hydroxycinnamic acid esters enzymatic synthesis and
TRANSCRIPT
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
Rights / License: In Copyright - Non-Commercial Use Permitted
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ETH Library
DISS. ETH NO. 23265
Enzymatic Synthesis and Hydrolysis of Linear Alkyl and Steryl
Hydroxycinnamic Acid Esters
A thesis submitted to attain the degree of
DOCTOR OF SCIENCES of ETH ZURICH
(Dr. sc. ETH Zurich)
presented by
Aline Lea Schär
MSc ETH in Food Science, ETH Zurich
born on 24.01.1987
citizen of Madiswil (BE)
accepted on the recommendation of
Prof. Dr. Laura Nyström, examiner
Dr. Pierre Villeneuve, co-examiner
Prof. Dr. Evangelos Topakas, co-examiner
2016
Wir treten auf. Wir spielen. Wir treten ab.
Moritz Leuenberger
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
Hydroxycinnamic acids
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
Hydroxycinnamic acids
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
Hydroxycinnamic acids
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
Hydroxycinnamic acids
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
Hydroxycinnamic acids
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
Hydroxycinnamic acids
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
Hydroxycinnamic acids
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.
General enzymatic reactions
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)
General enzymatic reactions
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
General enzymatic reactions
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
General enzymatic reactions
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.
Enzymatic lipophilization of hydroxycinnamic acids
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)
Hydroxycinnamic acid derivatives,
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)
Hydroxycinnamic acid derivatives,
methyl or ethyl esters thereof, oleyl
alcohol
Lipozyme RM IM,
Novozym 435 Solvent-free, 80 kPa
(Vosmann et al.,
2006) (Weitkamp et
al., 2006)
Enzymatic lipophilization of hydroxycinnamic acids
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.
Enzymatic lipophilization of hydroxycinnamic acids
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.
Enzymatic lipophilization of hydroxycinnamic acids
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.
Enzymatic lipophilization of hydroxycinnamic 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
Enzymatic lipophilization of hydroxycinnamic acids
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
n-hexane/butanol/water
CLEAs of A. niger
type A feruloyl
esterase
(Vafiadi et al., 2008a)
Methyl ferulate, 1-butanol in
n-hexane/butanol/water
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
Enzymatic lipophilization of hydroxycinnamic acids
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.
Esterification of phytosterols
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
Immobilized C. antarctica
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)
Esterification of phytosterols
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
Immobilized C. antarctica
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)
Esterification of phytosterols
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
Esterification of phytosterols
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
Esterification of phytosterols
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
Esterification of phytosterols
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
Esterification of phytosterols
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
Esterification of phytosterols
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.
Esterification of phytosterols
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
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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
Part B: Enzymatic esterification of ferulic acid
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
Part B: Enzymatic esterification of ferulic acid
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.
Part B: Enzymatic esterification of ferulic acid
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.
Part B: Enzymatic esterification of ferulic acid
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
Part B: Enzymatic esterification of ferulic acid
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
Part B: Enzymatic esterification of ferulic acid
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
Part B: Enzymatic esterification of ferulic acid
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%.
Part B: Enzymatic esterification of ferulic acid
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.
Part B: Enzymatic esterification of ferulic acid
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
Part B: Enzymatic esterification of ferulic acid
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).
Part B: Enzymatic esterification of ferulic acid
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.
Part B: Enzymatic esterification of ferulic acid
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%.
Part B: Enzymatic esterification of ferulic acid
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
Part B: Enzymatic esterification 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.
Part B: Enzymatic esterification of ferulic acid
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.
Part B: Enzymatic esterification of ferulic acid
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.
Part B: Enzymatic esterification of ferulic acid
72
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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
Part B: Enzymatic synthesis of steryl ferulates
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.
