmaceutical Bulletin, 36(8), 3147-3152.

Koseki, T., Takahashi, K., Fushinobu, S., Iefuji, H., Iwano, K., Hashizume, K., & Matsuzawa, H. (2005). Mutational analysis of a feruloyl esterase from Aspergillus awamori involved in substrate discrimination and pH dependence. Biochimica et Biophysica Acta, 1722(2), 200-208.

Kosma, D. K., Molina, I., Ohlrogge, J. B., & Pollard, M. (2012). Identification of an Arabidopsis fatty alcohol:caffeoyl-Coenzyme A acyltransferase required for the synthesis of alkyl hydroxycinnamates in root waxes. Plant Physiology, 160(1), 237-248.

Kosma, D. K., Rice, A., & Pollard, M. (2015). Analysis of aliphatic waxes associated with root periderm or exodermis from eleven plant species. Phytochemistry, 117, 351-362.

Kroon, P. A., Faulds, C. B., Ryden, P., Robertson, J. A., & Williamson, G. (1997). Release of covalently bound ferulic acid from fiber in the human colon. Journal of Agricultural and Food Chemistry, 45(3), 661-667.

Kroon, P. A., Garcia-Conesa, M. T., Fillingham, I. J., Hazlewood, G. P., & Williamson, G. (1999). Release of ferulic acid dehydrodimers from plant cell walls by feruloyl esterases. Journal of the Science of Food and Agriculture, 79(3), 428-434.

Kumar, A., & Kanwar, S. S. (2011). Synthesis of ethyl ferulate in organic medium using celite-immobilized lipase. Bioresource Technology, 102(3), 2162-2167.

References

44

Kumar, M. S. S., Dahuja, A., Rai, R. D., Walia, S., & Tyagi, A. (2014). Role of -oryzanol in drought-tolerant and susceptible cultivars of rice (Oryza sativa L.). Indian Journal of Biochemistry & Biophysics, 51(1), 75-80.

Kumar, V., Jahan, F., Raghuwanshi, S., Mahajan, R. V., & Saxena, R. K. (2013). Immobilization of Rhizopus oryzae lipase on magnetic Fe3O4-chitosan beads and its potential in phenolic acids ester synthesis. Biotechnology and Bioprocess Engineering, 18(4), 787-795.

Kurata, A., Kitamura, Y., Irie, S., Takemoto, S., Akai, Y., Hirota, Y., Fujita, T., Iwai, K., Furusawa, M., & Kishimoto, N. (2010). Enzymatic synthesis of caffeic acid phenethyl ester analogues in ionic liquid. Journal of Biotechnology, 148(2-3), 133-138.

Laguerre, M., Bayrasy, C., Lecomte, J., Chabi, B., Decker, E. A., Wrutniak-Cabello, C., Cabello, G., & Villeneuve, P. (2013). How to boost antioxidants by lipophilization? Biochimie, 95(1), 20-26.

Laguerre, M., Giraldo, L. J. L., Lecomte, J., Figueroa-Espinoza, M. C., Barea, B., Weiss, J., Decker, E. A., & Villeneuve, P. (2009). Chain Length Affects Antioxidant Properties of Chlorogenate Esters in Emulsion: The Cutoff Theory Behind the Polar Paradox. Journal of Agricultural and Food Chemistry, 57(23), 11335-11342.

Laszlo, J. A., & Compton, D. L. (2006). Enzymatic glycerolysis and transesterification of vegetable oil for enhanced production of feruloylated glycerols. Journal of the American Oil Chemists Society, 83(9), 765-770.

Laszlo, J. A., Compton, D. L., Eller, F. J., Taylor, S. L., & Isbell, T. A. (2003). Packed-bed bioreactor synthesis of feruloylated monoacyl- and diacylglycerols: clean production of a “green” sunscreen. Green Chemistry, 5(4), 382.

Lee, G. S., Widjaja, A., & Ju, Y. H. (2006). Enzymatic synthesis of cinnamic acid derivatives. Biotechnology Letters, 28(8), 581-585.

Li, R., Jia, C. S., Yue, L., Zhang, X. M., Xia, Q. Y., Zhao, S. L., Feng, B., Zhong, F., & Chen, W. J.

(2010). Lipase-Catalyzed Synthesis of Conjugated Linoleyl -Sitosterol and Its Cholesterol-Lowering Properties in Mice. Journal of Agricultural and Food Chemistry, 58(3), 1898-1902.

Li, W., Li, N., Tang, Y., Li, B., Liu, L., Zhang, X., Fu, H., & Duan, J. A. (2012). Biological activity evaluation and structure-activity relationships analysis of ferulic acid and caffeic acid derivatives for anticancer. Bioorganic & Medicinal Chemistry Letters, 22(19), 6085-6088.

Lofty, S., Negrel, J., & Javelle, F. (1994). Formation of Omega-Feruloyloxypalmitic Acid by an Enzyme from Wound-Healing Potato-Tuber Disks. Phytochemistry, 35(6), 1419-1424.

Lubinus, T., Barnsteiner, A., Skurk, T., Hauner, H., & Engel, K. H. (2013). Fate of dietary phytosteryl/-stanyl esters: analysis of individual intact esters in human feces. European Journal of Nutrition, 52(3), 997-1013.

Madhavi Latha, G., Srinivas, P., & Muralikrishnah, G. (2007). Purification and characterization of ferulic acid esterase from malted finger millet (Eleusine coracana, indaf-15). Journal of Agricultural and Food Chemistry, 55(23), 9704-9712.

Manach, C., Scalbert, A., Morand, C., Remesy, C., & Jimenez, L. (2004). Polyphenols: food sources and bioavailability. American Journal of Clinical Nutrition, 79(5), 727-747.

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.

References

45

Martinez, I., Markovits, A., Chamy, R., & Markovits, A. (2004). Lipase-catalyzed solvent-free transesterification of wood sterols. Applied Biochemistry and Biotechnology, 112(1), 55-62.

Miller, A., & Engel, K. H. (2006). Content of -oryzanol and composition of steryl ferulates in brown rice (Oryza sativa L.) of European origin. Journal of Agricultural and Food Chemistry, 54(21), 8127-8133.

Molina, I., Li-Beisson, Y., Beisson, F., Ohlrogge, J. B., & Pollard, M. (2009). Identification of an Arabidopsis feruloyl-coenzyme A transferase required for suberin synthesis. Plant Physiology, 151(3), 1317-1328.

Morinaga, N., Maeda, A., Mizuno, T., Bunya, M., Sugihara, S., & Sugihara, A. (2011). Synthesis of fatty acid sterol esters using cholesterol esterase from Trichoderma sp AS59. Enzyme and Microbial Technology, 48(6-7), 498-504.

Moukouli, M., Topakas, E., & Christakopoulos, P. (2008). Cloning, characterization and functional expression of an alkalitolerant type C feruloyl esterase from Fusarium oxysporum. Applied Microbiology and Biotechnology, 79(2), 245-254.

Negishi, S., Hidaka, I., Takahashi, I., & Kunita, S. (2003). Transesterification of phytosterol and edible oil by lipase powder at high temperature. Journal of the American Oil Chemists Society, 80(9), 905-907.

Nićiforović, N., & Abramovič, H. (2014). Sinapic Acid and Its Derivatives: Natural Sources and Bioactivity. Comprehensive Reviews in Food Science and Food Safety, 13(1), 34-51.

No, D. S., Zhao, T., Lee, J., Lee, J. S., & Kim, I. H. (2013). Synthesis of Phytosteryl Ester Containing Pinolenic Acid in a Solvent-Free System Using Immobilized Candida rugosa Lipase. Journal of Agricultural and Food Chemistry, 61(37), 8934-8940.

Norton, R. A. (1995). Quantitation of Steryl Ferulate and p-Coumarate Esters from Corn and Rice. Lipids, 30(3), 269-274.

