Production of canolol from canola meal phenolics via hydrolysis and microwave-induced decarboxylation

Article abstract of DOI:10.1007/s11746-013-2345-6

A potent antioxidant, anti-inflammatory and anti-mutagenic agent; 4-vinyl-2,6-dimethoxyphenol (canolol) was obtained from canola meal in a significant yield via alkaline (NaOH)/enzymatic (ferulic acid esterase) hydrolysis followed by microwave-assisted decarboxylation. The hydrolysis was carried out either through using canola meal directly as a substrate or by using the 70 % aqueous methanolic extract filtrates. The hydrolyzed extracts underwent RP-HPLC analysis which showed that 81.0 and 94.8 % of the total phenolics were hydrolyzed to sinapic acid after the alkaline hydrolysis of the meal and the methanolic extracts, respectively. The enzymatic hydrolysis showed lower conversion rates (49.5 and 58.3 %). The hydrolyzed extracts were consequently decarboxylated using 8-diazabicyclo[5.4.0]undec-7-ene under microwave irradiation at different conditions. The HPLC profiling of decarboxylated extracts showed that using microwave at 300 Wof microwave power for 12 min brought the highest sinapic acid conversion to canolol (58.3 %) yielding 4.2 mg canolol from each gram of canola meal suggesting that the process could be commercially economical. AOCS 2013.

Full text of DOI:10.1007/s11746-013-2345-6

J Am Oil Chem Soc (2014) 91:89–97
DOI 10.1007/s11746-013-2345-6
ORIGINAL PAPER
Production of Canolol from Canola Meal Phenolics via Hydrolysis
and Microwave-Induced Decarboxylation
Rabie Y. Khattab
Usha Thiyam-Hollander
•
Michael N. A. Eskin
•
Received: 5 June 2013 / Revised: 16 August 2013 / Accepted: 4 September 2013 / Published online: 29 September 2013
Ó AOCS 2013
Abstract A potent antioxidant, anti-inflammatory and
anti-mutagenic agent; 4-vinyl-2,6-dimethoxyphenol (can-
olol) was obtained from canola meal in a significant yield
via alkaline (NaOH)/enzymatic (ferulic acid esterase)
hydrolysis followed by microwave-assisted decarboxyl-
ation. The hydrolysis was carried out either through using
canola meal directly as a substrate or by using the 70 %
aqueous methanolic extract filtrates. The hydrolyzed
extracts underwent RP-HPLC analysis which showed that
Keywords Antioxidant Á 4-Vinyl-2,
6-dimethoxyphenol (canolol) Á Canola meal Á
Hydrolysis Á Microwave Á Decarboxylation
Introduction
Free radicals induce oxidative damage in vivo causing
aging and various diseases [1, 2]. They are also the major
cause for deterioration, quality loss, and shelf life reduction
in oils/fats and fat-containing systems. There are many
studies on the antioxidative effects of natural extracts
derived from rapeseed [3, 4], vegetables [5, 6] and olives
[7]. Most of these extracts contain water-soluble compo-
nents as active ingredients. Thus, there is a need for nat-
urally derived lipid-soluble antioxidants for utilization in
oil/fat products as well as in vivo. Examples of lipid-sol-
uble antioxidants include butylhydroxyanisole (BHA), bu-
tylhydroxytoluene (BHT), tert-butylhydroquinone (TBHQ)
and propylgallate (PG). Their safety has been extensively
questioned and attempts to eliminate them from human diet
continued. Consequently, there is a need for developing
highly active fat-soluble antioxidants from natural sources,
especially from the under-utilized by-products such as
meals.
8
1.0 and 94.8 % of the total phenolics were hydrolyzed to
sinapic acid after the alkaline hydrolysis of the meal and
the methanolic extracts, respectively. The enzymatic
hydrolysis showed lower conversion rates (49.5 and
5
8.3 %). The hydrolyzed extracts were consequently
decarboxylated using 8-diazabicyclo[5.4.0]undec-7-ene
under microwave irradiation at different conditions. The
HPLC profiling of decarboxylated extracts showed that
using microwave at 300 W of microwave power for 12 min
brought the highest sinapic acid conversion to canolol
(
58.3 %) yielding 4.2 mg canolol from each gram of
canola meal suggesting that the process could be com-
mercially economical.