Part B: Enzymatic synthesis of steryl ferulates
77
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
Part B: Enzymatic synthesis of steryl ferulates
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. Materials and Methods
2.1 Chemicals and enzymes
Ferulic acid, ≥99% and β-sitosterol, ≥70% (impurities being mainly campesterol and
β-sitostanol) were obtained from Sigma-Aldrich, Switzerland. γ-oryzanol, ≥ 99%, was
purchased from Wako Pure Chemical Industries, Japan. Ethyl ferulate was purchased from
Alfa Aesar, Germany (98% purity), and also synthesized as reported earlier (Schär &
Nyström, 2015). All used solvents were of HPLC grade and all enzymes were purchased
from Sigma-Aldrich, Switzerland. Five lipases were used in this study, namely lipase from
Rhizomucor miehei (formerly known as mucor miehei) immobilized on macroporous
ion-exchange resin, >30 U/g (1 U sets free 1 μmol stearic acid at pH 8.0 and 70°C per
minute), lipase from C. rugosa type VII, ≥11.7 U/mg (at pH 7.2 and 37°C 1 U will hydrolyze
1.0 microequivalent of fatty acid from a triglyceride per minute), lipase A from C. antarctica
immobilized on Immobead 150, recombinant from Aspergillus oryzae ≥500 U/g (1 U
corresponds to the amount of enzyme which liberates 1 μmol butyric acid per minute at pH
10.0 and 40°C), lipase type B from C. antarctica, recombinant from A. niger, immobilized on
acrylic resin ≥5,000 U/g (propyl laurate units), and lipase from C. rugosa, immobilized on
Part B: Enzymatic synthesis of steryl ferulates
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
Part B: Enzymatic synthesis of steryl ferulates
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:
Part B: Enzymatic synthesis of steryl ferulates
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.
Part B: Enzymatic synthesis of steryl ferulates
82
3. Results and Discussion
3.1 Optimization of transesterification
In a first step different immobilized lipases were tested on their activity to transesterify ferulic
acid from ethyl ferulate to β-sitosteryl ferulate. For the duplicate sample with the immobilized
C. rugosa lipase an average molar yield of 9.2% was measured. For the lipase A from
C. antarctica and the lipase from R. miehei a very small amount of steryl ferulates could be
detected, however, smaller than the quantification limit. In the samples with C. antarctica
lipase B no clear difference to the blank could be measured after 5 days of incubation. This
findings correspond well with other studies, were C. rugosa lipase has been the only lipase
able to catalyze the synthesis of steryl ferulates starting from vinyl ferulate (Chigorimbo-
Murefu et al., 2009; Tan & Shahidi, 2011). Recently the C. antarctica lipase A was shown to
esterify sterols with fatty acids most efficiently (Panpipat et al., 2013). However, based on
this data it seems that more complex acid substrates such as hydroxycinnamic acids are not
amongst good substrates of C. antarctica lipase A. Conclusively, a lipase from C. rugosa was
selected for later use. However, since the yield of steryl ferulates with the immobilized lipase
was rather low and the enzyme amount very high, a non-immobilized enzyme preparation
with a higher activity was chosen with the lipase type VII from C. rugosa.
The transesterification of ferulic aid from ethyl ferulate to sitosteryl ferulate using C. rugosa
lipase was optimized regarding four parameters: temperature, enzyme-to-sterol ratio, sterol
amount, and substrate ratio (Table 2). The three center points gave a yield of 48.7(2.9)% and
the experimental data was fitted to the second-order polynomial equation (equation 1). The
analysis of variance (Table 3) shows a strong correlation between the model and the
experimental data, as indicated by the low p-values for all model variables and a very high R2
and lack of fit.
Part B: Enzymatic synthesis of steryl ferulates
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
Part B: Enzymatic synthesis of steryl ferulates
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.
Direct esterification Transesterification
β-coefficient p-value β-coefficient p-value
β0 23.33 48.73
x1 2.66 0.00* 5.15 0.00*
x2 7.58 0.00* 10.85 0.00*
x3 2.33 0.01* 1.13 0.06
x4 -0.58 0.44 1.71 0.01*
x1* x2 0.05 0.97 0.28 0.78
x1* x3 1.88 0.16 3.00 0.01*
x2* x3 -2.98 0.03* -1.08 0.28
x1* x4 -1.85 0.16 -0.82 0.40
x2* x4 -1.48 0.26 4.45 0.00*
x3* x4 -1.65 0.21 -1.15 0.25
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
Part B: Enzymatic synthesis of steryl ferulates
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.
Direct esterification Transesterification
x1: temperature [°C] 63 63
x2: enzyme-to-sterol ratio [g/g] 3 3
x3: sterol amount [mg/3 mL] 23.8 11.2
x4: substrate ratio [mol/mol] 1 2.5
Predicted Yield [%] 31.0 57.2
Measured Yield [%] 34.8 (1.5); n=10 54.9 (2.5); n=9
n= number of conducted replicates, yield reflects the molar percentage of sterols (β-sitosterol
and sterol impurities campesterol and β-sitostanol) converted to steryl ferulates.
Part B: Enzymatic synthesis of 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
Part B: Enzymatic synthesis of steryl ferulates
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.,
Part B: Enzymatic synthesis of steryl ferulates
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
Part B: Enzymatic synthesis of steryl ferulates
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
Part B: Enzymatic synthesis of steryl ferulates
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.