Pan, X. X., Chen, B. Q., Wang, J., Zhang, X. Z., Zhul, B. Y., & Tan, T. W. (2012). Enzymatic Synthesizing of Phytosterol Oleic Esters. Applied Biochemistry and Biotechnology, 168(1), 68-77.

Pang, N., Gu, S. S., Jun, W. A., Hong-Sheng, C. A., Fang-Qin, W. A., Xi, L. A., Xing-Yu, Z. A., & Wu, F. A. (2013). A novel chemoenzymatic synthesis of propyl caffeate using lipase-catalyzed transesterification in ionic liquid. Bioresource Technology, 139, 337-342.

Panpipat, W., Xu, X., & Guo, Z. (2013). Improved acylation of phytosterols catalyzed by Candida antarctica lipase A with superior catalytic activity. Biochemical Engineering Journal, 70, 55-62.

Piironen, V., Lindsay, D. G., Miettinen, T. A., Toivo, J., & Lampi, A. M. (2000). Plant sterols: biosynthesis, biological function and their importance to human nutrition. Journal of the Science of Food and Agriculture, 80(7), 939-966.

Porter, W. L., Black, E. D., & Drolet, A. M. (1989). Use of Polyamide Oxidative Fluorescence Test on Lipid Emulsions - Contrast in Relative Effectiveness of Antioxidants in Bulk Versus Dispersed Systems. Journal of Agricultural and Food Chemistry, 37(3), 615-624.

Rodrigues, R. C., & Fernandez-Lafuente, R. (2010a). Lipase from Rhizomucor miehei as a biocatalyst in fats and oils modification. Journal of Molecular Catalysis B: Enzymatic, 66(1-2), 15-32.

Rodrigues, R. C., & Fernandez-Lafuente, R. (2010b). Lipase from Rhizomucor miehei as an industrial biocatalyst in chemical process. Journal of Molecular Catalysis B: Enzymatic, 64(1-2), 1-22.

References

46

Sabally, K., Karboune, S., Yeboah, F. K., & Kermasha, S. (2005). Lipase-catalyzed esterification of selected phenolic acids with linolenyl alcohols in organic solvent media. Applied Biochemistry and Biotechnology, 127(1), 17-27.

Safari, M., Karboune, S., St-Louis, R., & Kermasha, S. (2006). Enzymatic synthesis of structured phenolic lipids by incorporation of selected phenolic acids into triolein. Biocatalysis and Biotransformation, 24(4), 272-279.

Sancho, A. I., Faulds, C. B., Bartolome, B., & Williamson, G. (1999). Characterisation of feruloyl esterase activity in barley. Journal of the Science of Food and Agriculture, 79(3), 447-449.

Sandoval, G., Quintana, P. G., Baldessari, A., Ballesteros, A. O., & Plou, F. J. (2015). Lipase-catalyzed preparation of mono- and diesters of ferulic acid. Biocatalysis and Biotransformation, 33(2), 89-97.

Schubot, F. D., Kataeva, I. A., Blum, D. L., Shah, A. K., Ljungdahl, L. G., Rose, J. P., & Wang, B. C. (2001). Structural basis for the substrate specificity of the feruloyl esterase domain of the cellulosomal xylanase Z from Clostridium thermocellum. Biochemistry, 40(42), 12524-12532.

Seino, H. (1987). Enzymatic Esterification of Ferulic Acid with Sterols. Journal of the American Oil Chemists Society, 64(9), 1262-1262.

Seitz, L. M. (1989). Stanol and sterol esters of ferulic and p-coumaric acids in wheat, corn, rye, and triticale. J Agric Food Chem, 37(3), 662-667.

Sengupta, A., & Ghosh, M. (2011). The kinetics of enzyme catalyzed synthesis of sterol ester. European Journal of Lipid Science and Technology, 113(6), 763-767.

Serafim, T. L., Carvalho, F. S., Marques, M. P., Calheiros, R., Silva, T., Garrido, J., Milhazes, N., Borges, F., Roleira, F., Silva, E. T., Holy, J., & Oliveira, P. J. (2011). Lipophilic caffeic and ferulic acid derivatives presenting cytotoxicity against human breast cancer cells. Chemical Research in Toxicology, 24(5), 763-774.

Shahidi, F., & Chandrasekara, A. (2009). Hydroxycinnamates and their in vitro and in vivo antioxidant activities. Phytochemistry Reviews, 9(1), 147-170.

Shang, C. Y., Li, W. X., Jiang, F., & Zhang, R. F. (2015). Improved enzymatic properties of Candida rugosa lipase immobilized on ZnO nanowires/macroporous SiO2 microwave absorbing supports. Journal of Molecular Catalysis B: Enzymatic, 113, 9-13.

Sharma, S., Dogra, P., Chauhan, G. S., & Kanwar, S. S. (2014). Synthesis of alkyl coumarate esters by celite-bound lipase of Bacillus lichenzformis SCD11501. Journal of Molecular Catalysis B: Enzymatic, 101, 80-86.

Shimada, Y., Hirota, Y., Baba, T., Sugihara, A., Moriyama, S., Tominaga, Y., & Terai, T. (1999). Enzymatic synthesis of steryl esters of polyunsaturated fatty acids. Journal of the American Oil Chemists Society, 76(6), 713-716.

Sorour, N., Karboune, S., Saint-Louis, R., & Kermasha, S. (2012). Lipase-catalyzed synthesis of structured phenolic lipids in solvent-free system using flaxseed oil and selected phenolic acids as substrates. Journal of Biotechnology, 158(3), 128-136.

Stamatis, H., Sereti, V., & Kolisis, F. N. (1999). Studies on the enzymatic synthesis of lipophilic derivatives of natural antioxidants. Journal of the American Oil Chemists Society, 76(12), 1505-1510.

Stamatis, H., Sereti, V., & Kolisis, F. N. (2001). Enzymatic synthesis of hydrophilic and hydrophobic derivatives of natural phenolic acids in organic media. Journal of Molecular Catalysis B: Enzymatic, 11(4-6), 323-328.

References

47

Stevenson, D. E., Parkar, S. G., Zhang, J., Stanley, R. A., Jensen, D. J., & Cooney, J. M. (2007). Combinatorial enzymic synthesis for functional testing of phenolic acid esters catalysed by Candida antarctica lipase B (Novozym 435

®). Enzyme and Microbial Technology, 40(5), 1078-

1086.

Sun, S., Song, F., Bi, Y., Yang, G., & Liu, W. (2012). Solvent-free enzymatic transesterification of ethyl ferulate and monostearin: optimized by response surface methodology. Journal of Biotechnology, 164(2), 340-345.

Sun, S., & Zhou, W. (2014). Enhanced Enzymatic Preparation of Lipophilic Feruloylated Lipids Using Distearin as Feruloyl Acceptors: Optimization by Response Surface Methodology. Journal of Oleo Science, 63(10), 1011-1018.

Swell, L., Field, H., & Treadwell, C. R. (1954). Sterol Specificity of Pancreatic Cholesterol Esterase. Proceedings of the Society for Experimental Biology and Medicine, 87(1), 216-218.

Tada, M., Matsumoto, R., Yamaguchi, H., & Chiba, K. (1996). Novel antioxidants isolated from Perilla frutescens Britton var. crispa (Thunb.). Bioscience Biotechnology and Biochemistry, 60(7), 1093-1095.

Takagi, T., & Iida, T. (1980). Antioxidant for Fats and Oils from Canary Seed - Sterol and Triterpene Alcohol Esters of Caffeic Acid. Journal of the American Oil Chemists Society, 57(10), 326-330.

Takeda, Y., Aono, R., & Doukyu, N. (2006). Purification, characterization, and molecular cloning of organic-solvent-tolerant cholesterol esterase from cyclohexane-tolerant Burkholderia cepacia strain ST-200. Extremophiles, 10(4), 269-277.