R. Y. Khattab
Food Science Department, Faculty of Agriculture (Saba Basha),
Alexandria University, Alexandria, Egypt
2,6-Dimethoxy-4-vinylphenol (known as canolol or
4-vinylsyringol) is a well-known lipid-soluble potent anti-
oxidant and antimutagenic compound [1, 2, 8, 9] formed in
canola oil via sinapic acid decarboxylation during oil
pressing at high temperature and pressure [1, 2]. The anti-
radical scavenging activity of canolol is much greater than
that of well-known antioxidants, including vitamin C, b-
carotene, a-tocopherol, rutin and quercetin [10]. Unluckily,
canolol is almost completely lost during oil refining [10,
11] which advocates its isolation from meal and adding it
M. N. A. Eskin Á U. Thiyam-Hollander (&)
Human Nutritional Sciences Department, University
of Manitoba, Winnipeg, MB, Canada
e-mail: thiyam@cc.umanitoba.ca;
usha.thiyam@ad.umanitoba.ca
U. Thiyam-Hollander
Richardson Centre for Functional Foods and Nutraceuticals,
Winnipeg, MB, Canada
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J Am Oil Chem Soc (2014) 91:89–97
back to the oil. The lipophilic characteristics of canolol
might account for its high affinity to the cell membranes
and other biological membranes and hence its reactivity
inside the body where water-soluble antioxidants are hard
to react, thus establishing its outstanding role. The indus-
trial demand of vinyl phenols and canolol is relatively
satisfied by chemical synthesis and not from natural sour-
ces. Investigations have explored large scale synthesis of
decarboxylation of the hydrolyzed substrates under
microwave irradiation. The conversion of sinapine and
other sinapic acid derivatives to canolol will pave the way
for new and unlimited opportunities, especially for by-
product utilization, canola oil refining and value-added
processing of canola meal.
4
-vinylphenols through microbial or chemical decarbox-
Materials and Methods
ylation of cinnamic acids such as p-coumaric acid, ferulic
acid, sinapic acid and caffeic acid from plant sources
including barley [12], wheat bran [13] and sunflower seeds
Materials
[
14].
Investigations to obtain this bioactive compound from
Canola seeds were procured from commercial suppliers
including ADM Bunge Canada (Oakville, ON, Canada),
Viterra Canola (Ste. Agathe, MB, Canada), and Formely
Canbra Foods Ltd. and currently Richardson (Lethbridge,
AB, Canada). Phenolic standards; sinapic, ferulic, caffeic,
trans-cinnamic, syringic, p-coumaric, salicylic, vanillic,
gallic acid, kaempferol and syringaldehyde were pur-
chased from Sigma-Aldrich (St. Louis, MO, USA). Sina-
pine thiocyanate was purchased from EPL Bio Analytical
Services (Niantic, IL, USA). Authentic standard of syn-
thetic canolol was kindly provided by Dr. Amy Logan,
CSIRO Animal Food and Health Sciences, Werribee,
Australia and Dr. Bertrand Mathaus, Max Rubner Insti-
tute, Germany. Sinapoyl glucose were kindly donated by
Dr. A. Baumert, IPB-Halle, Germany. Sinapinaldehyde
was purchased from ChromaDex, Inc. (Irvine, CA, USA).
Ferulic acid esterase (FAE) was kindly donated by
Novozymes (Bagsvaerd, Denmark). All solvents (analyti-
cal and HPLC grades) were purchased from Sigma-
Aldrich (Oakville, ON, Canada).
the natural sources are rare and not really successful.
Although many attempts were tried, the amounts obtained
from this compound from either natural sources or chem-
ical synthesis were not able to meet the ever-increasing
industrial demand due to the limited amount of this com-
pound in plant sources and the less yield and high cost in
the chemical synthesis. Consequently, there is a growing
interest in developing alternative natural sources and more
economic procedures to obtain this phenol in significant
amounts to fulfill the industry demand.