Part B: Enzymatic synthesis of steryl ferulates
91
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Maeda, A., Mizuno, T., Bunya, M., Sugihara, S., Nakayama, D., Tsunasawa, S., Hirota, Y., & Sugihara, A. (2008). Characterization of novel cholesterol esterase from Trichoderma sp. AS59 with high ability to synthesize steryl esters. Journal of Bioscience and Bioengineering, 105(4), 341-349.
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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.
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Weitkamp, P., Vosmann, K., & Weber, N. (2006). Highly efficient preparation of lipophilic hydroxycinnamates by solvent-free lipase-catalyzed transesterification. Journal of Agricultural and Food Chemistry, 54(19), 7062-7068.
Wilson, T. A., Nicolosi, R. J., Woolfrey, B., & Kritchevsky, D. (2007). Rice bran oil and oryzanol reduce plasma lipid and lipoprotein cholesterol concentrations and aortic cholesterol ester accumulation to a greater extent than ferulic acid in hypercholesterolemic hamsters. Journal of Nutritional Biochemistry, 18(2), 105-112.
Winkler-Moser, J. K., Hwang, H. S., Bakota, E. L., & Palmquist, D. A. (2015). Synthesis of steryl ferulates with various sterol structures and comparison of their antioxidant activity. Food Chemistry, 169, 92-101.
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Yu, Y., Zheng, Y., Quan, J., Wu, C. Y., Wang, Y. J., Branford-White, C., & Zhu, L. M. (2010). Enzymatic Synthesis of Feruloylated Lipids: Comparison of the Efficiency of Vinyl Ferulate and Ethyl Ferulate as Substrates. Journal of the American Oil Chemists Society, 87(12), 1443-1449.
Zheng, Y., Wu, X. M., Branford-White, C., Ning, X., Quan, J., & Zhu, L. M. (2009). Enzymatic synthesis and characterization of novel feruloylated lipids in selected organic media. Journal of Molecular Catalysis B: Enzymatic, 58(1-4), 65-71.
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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%
Part B: Enzymatic synthesis of steryl ferulates
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%
Part B: Enzymatic synthesis of steryl ferulates
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
Part B: Esterification of hydroxycinnamic acids and their antioxidant activity
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
Part B: Esterification of hydroxycinnamic acids and their antioxidant activity
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
Part B: Esterification of hydroxycinnamic acids and their antioxidant activity
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.
Part B: Esterification of hydroxycinnamic acids and their antioxidant activity
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
Part B: Esterification of hydroxycinnamic acids and their antioxidant activity
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
Part B: Esterification of hydroxycinnamic acids and their antioxidant activity
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
Part B: Esterification of hydroxycinnamic acids and their antioxidant activity
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.
Part B: Esterification of hydroxycinnamic acids and their antioxidant activity
105
3. Results and Discussion
3.1 Esterification of cinnamic acid derivatives by R. miehei lipase
The esterification of the hydroxycinnamic acid was optimized for each acid individually. The
yield of ethyl sinapate in comparison with ethyl ferulate was slightly lower, also the amount of
immobilized enzyme needed was little higher (enzyme-to-substrate ratio (m/m) of 3.1 instead
of 2.5) (Table 1). Thus, the second methoxy group does not influence the enzymatic
esterification strongly. On the other hand, esterification yield was drastically lower for caffeic
acid (yield 16.1%), which has a second hydroxyl group in meta-position, instead of a
methoxy group. This is in accordance to previous published work, where immobilized R.
miehei lipase was employed in ionic liquid to esterify hydroxycinnamic acids with octanol
(Katsoura et al., 2009). In this earlier study, the yields of octyl ferulate and octyl sinapate
were similar, whereas the yield of octyl caffeate was significantly lower. Also in solvent-free
reaction system with 1-octanol, ferulic acid was esterified more efficiently than caffeic acid by
immobilized R. miehei lipase (Stamatis et al., 1999).
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)
Part B: Esterification of hydroxycinnamic acids and their antioxidant activity
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).
Part B: Esterification of hydroxycinnamic acids and their antioxidant activity
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.
Part B: Esterification of hydroxycinnamic acids and their antioxidant activity
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
Part B: Esterification of hydroxycinnamic acids and their antioxidant activity
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).
Part B: Esterification of hydroxycinnamic acids and their antioxidant activity
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.
Part B: Esterification of hydroxycinnamic acids and their antioxidant activity
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.
Part B: Esterification of hydroxycinnamic acids and their antioxidant activity
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.
Part B: Esterification of hydroxycinnamic acids and their antioxidant activity
113
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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.