Tan, Z., & Shahidi, F. (2011). Chemoenzymatic Synthesis of Phytosteryl Ferulates and Evaluation of Their Antioxidant Activity. Journal of Agricultural and Food Chemistry, 59(23), 12375-12383.

Tan, Z., & Shahidi, F. (2012a). A novel chemoenzymatic synthesis of phytosteryl caffeates and assessment of their antioxidant activity. Food Chemistry, 133(4), 1427-1434.

Tan, Z., & Shahidi, F. (2013). Phytosteryl sinapates and vanillates: chemoenzymatic synthesis and antioxidant capacity assessment. Food Chemistry, 138(2-3), 1438-1447.

Tan, Z. L., Le, K., Moghadasian, M., & Shahidi, F. (2012). Enzymatic synthesis of phytosteryl docosahexaneates and evaluation of their anti-atherogenic effects in apo-E deficient mice. Food Chemistry, 134(4), 2097-2104.

Tan, Z. L., & Shahidi, F. (2012b). Optimization of Enzymatic Synthesis of Phytosteryl Caprylates Using Response Surface Methodology. Journal of the American Oil Chemists Society, 89(4), 657-666.

Teixeira, A. R. S., Santos, J. L. C., & Crespo, J. G. (2011). Production of Steryl Esters from Vegetable Oil Deodorizer Distillates by Enzymatic Esterification. Industrial & Engineering Chemistry Research, 50(5), 2865-2875.

Teixeira, A. R. S., Santos, J. L. C., & Crespo, J. G. (2012). Lipase-Catalyzed Consecutive Batch Reaction for Production of Steryl Esters from Vegetable Oil Deodorizer Distillates. Industrial & Engineering Chemistry Research, 51(15), 5443-5455.

Teixeira, A. R. S., Santos, J. L. C., & Crespo, J. G. (2014). Production of Steryl Esters Using Alternative Sources of Sterols and Free Fatty Acids: Modeling and Guidelines. Industrial & Engineering Chemistry Research, 53(26), 10534-10539.

Theng, Y., Wu, X.-M., Branford-White, C., Quan, J., & Zhu, L.-M. (2009). Dual response surface-optimized process for feruloylated diacylglycerols by selective lipase-catalyzed transesterification in solvent free system. Bioresource Technology, 100(12), 2896-2901.

References

48

Thorn, C., Gustafsson, H., & Olsson, L. (2011). Immobilization of feruloyl esterases in mesoporous materials leads to improved transesterification yield. Journal of Molecular Catalysis B: Enzymatic, 72(1-2), 57-64.

Topakas, E., Moukouli, M., Dimarogona, M., & Christakopoulos, P. (2012). Expression, characterization and structural modelling of a feruloyl esterase from the thermophilic fungus Myceliophthora thermophila. Applied Microbiology and Biotechnology, 94(2), 399-411.

Topakas, E., Stamatis, H., Biely, P., Kekos, D., Macris, B. J., & Christakopoulos, P. (2003). Purification and characterization of a feruloyl esterase from Fusarium oxysporum catalyzing esterification of phenolic acids in ternary water-organic solvent mixtures. Journal of Biotechnology, 102(1), 33-44.

Topakas, E., Vafiadi, C., & Christakopoulos, P. (2007). Microbial production, characterization and applications of feruloyl esterases. Process Biochemistry, 42(4), 497-509.

Topakas, E., Vafiadi, C., Stamatis, H., & Christakopoulos, P. (2005). Sporotrichum thermophile type C feruloyl esterase (StFaeC): purification, characterization, and its use for phenolic acid (sugar) ester synthesis. Enzyme and Microbial Technology, 36(5-6), 729-736.

Torrelo, G., Torres, C. F., & Reglero, G. (2012). Enzymatic strategies for solvent-free production of short and medium chain phytosteryl esters. European Journal of Lipid Science and Technology, 114(6), 670-676.

Torrelo, G., Torres, C. F., Senorans, F. J., Blanco, R. M., & Reglero, G. (2009). Solvent-free preparation of phytosteryl esters with fatty acids from butterfat in equimolecular conditions in the presence of a lipase from Candida rugosa. Journal of Chemical Technology and Biotechnology, 84(5), 745-750.

Torres, C. F., Torrelo, G., Señorans, F. J., & Reglero, G. (2007). A two steps enzymatic procedure to obtain sterol esters, tocopherols and fatty acid ethyl esters from soybean oil deodorizer distillate. Process Biochemistry, 42(9), 1335-1341.

Tsuchiyama, M., Sakamoto, T., Fujita, T., Murata, S., & Kawasaki, H. (2006). Esterification of ferulic acid with polyols using a ferulic acid esterase from Aspergillus niger. Biochimica et Biophysica Acta, 1760(7), 1071-1079.

Udatha, D. B., Kouskoumvekaki, I., Olsson, L., & Panagiotou, G. (2011). The interplay of descriptor-based computational analysis with pharmacophore modeling builds the basis for a novel classification scheme for feruloyl esterases. Biotechnology Advances, 29(1), 94-110.

Uwai, K., Osanai, Y., Imaizumi, T., Kanno, S., Takeshita, M., & Ishikawa, M. (2008). Inhibitory effect of the alkyl side chain of caffeic acid analogues on lipopolysaccharide-induced nitric oxide production in RAW264.7 macrophages. Bioorganic and Medicinal Chemistry, 16(16), 7795-7803.

Vafiadi, C., Topakas, E., Alissandratos, A., Faulds, C. B., & Christakopoulos, P. (2008a). Enzymatic synthesis of butyl hydroxycinnamates and their inhibitory effects on LDL-oxidation. Journal of Biotechnology, 133(4), 497-504.

Vafiadi, C., Topakas, E., & Christakopoulos, P. (2008b). Preparation of multipurpose cross-linked enzyme aggregates and their application to production of alkyl ferulates. Journal of Molecular Catalysis B: Enzymatic, 54(1-2), 35-41.

Vafiadi, C., Topakas, E., Christakopoulos, P., & Faulds, C. B. (2006). The feruloyl esterase system of Talaromyces stipitatus: Determining the hydrolytic and synthetic specificity of TsFaeC. Journal of Biotechnology, 125(2), 210-221.

References

49

Vafiadi, C., Topakas, E., Wong, K. K. Y., Suckling, I. D., & Christakopoulos, P. (2005). Mapping the hydrolytic and synthetic selectivity of a type C feruloyl esterase (StFaeC) from Sporotrichum thermophile using alkyl ferulates. Tetrahedron: Asymmetry, 16(2), 373-379.

Vardakou, M., Nueno Palop, C., Gasson, M., Narbad, A., & Christakopoulos, P. (2007). In vitro three-stage continuous fermentation of wheat arabinoxylan fractions and induction of hydrolase activity by the gut microflora. International Journal of Biological Macromolecules, 41(5), 584-589.

Villeneuve, P. (2007). Lipases in lipophilization reactions. Biotechnology Advances, 25(6), 515-536.

Villeneuve, P., Muderhwa, J. M., Graille, J., & Haas, M. J. (2000). Customizing lipases for biocatalysis: a survey of chemical, physical and molecular biological approaches. Journal of Molecular Catalysis B: Enzymatic, 9(4-6), 113-148.

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.

Vu, P. L., Shin, J. A., Lim, C. H., & Lee, K. T. (2004). Lipase-catalyzed production of phytosteryl esters and their crystallization behavior in corn oil. Food Research International, 37(2), 175-180.

Wang, J., Gu, S. S., Cui, H. S., Wu, X. Y., & Wu, F. A. (2014). A novel continuous flow biosynthesis of caffeic acid phenethyl ester from alkyl caffeate and phenethanol in a packed bed microreactor. Bioresource Technology, 158, 39-47.