Microwave pre-treatment of seeds is a simple and
useful method for production of higher quality and more
functional vegetable oils. In addition to improving the oil
yield and quality, lowering energy consumption and
reducing processing time and solvent consumption, it
offers better retention and availability of desirable nu-
traceuticals including phytosterols, tocopherols and phe-
nolic compounds in the extracted oils [15]. In the past
few years, microwave-assisted chemical synthesis was
used extensively as an environmentally friendly, rapid,
and high-yielding technique in a laboratory scale [16].
Currently, efforts are being made to develop bulky
reactors that will allow the scaling up of this technique
to fit the industrial needs [17]. It is well documented that
microwave irradiation has been used to enhance the
decarboxylation of organic compounds [18–22]. Nomura
et al. [23] reported a base-catalyzed decarboxylation and
amide-forming reaction of substituted cinnamic acids via
microwave heating. However, in these experimental
conditions, the recovery and the yields of the 4-vinyl-
phenols were very low. Bernini et al. [22] described new
experimental conditions to achieve satisfactory yield
from the decarboxylation of pure sinapic acid under
microwave irradiation in the presence of 1,8-diazabicy-
clo[5.4.0]undec-7-ene (DBU), aluminum oxide, and
hydroquinone.
Experimental Procedures
Sample Preparation
Canola seeds were cleaned, ground, and sieved through a
1,410-lm screen. The meals were obtained by defatting the
seeds using a Soxtec 2050 system (Foss-Tecator, Foss
North America, MN, USA) according to FOSFA [24] in
two cycles. The meals were de-solventized overnight at
room temperature in the fume hood.
Extraction of Canola Meal Phenolics
Phenolic compounds were extracted from the meal
according to the procedure of Thiyam et al. [25]. Meal
samples (1 g each) were extracted three times with 9 mL of
70 % methanol using ultra-sonication (1 min) followed by
centrifugation under refrigerated conditions (10 min at
5,0009g) and filtered through Whatman No. 1 filter paper.
This paper reports a new procedure for the production of
canolol from under-utilized canola meal through a
sequence of alkaline/enzymatic hydrolysis followed by
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J Am Oil Chem Soc (2014) 91:89–97
91
The filtrates from the three extractions were combined and
made up to a total volume of 25 mL with the extraction
solution.
conical glass flask and 100 lL of 1,8-diazabicy-
clo[5.4.0]undec-7-ene (DBU), 5 mg hydroquinone and
2.0 g aluminum oxide were added. The mixture was vor-
texed and the solvent was completely evaporated under N2
at low pressure. The flasks were then heated in a micro-
wave oven (Panasonic, Model NN-CS597S, 1,000 W) at
different power levels (power 1; 100 W to power 7;
700 W) for different times (1–25 min). The flasks were
cooled at room temperature, neutralized with 1 M HCl and
extracted with 1:1 (v/v) diethyl ether/ethyl acetate mixture
(3 9 5 mL) using a refrigerated centrifuge (4 °C) at
5,0009g for 10 min. The organic layers were combined
Alkaline Hydrolysis of Phenolic Compounds
Hydrolyzing Phenolic Extracts
Twenty milliliters of 4 M NaOH was added to 20 mL of
the 70 % methanolic extract in a disposable 50-mL grad-
uated centrifuge tube. The tubes were filled with N2,
instantly capped and shaken in a Barnstead Lab-Line sha-
ker at 150 rpm for 2 h at 20 °C after which the mixture was
acidified to pH 2.0 with 6 M HCl. The mixture was then
centrifuged at 5,0009g for 10 min at 4 °C. Twenty milli-
liters of diethyl ether/ethyl acetate mixture was added to a
separation funnel followed by the acidified phenolic extract
and dried under N . The dried residues were then re-dis-
2
solved in 2 mL of 70 % methanol; syringe filtered and kept
at -20 °C until HPLC analysis.