Part B: Hydrolysis of alkyl ferulates by feruloyl esterases
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
Part B: Hydrolysis of alkyl ferulates by feruloyl esterases
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
Part B: Hydrolysis of alkyl ferulates by feruloyl esterases
120
lipophilicity were synthesized and evaluated as substrates for four types of feruloyl esterases
and one lipase as control.
2. Materials and Methods
2.1 Chemicals
Ferulic acid (≥99%), MOPS (3-(N-morpholino)propanesulfonic acid, ≥99.5%) and MES (2-(N-
morpholino)ethanesulfonic acid, ≥99%) were obtained from Sigma-Aldrich, Buchs,
Switzerland. Methyl ferulate (99%) and ethyl ferulate (98%) were purchased from Alfa Aesar,
Germany. γ-Oryzanol was obtained from Wako Pure Chemical Industries, Osaka, Japan. All
solvents used were of HPLC grade or of higher purity.
2.2 Enzymes
Lipozyme® RM IM was provided by Novozymes A/S, Bagsvaerd, Denmark. Feruloyl
esterases from rumen microorganism, ROFae (600 U/mL where 1 U corresponds to 1 µmol
ferulic acid released from ethyl ferulate per minute at pH 6.5 and 40°C) and from XynZ
domain of Clostridium thermocellum, CtFae (10 U/mL where 1 U corresponds to 1 µmol
ferulic acid released from ethyl ferulate per minute at pH 6 and 50°C) were obtained from
Megazyme, Bray, Ireland. Recombinant feruloyl esterase type-A from A. niger, AnFaeA, was
produced according to Juge and co-workers (Juge et al., 2001). The lyophilized enzyme was
redissolved in buffer (MOPS, pH 6). The type-B feruloyl esterase from Myceliophthora
thermophila, MtFaeB, was prepared according to Topakas et al. without the chromatographic
purification (Topakas et al., 2012). Lipase from Rhizomucor miehei (≥20000 U/g) was
purchased from Sigma-Aldrich, Buchs, Switzerland. Protein contents of enzyme preparations
were analyzed by Bradford assay using Bradford reagent from Sigma-Aldrich, Buchs,
Switzerland and bovine serum albumin as standard.
2.3 Preparation of n-alkyl ferulates
Propyl, hexyl, decyl and octadecyl ferulates (Figure 1) were enzymatically esterified using
Lipozyme® RM IM as published earlier (Schär & Nyström, 2015). To remove the ferulic acid
from the propyl ferulate, the reaction mixture in n-hexane was washed with water. After
evaporation of the unreacted propanol and the solvent n-hexane at 50°C, the propyl ferulate
product was redissolved in acetone and ready for hydrolytic reactions. The other ferulates
were purified by a base-acid wash adapted from Hakala and co-workers (Hakala et al.,
Part B: Hydrolysis of alkyl ferulates by feruloyl esterases
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
Part B: Hydrolysis of alkyl ferulates by feruloyl esterases
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.
3. Results and Discussion
The kinetic constants using the Michaelis-Menten equation were determined for four feruloyl
esterases and one control lipase using methyl, ethyl, propyl, hexyl, and decyl ferulate as
substrates (Table 1). For the substrate with the longest alkyl chain, the octadecyl ferulate, no
hydrolysis could be measured for AnFaeA, MtFaeB and ROFae, even if the incubation time
was increased to 24h. In contrast, CtFae and the control lipase liberated ferulic acid,
however the activity was too low to determine kinetic constants. Generally, Km and kcat values
decreased with increasing chain length for the feruloyl esterases. Although with increasing
lipophilicity of the substrate Km is decreasing stronger compared to the kcat values, the
catalytic efficiency kcat/Km is increasing mainly in the case of AnFaeA and MtFaeB. For CtFae
and the control lipase the pattern was not as clear. Also the coefficient of determination (R2)
of the experimental data fitted to the Michaelis-Menten kinetics showed a decreasing trend
with increasing chain length of the n-alkyl ferulate.
Part B: Hydrolysis of alkyl ferulates by feruloyl esterases
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.
Part B: Hydrolysis of alkyl ferulates by feruloyl esterases
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
Part B: Hydrolysis of alkyl ferulates by feruloyl esterases
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
Part B: Hydrolysis of alkyl ferulates by feruloyl esterases
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
Part B: Hydrolysis of alkyl ferulates by feruloyl esterases
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.
Part B: Hydrolysis of alkyl ferulates by feruloyl esterases
128
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Conclusion and Outlook
131
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
Conclusion and Outlook
132
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
Conclusion and Outlook
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.