Wang, J., Gu, S. S., Cui, H. S., Yang, L. Q., & Wu, X. Y. (2013). Rapid synthesis of propyl caffeate in ionic liquid using a packed bed enzyme microreactor under continuous-flow conditions. Bioresource Technology, 149, 367-374.

Wang, J., & Shahidi, F. (2014a). Acidolysis of p-coumaric acid with omega-3 oils and antioxidant activity of phenolipid products in in vitro and biological model systems. Journal of Agricultural and Food Chemistry, 62(2), 454-461.

Wang, J., & Shahidi, F. (2014b). Antioxidant activity of monooleyl and dioleyl p-coumarates in in vitro and biological model systems. European Journal of Lipid Science and Technology, 116(4), 370-379.

Wang, J., Wang, S. S., Li, Z. J., Gu, S. S., Wu, X. Y., & Wu, F. (2015a). Ultrasound irradiation accelerates the lipase-catalyzed synthesis of methyl caffeate in an ionic liquid. Journal of Molecular Catalysis B: Enzymatic, 111, 21-28.

Wang, X. K., Geng, X., Egashira, Y., & Sanada, H. (2004). Purification and characterization of a feruloyl esterase from the intestinal bacterium Lactobacillus acidophilus. Applied and Environmental Microbiology, 70(4), 2367-2372.

Wang, Z., Hwang, S. H., & Lim, S. S. (2015b). 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, H. K., Stecher, H., & Faber, K. (1995). Sensitivity of Microbial Lipases to Acetaldehyde Formed by Acyl-Transfer Reactions from Vinyl Esters. Biotechnology Letters, 17(8), 803-808.

References

50

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.

Weber, N., Weitkamp, P., & Mukherjee, K. D. (2002). Cholesterol-lowering food additives: lipase-catalysed preparation of phytosterol and phytostanol esters. Food Research International, 35(2-3), 177-181.

Weber, N., Weitkamp, P., & Mukherjee, K. D. (2003). Thermomyces lanuginosus lipase-catalyzed transesterification of sterols by methyl oleate. European Journal of Lipid Science and Technology, 105(10), 624-626.

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.

Widjaja, A., Yeh, T. H., & Ju, Y. H. (2008). Enzymatic synthesis of caffeic acid phenethyl ester. Journal of the Chinese Institute of Chemical Engineers, 39(5), 413-418.

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.

Xiao, P. L., Zhang, A. J., Zheng, L. Y., & Song, Y. Q. (2014). Straightforward Enzyme-Catalyzed Asymmetric Synthesis of Caffeic Acid Esters in Enantioenriched Form. Industrial & Engineering Chemistry Research, 53(29), 11638-11645.

Yang, H., Mu, Y., Chen, H., Xiu, Z., & Yang, T. (2013). Enzymatic synthesis of feruloylated lysophospholipid in a selected organic solvent medium. Food Chemistry, 141(4), 3317-3322.

Yang, Z., Glasius, M., & Xu, X. B. (2012a). Enzymatic Transesterification of Ethyl Ferulate with Fish Oil and Reaction Optimization by Response Surface Methodology. Food Technology and Biotechnology, 50(1), 88-97.

Yang, Z., Guo, Z., & Xu, X. (2012b). Enzymatic lipophilisation of phenolic acids through esterification with fatty alcohols in organic solvents. Food Chemistry, 132(3), 1311-1315.

Yang, Z., Guo, Z., & Xu, X. B. (2012c). Ionic Liquid-Assisted Solubilization for Improved Enzymatic Esterification of Phenolic Acids. Journal of the American Oil Chemists Society, 89(6), 1049-1055.

Yoshida-Shimokawa, T., Yoshida, S., Kakegawa, K., & Ishii, T. (2001). Enzymic feruloylation of arabinoxylan-trisaccharide by feruloyl-CoA:arabinoxylan-trisaccharide O-hydroxycinnamoyl transferase from Oryza sativa. Planta, 212(3), 470-474.

Yoshida, Y., Kimura, Y., Kadota, M., Tsuno, T., & Adachi, S. (2006). Continuous synthesis of alkyl ferulate by immobilized Candida antarctica lipase at high temperature. Biotechnology Letters, 28(18), 1471-1474.

References

51

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.

Zadernowski, R., Naczk, M., & Nesterowicz, J. (2005). Phenolic acid profiles in some small berries. Journal of Agricultural and Food Chemistry, 53(6), 2118-2124.

Zeng, C. X., Qi, S. J., Li, Z. G., Luo, R. M., Yang, B., & Wang, Y. H. (2015). Enzymatic synthesis of phytosterol esters catalyzed by Candida rugosa lipase in water-in-[Bmim]PF6 microemulsion. Bioprocess and Biosystems Engineering, 38(5), 939-946.

Zeng, Y., Yin, X., Wu, M.-C., Yu, T., Feng, F., Zhu, T.-D., & Pang, Q.-F. (2014). Expression of a novel feruloyl esterase from Aspergillus oryzae in Pichia pastoris with esterification activity. Journal of Molecular Catalysis B: Enzymatic, 110, 140-146.

Zerner, B., & Bender, M. L. (1964). Kinetic Consequences of Acyl-Enzyme Mechanism for Reactions of Specific Substrates with Chymotrypsin. Journal of the American Chemical Society, 86(18), 3669-&.

Zerner, B., Bond, R. P. M., & Bender, M. L. (1964). 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.

Zhao, Z. H., & Moghadasian, M. H. (2010). Bioavailability of hydroxycinnamates: a brief review of in vivo and in vitro studies. Phytochemistry Reviews, 9(1), 133-145.

Zheng, M. M., Dong, L., Lu, Y., Guo, P. M., Deng, Q. C., Li, W. L., Feng, Y. Q., & Huang, F. H. (2012a). Immobilization of Candida rugosa lipase on magnetic poly(allyl glycidyl ether-co-ethylene glycol dimethacrylate) polymer microsphere for synthesis of phytosterol esters of unsaturated fatty acids. Journal of Molecular Catalysis B: Enzymatic, 74(1-2), 16-23.

Zheng, M. M., Huang, Q., Huang, F. H., Guo, P. M., Xiang, X., Deng, Q. C., Li, W. L., Wan, C. Y., & Zheng, C. (2014). Production of Novel "Functional Oil" Rich in Diglycerides and Phytosterol Esters with "One-Pot" Enzymatic Transesterification. Journal of Agricultural and Food Chemistry, 62(22), 5142-5148.

Zheng, M. M., Lu, Y., Dong, L., Guo, P. M., Deng, Q. C., Li, W. L., Feng, Y. Q., & Huang, F. H. (2012b). Immobilization of Candida rugosa lipase on hydrophobic/strong cation-exchange functional silica particles for biocatalytic synthesis of phytosterol esters. Bioresource Technology, 115, 141-146.

Zheng, M. M., Lu, Y., Huang, F. H., Wang, L., Guo, P. M., Feng, Y. Q., & Deng, Q. C. (2013). Lipase Immobilization on Hyper-Cross-Linked Polymer-Coated Silica for Biocatalytic Synthesis of Phytosterol Esters with Controllable Fatty Acid Composition. Journal of Agricultural and Food Chemistry, 61(1), 231-237.

Zheng, M. M., Wang, L., Huang, F. H., Dong, L., Guo, P. M., Deng, Q. C., Li, W. L., & Zheng, C. (2012c). Ultrasonic pretreatment for lipase-catalyed synthesis of phytosterol esters with different acyl donors. Ultrasonics Sonochemistry, 19(5), 1015-1020.

Zheng, M. M., Zhu, J. X., Huang, F. H., Xiang, X., Shi, J., Deng, Q. 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.

References

52

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.

Part B - Research Papers

53

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).