HPLC Analysis
(
the supernatant produced after the centrifugation). The
Canola extracts (before hydrolysis, after hydrolysis and
after decarboxylation) were analyzed by the reversed-phase
HPLC-DAD (Ultimate 3000; Dionex, Sunnyvale, CA,
USA) following the procedure of Khattab et al. [26].
Gradient elution was performed using water/methanol
(90:10) with 1.2 % o-phosphoric acid as solvent A, and
methanol (100 %) with 0.1 % o-phosphoric acid as solvent
separating funnel was capped and shaken well then left to
stand for 5 min to get two separated layers. The organic
upper layer was separated to a round-bottom flask and the
water phase was re-extracted the same way for three times
and the upper phases were combined. The organic phase
was rotary evaporated followed by N flushing. The residue
2
˚
was re-dissolved in 20 mL of 70 % MeOH and stored at
B using C18 column (Synergi 4 lm Fusion-RP 80 A;
-
20 °C until further experiments.
150 9 4.0 mm 4 lm) (Phenomenex, Canada) at 0, 7, 20,
25, 28, 31 and 40 min with 10, 20, 45, 70, 100, 100 and
Hydrolyzing Canola Meal
10 % B. The column was maintained at 25 °C at 0.8 mL/
min flow rate. The chromatograms were acquired at 219,
270, 275 and 330 nm and data were analyzed using
Chromeleon software (Version 6.8). Peaks were identified
by comparing their relative retention times with those of
the authentic standards. The principal constituents; sina-
pine, sinapoyl glucose, and sinapic acid were quantified
based on the calibration curves generated by plotting the
concentration of each compound against the area. All
phenolics including principal components were detected at
330 nm as previously reported [26]. The content of canolol
was expressed as sinapic acid equivalents (SAE) using
detection at 270 nm based on the sinapic acid calibration
curve generated at 270 nm. Total phenolic content (TP,
330 nm) were quantified as sinapic acid equivalents (SAE)
by using all the peaks including unidentified, non-principal
phenolics based on the calibration curve generated by using
sinapic acid.
Canola meal samples (one gram each) were hydrolyzed by
2
0 mL of 4 M NaOH for 4 h at room temperature in the
Barnstead Lab-Line shaker at 150 rpm. The tubes were
flushed with N2 before the extraction. The hydrolyzed
mixture was then centrifuged (5,0009g for 10 min at 4 °C)
and the supernatant was acidified to pH 2.0 and extracted
the same way as mentioned above.
Enzymatic Hydrolysis of Phenolic Compounds
Enzymatic hydrolysis was conducted on canola meal phe-
nolic extracts as well as on the meal itself. Two milliliters
of 70 % methanolic extract was hydrolyzed with 50, 100 or
2
00 lL of ferulic acid esterase (FAE) at 5.5, 6.5 and 7.5 pH
at 37 °C for 4 h. For canola meal, 0.2 g was hydrolyzed
with 50 or 100 lL of FAE. The hydrolysis was investigated
at the same conditions mentioned above.
Statistical Analysis
Decarboxylation of the Hydrolyzed Extracts
The data presented are the means of three repli-
cates ± standard deviations. Data were analyzed using a
one factor analysis of variance (ANOVA) and Tukey mean
separation for multiple comparisons with the Statistical
The decarboxylation of sinapic acid was adapted from the
procedure of Bernini et al. [22] with some modifications.
Briefly, 2 mL of the hydrolyzed extract was put in a 50-mL
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Analysis System (SAS) Program (SAS Institute, Carey,
10.6 ± 0.01 mg/g meal (sinapic acid equivalents; SAEs)
estimated using the HPLC analysis from the total area
under all peaks (Table 1). The results are in accordance
with previous work [26, 29]. After hydrolysis, it ranged
from 8.1 to 11.0 mg/g reaching its highest value after
hydrolyzing for 2 h using 20 mL of 4 M NaOH for 20 mL
of the extract. The amount of released sinapic acid showed
how efficient different hydrolyzing conditions were in
releasing sinapic acid from its esters. Hydrolysis is con-
sidered complete if the value of released sinapic acid is
100 % of the total phenolic content in the original extract.
