Research Collection Doctoral Thesis Enzymatic Synthesis and Hydrolysis of Linear Alkyl and Steryl Hydroxycinnamic Acid Esters Author(s): Schär, Aline Lea Publication Date: 2016 Permanent Link: https://doi.org/10.3929/ethz-a-010670414 Rights / License: In Copyright - Non-Commercial Use Permitted This page was generated automatically upon download from the ETH Zurich Research Collection . For more information please consult the Terms of use . ETH Library
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Research Collection
Doctoral Thesis
Enzymatic Synthesis and Hydrolysis of Linear Alkyl and SterylHydroxycinnamic Acid Esters
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.
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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).
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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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
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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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
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
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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
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
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
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