Part B: Enzymatic esterification of ferulic acid

55

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


56

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


57

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.


58

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.

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

Figure 1: General enzymatically catalyzed reaction examined in this study.


59

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


60

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:

𝑌 = 𝛽𝑘0 + ∑ 𝛽𝑘𝑖𝑥𝑖 + ∑ 𝛽𝑘𝑖𝑖𝑥𝑖2 + ∑

4

𝑖=1

5

𝑖=1

∑ 𝛽𝑘𝑖𝑗𝑥𝑖𝑥𝑗

5

𝑗=𝑖+1

5

𝑖=1

(1)

where 𝑌 corresponds to the response (molar yield %), 𝛽𝑘0 , 𝛽𝑘𝑖 , 𝛽𝑘𝑖𝑖 , and 𝛽𝑘𝑖𝑗 are constant

coefficients, and the uncoded independent variables are represented by 𝑥𝑖.

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

x5: enzyme:

substrate ratio 0 0 -1 0 0 1 0 0 0 -1 0 0 1 0 -1 0 0 1 -1 -1

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

x5: enzyme:

substrate ratio 1 1 -1 0 0 1 0 -1 0 0 1 0 0 0 -1 0 0 1 0 0


61

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


62

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%.


63

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.


64

Table 3: β-coefficients and corresponding p-values of

the ethyl ferulate and decyl ferulate synthesis

optimization. The variables refer to x1: temperature;

x2: time; x3: ferulic acid; x4: alcohol; x5: enzyme-to-

substrate ratio. p-value = level of significance.

Ethyl ferulate Decyl ferulate

β-coefficient p-value β -coefficient p-value

β0 69.06

88.46

x1 1.89 0.0788 4.09 0.0040*

x2 4.22 0.0004* 12.25 0.0000*

x3 -10.14 0.0000* -0.76 0.5611

x4 14.66 0.0000* 5.03 0.0006*

x5 9.76 0.0000* 3.00 0.0286*

x1*x2 -1.11 0.5968 -4.39 0.1012

x1*x3 -1.16 0.5796 -1.44 0.5822

x2*x3 1.96 0.3504 -0.11 0.9663

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

x12 -3.51 0.0188* -5.71 0.0031*

x22 -4.94 0.0016* -6.35 0.0013*

x32 -7.78 0.0000* -2.93 0.1056

x42 -16.72 0.0000* -2.17 0.2246

x52 -6.89 0.0000* -11.20 0.0000*

*significant at p = 0.05


65

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).


66

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.


67

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%.


68

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


69

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.


70

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.


71

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.

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.


72

References


Cione, E., Tucci, P., Senatore, V., Perri, M., Trombino, S., Iemma, F., Picci, N., & Genchi, G. (2008). Synthesized esters of ferulic acid induce release of cytochrome c from rat testes mitochondria. Journal of Bioenergetics and Biomembranes, 40(1), 19-26.



Garrido, J., Gaspar, A., Garrido, E. M., Miri, R., Tavakkoli, M., Pourali, S., Saso, L., Borges, F., & Firuzi, O. (2012). Alkyl esters of hydroxycinnamic acids with improved antioxidant activity and lipophilicity protect PC12 cells against oxidative stress. Biochimie, 94(4), 961-967.

Ghamgui, H., Karra-Chaâbouni, M., & Gargouri, Y. (2004). 1-Butyl oleate synthesis by immobilized lipase from Rhizopus oryzae: a comparative study between n-hexane and solvent-free system. Enzyme and Microbial Technology, 35(4), 355-363.

Giuliani, S., Piana, C., Setti, L., Hochkoeppler, A., Pifferi, P. G., Williamson, G., & Faulds, C. B. (2001). Synthesis of pentylferulate by a feruloyl esterase from Aspergillus niger using water-in-oil microemulsions. Biotechnology Letters, 23(4), 325-330.


Hari Krishna, S., & Karanth, N. G. (2002). Lipases and Lipase-Catalyzed Esterification Reactions in Nonaqueous Media. Catalysis Reviews - Science and Engineering, 44(4), 499-591.

Hashizume, K., Ito, T., Shimohashi, M., Ishizuka, T., & Okuda, M. (2013). Ferulic Acid and Ethyl Ferulate in Sake: Comparison of Levels between Sake and Mirin and Analysis of Their Sensory Properties. Food Science and Technology Research, 19(4), 705-709.

He, W. S., Li, J. J., Pan, X. X., Zhou, Y., Jia, C. S., Zhang, X. M., & Feng, B. (2012). Lipase-mediated synthesis of water-soluble plant stanol derivatives in tert-butanol. Bioresource Technology, 114, 1-5.

Hixson, J. L., Sleep, N. R., Capone, D. L., Elsey, G. M., Curtin, C. D., Sefton, M. A., & Taylor, D. K. (2012). Hydroxycinnamic Acid Ethyl Esters as Precursors to Ethylphenols in Wine. Journal of Agricultural and Food Chemistry, 60(9), 2293-2298.

Jayaprakasam, B., Vanisree, M., Zhang, Y. J., Dewitt, D. L., & Nair, M. G. (2006). Impact of alkyl esters of caffeic and ferulic acids on tumor cell proliferation, cyclooxygenase enzyme, and lipid peroxidation. Journal of Agricultural and Food Chemistry, 54(15), 5375-5381.


Kondo, H., Sugiyama, H., Katayama, S., & Nakamura, S. (2013). Enhanced antiamyloidal activity of hydroxy cinnamic acids by enzymatic esterification with alkyl alcohols. Biotechnology and Applied Biochemistry.

Laguerre, M., Bayrasy, C., Lecomte, J., Chabi, B., Decker, E. A., Wrutniak-Cabello, C., Cabello, G., & Villeneuve, P. (2013). How to boost antioxidants by lipophilization? Biochimie, 95(1), 20-26.

Lee, G. S., Widjaja, A., & Ju, Y. H. (2006). Enzymatic synthesis of cinnamic acid derivatives. Biotechnology Letters, 28(8), 581-585.

Mancuso, C., & Santangelo, R. (2014). Ferulic acid: Pharmacological and toxicological aspects. Food and Chemical Toxicology, 65, 185-195.


73

Sabally, K., Karboune, S., Yeboah, F. K., & Kermasha, S. (2005). Lipase-catalyzed esterification of selected phenolic acids with linolenyl alcohols in organic solvent media. Applied Biochemistry and Biotechnology, 127(1), 17-27.

Šabeder, S., Habulin, M., & Knez, Ž. (2006). Lipase-catalyzed synthesis of fatty acid fructose esters. Journal of Food Engineering, 77(4), 880-886.

Safari, M., Karboune, S., St-Louis, R., & Kermasha, S. (2006). Enzymatic synthesis of structured phenolic lipids by incorporation of selected phenolic acids into triolein. Biocatalysis and Biotransformation, 24(4), 272-279.



Sun, S., Song, F., Bi, Y., Yang, G., & Liu, W. (2012). Solvent-free enzymatic transesterification of ethyl ferulate and monostearin: optimized by response surface methodology. Journal of Biotechnology, 164(2), 340-345.

Vafiadi, C., Topakas, E., Alissandratos, A., Faulds, C. B., & Christakopoulos, P. (2008). Enzymatic synthesis of butyl hydroxycinnamates and their inhibitory effects on LDL-oxidation. Journal of Biotechnology, 133(4), 497-504.

Yang, Z., Glasius, M., & Xu, X. B. (2012a). Enzymatic Transesterification of Ethyl Ferulate with Fish Oil and Reaction Optimization by Response Surface Methodology. Food Technology and Biotechnology, 50(1), 88-97.

Yang, Z., Guo, Z., & Xu, X. (2012b). Enzymatic lipophilisation of phenolic acids through esterification with fatty alcohols in organic solvents. Food Chemistry, 132(3), 1311-1315.