Under optimum conditions all phenolics were hydrolyzed
to sinapic acid which increased from 0.14 to 10.2 mg/g
representing 94.8 % of the total phenolic content in the
original extract.
NC, USA). Significance was accepted at p B 0.05.
Results and Discussion
Alkaline Hydrolysis
The phenolic extracts were hydrolyzed with NaOH to
release the esterified phenolics. When the pH was below 2,
the released phenolics were available as phenolic acids
instead of ionic forms and could be extracted with diethyl
ether/ethyl acetate mixture [27, 28]. Insoluble-bound
phenolics were not extractable using the normal common
extraction procedures and potentially retained in the meal.
Their proportion is small, but they can be released from
canola by alkaline hydrolysis. As sinapine, the choline
ester of sinapic acid, is the major phenolic compound in
canola meal extracts, the major role of the alkaline treat-
ment is to break the ester linkage in sinapine thereby lib-
erating sinapic acid and choline (Fig. 1). The released
phenolics were extracted with either diethyl ether, or ethyl
acetate, or both. The conversion efficiency and the yield
obtained were higher with alkaline hydrolysis compared to
other methods including enzyme hydrolysis. Alkaline
hydrolysis was conducted on the methanolic extracts as
well as on the meal itself. Sinapine was the predominant
phenolic compound in rapeseed meal, while the amount of
sinapic acid was considerably less. Concerning the meth-
anolic extracts, the total phenolic content was
Using canola meal as a substrate for alkaline hydrolysis,
the released sinapic acid content was lower compared to
using the methanolic extracts (Table 2). The original meal
contained 6.8, 1.1, 0.3 and 8.5 mg/g sinapine, sinapoyl
glucose, sinapic acid and total phenolics, respectively.
After hydrolysis, the released sinapic acid content was
7.1 mg/g (81.0 and 93.9 % of the total phenolics in the
original and hydrolyzed meal, respectively).
Enzymatic Hydrolysis
As NaOH is a strong base, the liberated phenolics may
undergo further degradation and breakdown and a milder
procedure is preferable. Furthermore, if the left-over meal
is intended to be used for the food/feed industries, the
Fig. 1 Alkaline hydrolysis of
sinapine to sinapic acid and
choline
Table 1 Effect of different alkaline hydrolysis conditions on the major phenolic contents in canola extracts
Phenolic content (mg/g)
SP
SG
SA
0.14 ± 0.01
TP (SAE)
a
a
d
c
b
b
a
a
c
b
b
a
Original extract
7.88 ± 0.10
b
0.00
2.78 ± 0.06
b
0.00
10.63 ± 0.21
8.14 ± 0.18
10.49 ± 0.21
10.37 ± 0.20
10.96 ± 0.22
3
1
1
2
.75 mL NaOH for 4 h
0 mL NaOH for 2 h
0 mL NaOH for 4 h
0 mL NaOH for 2 h
7.54 ± 0.12
9.56 ± 0.18
9.75 ± 0.18
10.22 ± 0.20
b
b
0.00
0.00
b
b
0.00
0.00
b
b
0.00
0.00
Results are means ± standard deviations (SD). Values in the same column followed by same superscript letters are not significantly different
p B 0.05)
SP sinapine, SG sinapoyl glucose, SA sinapic acid, TP (SAE) total phenolics as sinapic acid equivalents
(
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Table 2 Effect of alkaline hydrolysis on the major phenolic contents in canola meal
Phenolic content (mg/g)
SP
SG
SA
TP (SAE)
A
a
a
b
a
a
Original extract
6.80 ± 0.10
b
0.00
1.14 ± 0.04
b
0.00
0.27 ± 0.02
7.14 ± 0.10
8.52 ± 0.18
b
Hydrolyzed extract
7.60 ± 0.12
Results are means ± standard deviations (SD). Values in the same column followed by same superscript letters are not significantly different