Yoshida, Y., Kimura, Y., Kadota, M., Tsuno, T., & Adachi, S. (2006). Continuous synthesis of alkyl ferulate by immobilized Candida antarctica lipase at high temperature. Biotechnology Letters, 28(18), 1471-1474.


Yunoki, K., Musa, R., Kinoshita, M., Tazaki, H., Oda, Y., & Masao, O. (2004). Presence of higher alcohols as ferulates in potato pulp and its radical-scavenging activity. Bioscience Biotechnology and Biochemistry, 68(12), 2619-2622.

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.

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.

Part B: Enzymatic synthesis of steryl ferulates

75

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


76

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.


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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


78

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.


2.1 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


79

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


80

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:


81

𝑌 = 𝛽𝑘0 + ∑ 𝛽𝑘𝑖𝑥𝑖 + ∑ 𝛽𝑘𝑖𝑖𝑥𝑖2 + ∑ 3

𝑖=14𝑖=1 ∑ 𝛽𝑘𝑖𝑗𝑥𝑖𝑥𝑗

4𝑗=𝑖+1

4𝑖=1 (1)

Y refers to the response (molar yield %), 𝛽𝑘0 , 𝛽𝑘𝑖 , 𝛽𝑘𝑖𝑖 , and 𝛽𝑘𝑖𝑗 are constant coefficient and

𝑥𝑖represents 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.


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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.


83

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


84

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.


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

x12 -3.82 0.00* -3.68 0.00*

x22 -2.80 0.02* -6.05 0.00*

x32 -3.03 0.02* -3.79 0.00*

x42 -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


85

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.


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.


86

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


87

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.,


88

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


89

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


90

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.


91

References



Condo, A. M., Baker, D. C., Moreau, R. A., & Hicks, K. B. (2001). Improved method for the synthesis

of trans-feruloyl--sitostanol. Journal of Agricultural and Food Chemistry, 49(10), 4961-4964.

Dominguez de Maria, P., Sanchez-Montero, J. M., Sinisterra, J. V., & Alcantara, A. R. (2006). Understanding Candida rugosa lipases: an overview. Biotechnology Advances, 24(2), 180-196.

Domínguez de María, P., Sinisterra, J. V., Sánchez-Montero, J. M., Lotti, M., Valero, F., & Alcántara, A. R. (2006). Acyl transfer strategy for the biocatalytical characterisation of Candida rugosa lipases in organic solvents. Enzyme and Microbial Technology, 38(1-2), 199-208.

Evershed, R. P., Spooner, N., Prescott, M. C., & Goad, L. J. (1988). Isolation and Characterization of Intact Steryl Ferulates from Seeds. Journal of Chromatography, 440, 23-35.


Hakala, P., Lampi, A. M., Ollilainen, V., Werner, U., Murkovic, M., Wahala, K., Karkola, S., & Piironen, V. (2002). Steryl phenolic acid esters in cereals and their milling fractions. Journal of Agricultural and Food Chemistry, 50(19), 5300-5307.

He, W., Jia, C., Ma, Y., Yang, Y., Zhang, X., Feng, B., & Yue, L. (2010). Lipase-catalyzed synthesis of phytostanyl esters in non-aqueous media. Journal of Molecular Catalysis B: Enzymatic, 67(1-2), 60-65.

Kondo, Y., Nakajima, K., Nozoe, S., & Suzuki, S. (1988). Isolation of Ovulatory-Active Substances from Crops of Job's Tears (Coix lacryma-jobi L. var. ma-yuen STAPF.). Chemical & Pharmaceutical Bulletin, 36(8), 3147-3152.

Lopez, N., Pernas, M. A., Pastrana, L. M., Sanchez, A., Valero, F., & Rua, M. L. (2004). Reactivity of pure Candida rugosa lipase isoenzymes (Lip1, Lip2, and Lip3) in aqueous and organic media. Influence of the isoenzymatic profile on the lipase performance in organic media. Biotechnology Progress, 20(1), 65-73.

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.


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.


92

Nyström, L., Achrenius, T., Lampi, A. M., Moreau, R. A., & Piironen, V. (2007). A comparison of the antioxidant properties of steryl ferulates with tocopherol at high temperatures. Food Chemistry, 101(3), 947-954.

Nyström, L., Makinen, M., Lampi, A. M., & Piironen, V. (2005). Antioxidant activity of steryl ferulate extracts from rye and wheat bran. Journal of Agricultural and Food Chemistry, 53(7), 2503-2510.

Nyström, L., Moreau, R. A., Lampi, A. M., Hicks, K. B., & Piironen, V. (2008). Enzymatic hydrolysis of steryl ferulates and steryl glycosides. European Food Research and Technology, 227(3), 727-733.



Schär, A., & Nyström, L. (2015). High yielding and direct enzymatic lipophilization of ferulic acid using lipase from Rhizomucor miehei. Journal of Molecular Catalysis B: Enzymatic, 118, 29-35.

Shieh, C. J., Akoh, C. C., & Yee, L. N. (1996). Optimized enzymatic synthesis of geranyl butyrate with lipase AY from Candida rugosa. Biotechnology and Bioengineering, 51(3), 371-374.



Tenkanen, M., Kontkanen, H., Isoniemi, R., Spetz, P., & Holmbom, B. (2002). Hydrolysis of steryl esters by a lipase (Lip 3) from Candida rugosa. Applied Microbiology and Biotechnology, 60(1-2), 120-127.

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.


93

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.



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.


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.


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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%


95

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%


96

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.

A

B

Part B: Esterification of hydroxycinnamic acids and their antioxidant activity

97

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


98

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


99

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


100

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.


101

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


102

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


103

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


104

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.


105


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).

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).

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-

cinnamic

acid [mg]

Enzyme/

substrate

[mg/mg]

Hexane

[µL]

Ethan

ol [µL]

Butanon

e [µL]

Time

[h]

Yield

[%]

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)


106

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).


107

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 derivative

Yield [%]

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.


108

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


109

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.

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

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).


110

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.


111

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.

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.


112

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.

To conclude, the esterification and transesterification of hydroxycinnamic acids by lipases

strongly depends on the structure of the acid substrate and the lipase applieds. The

presence, location and numbers of hydroxyl groups and the unsaturation in the side chain

influence the esterification yield. For example ferulic acid is transesterified by C. rugosa

lipase to a sufficient extent, but the p-coumaric acid without the methoxy group was hardly

accepted as substrate. Depending on the oxidation system the esterification of a

hydroxycinnamic acid with a sterol does not necessarily increase its antioxidant activity.

4. Acknowledgements

This study was conducted with the financial support of the Swiss National Science

Foundation, SNSF (Project 200021_141268) and ETH Zurich.


113

References


Buisman, G. J. H., van Helteren, C. T. W., Kramer, G. F. H., Veldsink, J. W., Derksen, J. T. P., & Cuperus, F. P. (1998). Enzymatic esterifications of functionalized phenols for the synthesis of lipophilic antioxidants. Biotechnology Letters, 20(2), 131-136.

Chaiyasit, W., Elias, R. J., McClements, D. J., & Decker, E. A. (2007). Role of physical structures in bulk oils on lipid oxidation. Critical Reviews in Food Science and Nutrition, 47(3), 299-317.


Decker, E. A. (1998). Strategies for manipulating the prooxidative/antioxidative balance of foods to maximize oxidative stability. Trends in Food Science & Technology, 9(6), 241-248.






Kikuzaki, H., Hisamoto, M., Hirose, K., Akiyama, K., & Taniguchi, H. (2002). Antioxidant properties of ferulic acid and its related compounds. Journal of Agricultural and Food Chemistry, 50(7), 2161-2168.

Laguerre, M., Giraldo, L. J. L., Lecomte, J., Figueroa-Espinoza, M. C., Barea, B., Weiss, J., Decker, E. A., & Villeneuve, P. (2009). Chain Length Affects Antioxidant Properties of Chlorogenate Esters in Emulsion: The Cutoff Theory Behind the Polar Paradox. Journal of Agricultural and Food Chemistry, 57(23), 11335-11342.

Manach, C., Scalbert, A., Morand, C., Remesy, C., & Jimenez, L. (2004). Polyphenols: food sources and bioavailability. American Journal of Clinical Nutrition, 79(5), 727-747.


Norton, R. A. (1995). Quantitation of Steryl Ferulate and p-Coumarate Esters from Corn and Rice. Lipids, 30(3), 269-274.

Nyström, L., Makinen, M., Lampi, A. M., & Piironen, V. (2005). Antioxidant activity of steryl ferulate extracts from rye and wheat bran. Journal of Agricultural and Food Chemistry, 53(7), 2503-2510.



114


Porter, W. L., Black, E. D., & Drolet, A. M. (1989). Use of Polyamide Oxidative Fluorescence Test on Lipid Emulsions - Contrast in Relative Effectiveness of Antioxidants in Bulk Versus Dispersed Systems. Journal of Agricultural and Food Chemistry, 37(3), 615-624.

Saito, S., Okamoto, Y., & Kawabata, J. (2004). Effects of alcoholic solvents on antiradical abilities of protocatechuic acid and its alkyl esters. Bioscience Biotechnology and Biochemistry, 68(6), 1221-1227.


Schär, A., & Nyström, L. (2016). Enzymatic synthesis of steryl ferulates. European Journal of Lipid Science and Technology, doi: 10.1002/ejlt.201500586.

Seitz, L. M. (1989). Stanol and Sterol Esters of Ferulic and p-Coumaric Acids in Wheat, Corn, Rye, and Triticale. Journal of Agricultural and Food Chemistry, 37(3), 662-667.

Snook, M. E., Data, E. S., & Kays, S. J. (1994). Characterization and Quantitation of Hexadecyl, Octadecyl, and Eicosyl Esters of P-Coumaric Acid in the Vine and Root Latex of Sweet-Potato [Ipomoea-Batatas (L.) Lam.]. Journal of Agricultural and Food Chemistry, 42(11), 2589-2595.



Stockmann, H., Schwarz, K., & Huynh-Ba, T. (2000). The influence of various emulsifiers on the partitioning and antioxidant activity of hydroxybenzoic acids and their derivatives in oil-in-water emulsions. Journal of the American Oil Chemists Society, 77(5), 535-542.

Takagi, T., & Iida, T. (1980). Antioxidant for Fats and Oils from Canary Seed - Sterol and Triterpene Alcohol Esters of Caffeic Acid. Journal of the American Oil Chemists Society, 57(10), 326-330.


Tan, Z., & Shahidi, F. (2012). A novel chemoenzymatic synthesis of phytosteryl caffeates and assessment of their antioxidant activity. Food Chemistry, 133(4), 1427-1434.

Tan, Z., & Shahidi, F. (2013). Phytosteryl sinapates and vanillates: chemoenzymatic synthesis and antioxidant capacity assessment. Food Chemistry, 138(2-3), 1438-1447.

Tan, Z., & Shahidi, F. (2014). Antioxidant activity of phytosteryl phenolates. European Journal of Lipid Science and Technology, 116(12), 1701-1707.

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.



115


Xu, Z. M., & Godber, J. S. (2001). Antioxidant activities of major components of gamma-oryzanol from rice bran using a linoleic acid model. 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.

Part B: Hydrolysis of alkyl ferulates by feruloyl esterases

117

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.


118

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


119

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


120

lipophilicity were synthesized and evaluated as substrates for four types of feruloyl esterases

and one lipase as control.


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.,


121

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


122

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.


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.


123

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,

70.74 s-1 and 91 mM-1∙s-1 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

earlier and were found to be 270 µM, 6.4 s-1 and 23.7 mM-1∙s-1 for Km, kcat and kcat/Km,

respectively (Topakas et al., 2012). Comparing to that study, the turnover number obtained

matches quite well (8.8 s-1), 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.


124

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 ferulate

Ethyl ferulate

Propyl ferulate

Hexyl ferulate

Decyl ferulate

Octadecyl ferulate

AnFaeA

Km [µM] 1123 (71) 611 (60) 245 (18) 40 (3.9) 8 (2.0) n.d.

kcat [s-1

] 32.9 (1.2) 29.4 (1.2) 44.6 (1.2) 9.8 (0.3) 4.6 (0.3)

kcat/Km [mM-1

∙s-1

] 29 (2) 48 (5) 182 (14) 243 (25) 547 (136)

R2 0.996 0.985 0.989 0.948 0.709

n 9 9 11 13 11

a

MtFaeB

Km [µM] 51 (3.4) 48 (2.7) 27 (1.7) 10 (0.9) >0 n.d.

kcat [s-1

] 8.8 (0.3) 11.2 (0.4) 12.1 (0.4) 8.9 (0.3)

kcat/Km [mM-1

∙s-1

] 173 (13) 236 (15) 452 (32) 906 (89)

R2 0.988 0.991 0.985 0.918

n 9 9 9 13

CtFae

Km [µM] 2472 (170) 2578 (152) 1237 (358) 29 (5) 125 (27) >0

kcat [s-1

] 8.0 (0.3) 5.7 (0.2) 3.2 (0.5) 0.2 (0.006) 0.4 (0.02)

kcat/Km [mM-1

∙s-1

] 3.2 (0.3) 2.2 (0.1) 2.6 (0.8) 5.6 (0.9) 3.2 (0.7)

R2 0.994 0.996 0.93 0.909 0.907

n 6 6 10 11 10

ROFae

Km [µM] 134 (17) 149 (16) 81 (8) 27 (2.5) 3.3c (0.8) n.d.

kcat [s-1

] 33.5 (4.9) 30.7 (4.5) 31.7 (4.6) 6.1 (0.9) 2.6 (0.4)

kcat/Km [mM-1

∙s-1

] 250 (48) 206 (37) 391 (68) 225 (39) 780 (213)

R2 0.962 0.973 0.976 0.937 0.636

n 8 8 9 13 11

Lipase

Km [µM] 413 (79) 636 (168) 1848

b

(401) 88 (25) 146 (33) >0

kcat [s-1

] 0.002

(0.0002) 0.004

(0.0004) 0.022

(0.0030) 0.006

(0.0004) 0.010

(0.0009)

kcat/Km [mM-1

∙s-1

] 0.005

(0.001) 0.006

(0.002) 0.012

(0.003) 0.07 (0.02) 0.07 (0.02)

R2 0.941 0.939 0.979 0.811 0.894

n 7

a 7

a 10

a 9

a 8

Numbers in parentheses represent the estimated standard errors. R

2 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


125

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


126

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


127

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. The change in n-alkyl

ferulates concentration in comparison to the total liberated ferulic acid may be researched.

Potentially feruloyl esterases play a minor role in the natural decomposition and digestion of

nonpolar n-alkyl ferulates compared to lipases.

5. Acknowledgements

This study was financially supported by Swiss National Science Foundation, SNSF (project

200021_141268) and ETH Zurich.


128

References

Adom, K. K., Sorrells, M. E., & Liu, R. H. (2005). Phytochemicals and antioxidant activity of milled fractions of different wheat varieties. Journal of Agricultural and Food Chemistry, 53(6), 2297-2306.

Aliwan, F. O., Kroon, P. A., Faulds, C. B., Pickersgill, R., & Williamson, G. (1999). Ferulic acid esterase-III from Aspergillus niger does not exhibit lipase activity. Journal of the Science of Food and Agriculture, 79(3), 457-459.

Andreasen, M. F., Kroon, P. A., Williamson, G., & Garcia-Conesa, M. T. (2001). Esterase activity able to hydrolyze dietary antioxidant hydroxycinnamates is distributed along the intestine of mammals. Journal of Agricultural and Food Chemistry, 49(11), 5679-5684.

Benoit, I., Danchin, E. G. J., Bleichrodt, R. J., & de Vries, R. P. (2008). Biotechnological applications and potential of fungal feruloyl esterases based on prevalence, classification and biochemical diversity. Biotechnology Letters, 30(3), 387-396.


Couteau, D., McCartney, A. L., Gibson, G. R., Williamson, G., & Faulds, C. B. (2001). Isolation and characterization of human colonic bacteria able to hydrolyse chlorogenic acid. Journal of Applied Microbiology, 90(6), 873-881.

Crepin, V. F., Faulds, C. B., & Connerton, I. F. (2004). Functional classification of the microbial feruloyl esterases. Applied Microbiology and Biotechnology, 63(6), 647-652.

Derewenda, Z. S., Derewenda, U., & Dodson, G. G. (1992). The Crystal and Molecular-Structure of the Rhizomucor Miehei Triacylglyceride Lipase at 1.9 Angstrom Resolution. Journal of Molecular Biology, 227(3), 818-839.

deVries, R. P., Michelsen, B., Poulsen, C. H., Kroon, P. A., vandenHeuvel, R. H. H., Faulds, C. B., Williamson, G., vandenHombergh, J. P. T. W., & Visser, J. (1997). The faeA genes from Aspergillus niger and Aspergillus tubingensis encode ferulic acid esterases involved in degradation of complex cell wall polysaccharides. Applied and Environmental Microbiology, 63(12), 4638-4644.

Faulds, C. B. (2010). What can feruloyl esterases do for us? Phytochemistry Reviews, 9(1), 121-132.

Faulds, C. B., Kroon, P. A., Saulnier, L., Thibault, J. F., & Williamson, G. (1995). Release of Ferulic Acid from Maize Bran and Derived Oligosaccharides by Aspergillus Niger Esterases. Carbohydrate Polymers, 27(3), 187-190.

Faulds, C. B., Molina, R., Gonzalez, R., Husband, F., Juge, N., Sanz-Aparicio, J., & Hermoso, J. A. (2005). Probing the determinants of substrate specificity of a feruloyl esterase, AnFaeA, from Aspergillus niger. FEBS Journal, 272(17), 4362-4371.

Faulds, C. B., Perez-Boada, M., & Martinez, A. T. (2011). Influence of organic co-solvents on the activity and substrate specificity of feruloyl esterases. Bioresource Technology, 102(8), 4962-4967.

Faulds, C. B., & Williamson, G. (1994). Purification and characterization of a ferulic acid esterase (FAE-III) from Aspergillus niger: specificity for the phenolic moiety and binding to microcrystalline cellulose. Microbiology, 140, 779-787.

Faulds, C. B., & Williamson, G. (1999). The role of hydroxycinnamates in the plant cell wall. Journal of the Science of Food and Agriculture, 79(3), 393-395.


129

Graça, J. (2010). Hydroxycinnamates in suberin formation. Phytochemistry Reviews, 9(1), 85-91.



Hermoso, J. A., Sanz-Aparicio, J., Molina, R., Juge, N., Gonzalez, R., & Faulds, C. B. (2004). The crystal structure of feruloyl esterase A from Aspergillus niger suggests evolutive functional convergence in feruloyl esterase family. Journal of Molecular Biology, 338(3), 495-506.

Juge, N., Williamson, G., Puigserver, A., Cummings, N. J., Connerton, I. F., & Faulds, C. B. (2001). High-level production of recombinant Aspergillus niger cinnamoyl esterase (FAEA) in the methylotrophic yeast Pichia pastoris. FEMS Yeast Research, 1(2), 127-132.

Klibanov, A. M. (1997). Why are enzymes less active in organic solvents than in water? Trends in Biotechnology, 15(3), 97-101.

Koseki, T., Takahashi, K., Fushinobu, S., Iefuji, H., Iwano, K., Hashizume, K., & Matsuzawa, H. (2005). Mutational analysis of a feruloyl esterase from Aspergillus awamori involved in substrate discrimination and pH dependence. Biochimica et Biophysica Acta, 1722(2), 200-208.

Kroon, P. A., Faulds, C. B., Brezillon, C., & Williamson, G. (1997). Methyl phenylalkanoates as substrates to probe the active sites of esterases. European Journal of Biochemistry, 248(1), 245-251.

Li, L., Shewry, P. R., & Ward, J. L. (2008). Phenolic Acids in Wheat Varieties in the HEALTHGRAIN Diversity Screen. Journal of Agricultural and Food Chemistry, 56(21), 9732-9739.


Moukouli, M., Topakas, E., & Christakopoulos, P. (2008). Cloning, characterization and functional expression of an alkalitolerant type C feruloyl esterase from Fusarium oxysporum. Applied Microbiology and Biotechnology, 79(2), 245-254.

Ralet, M. C., Faulds, C. B., Williamson, G., & Thibault, J. F. (1994). Degradation of Feruloylated Oligosaccharides from Sugar-Beet Pulp and Wheat Bran by Ferulic Acid Esterases from Aspergillus-Niger. Carbohydrate Research, 263(2), 257-269.


Schubot, F. D., Kataeva, I. A., Blum, D. L., Shah, A. K., Ljungdahl, L. G., Rose, J. P., & Wang, B. C. (2001). Structural basis for the substrate specificity of the feruloyl esterase domain of the cellulosomal xylanase Z from Clostridium thermocellum. Biochemistry, 40(42), 12524-12532.

Sorensen, A. D., Durand, E., Laguerre, M., Bayrasy, C., Lecomte, J., Villeneuve, P., & Jacobsen, C. (2014). Antioxidant properties and efficacies of synthesized alkyl caffeates, ferulates, and coumarates. Journal of Agricultural and Food Chemistry, 62(52), 12553-12562.

Topakas, E., Christakopoulos, P., & Faulds, C. B. (2005). Comparison of mesophilic and thermophilic feruloyl esterases: Characterization of their substrate specificity for methyl phenylalkanoates. Journal of Biotechnology, 115(4), 355-366.


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Topakas, E., Moukouli, M., Dimarogona, M., & Christakopoulos, P. (2012). Expression, characterization and structural modelling of a feruloyl esterase from the thermophilic fungus Myceliophthora thermophila. Applied Microbiology and Biotechnology, 94(2), 399-411.

Vafiadi, C., Topakas, E., Christakopoulos, P., & Faulds, C. B. (2006). The feruloyl esterase system of Talaromyces stipitatus: Determining the hydrolytic and synthetic specificity of TsFaeC. Journal of Biotechnology, 125(2), 210-221.

Vafiadi, C., Topakas, E., Wong, K. K. Y., Suckling, I. D., & Christakopoulos, P. (2005). Mapping the hydrolytic and synthetic selectivity of a type C feruloyl esterase (StFaeC) from Sporotrichum thermophile using alkyl ferulates. Tetrahedron: Asymmetry, 16(2), 373-379.

Wang, X. K., Geng, X., Egashira, Y., & Sanada, H. (2004). Purification and characterization of a feruloyl esterase from the intestinal bacterium Lactobacillus acidophilus. Applied and Environmental Microbiology, 70(4), 2367-2372.

Wu, X. Y., Jaaskelainen, S., & Linko, Y. Y. (1996). Purification and partial characterization of Rhizomucor miehei lipase for ester synthesis. Applied Biochemistry and Biotechnology, 59(2), 145-158.

Zerner, B., Bond, R. P. M., & Bender, M. L. (1964). 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.

Conclusion and Outlook

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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


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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


133

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.

134

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.

Documents

Hydroxycinnamic Acid Esters Enzymatic Synthesis and