p B 0.05)
(
SP sinapine, SG sinapoyl glucose, SA sinapic acid, TP (SAE) total phenolics as sinapic acid equivalents estimated from the HPLC analysis
A
Nexera meal
Table 3 Effect of different enzymatic hydrolysis conditions on the major phenolic contents in canola extracts
Phenolic content (mg/g)
SP
SG
SA
TP (SAE)
A
a
a
b
a
a
a
b
b
Original extract
FAE (pH 5.5)
FAE (pH 6.5)
9.01 ± 0.20
b
0.00
0.59 ± 0.06
b
0.00
0.12 ± 0.02
5.47 ± 0.10
5.39 ± 0.10
9.17 ± 0.18
6.56 ± 0.12
6.35 ± 0.12
b
b
0.00
0.00
Results are means ± standard deviations (SD). Values in the same column followed by same superscript letters are not significantly different
p B 0.05)
(
SP sinapine, SG sinapoyl glucose, SA sinapic acid, TP (SAE) total phenolics as sinapic acid equivalents estimated from the HPLC analysis, FAE
ferulic acid esterase
A
Nexera meal
Table 4 Effect of different enzymatic hydrolysis conditions on the major phenolic contents in canola meal
Phenolic content (mg/g)
SP
SG
SA
TP (SAE)
A
a
a
d
b
c
a
a
a
a
a
c
d
c
c
d
b
Original extract
6.80 ± 0.12
b
0.00
1.14 ± 0.04
b
0.00
0.27 ± 0.02
3.85 ± 0.04
2.82 ± 0.04
4.25 ± 0.08
4.36 ± 0.08
4.15 ± 0.08
4.49 ± 0.08
8.52 ± 0.18
4.50 ± 0.08
3.06 ± 0.06
4.46 ± 0.08
4.65 ± 0.08
3.39 ± 0.06
5.21 ± 0.08
FAE (pH 5.5/50 lL)
FAE (pH 5.5/100 lL)
FAE (pH 6.5/50 lL)
FAE (pH 6.5/100 lL)
FAE (pH 6.5/200 lL)
FAE (pH 7.5/50 lL)
b
b
0.00
0.00
b
b
0.00
0.00
b
b
0.00
0.00
b
b
0.00
0.00
b
b
0.00
0.00
Results are means ± standard deviations (SD). Values in the same column followed by same superscript letters are not significantly different
p B 0.05)
(
SP sinapine, SG sinapoyl glucose, SA = sinapic acid, TP (SAE) total phenolics as sinapic acid equivalents estimated from the HPLC analysis,
FAE ferulic acid esterase
A
Nexera meal
presence of alkaline residues will make it unsafe and cause
adverse effects. Consequently, enzymes can be more spe-
cific and environmentally friendly agents. Enzymes have
been successfully used for phenolic hydrolysis in cereals
sinapic acid and choline. Results of the enzymatic hydro-
lysis of canola extracts and canola meal are listed in
Tables 3 and 4. FAE was not as effective in hydrolyzing
sinapic acid esters as NaOH under optimum conditions.
Ideally, it hydrolyzed 58.3 and 49.5 % of the total phen-
olics in the extracts and meals, respectively to free sinapic
acid. The original extract contained 9.01, 0.59, 0.12 and
9.17 mg/g of sinapine (SP), sinapoyl glucose (SG), sinapic
acid (SA) and total phenolics (TP), respectively (Table 3).
[
30] and canola meal [31]. In this study we used ferulic
acid esterase (FAE) which was reported in the earlier lit-
erature [31] to be the most potent for canola hydrolysis.
FAE is a Humicola insolens monocomponent enzyme that
is able to break the ester bond in sinapine to produce
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Fig. 2 Decarboxylation of
sinapic acid to canolol via
microwave irradiation
The use of FAE could totally hydrolyze SP and SG.
However, the TP content was reduced only by 28.46 %
suggesting that the hydrolysis of SP and SG was not
straightforward to SA. In addition to SA, this hydrolysis
yielded other unidentified phenolics which were counted
when calculating the TP content from the total area under
all peaks. Sinapic acid content was increased by 45.58
folds as compared to 73 folds in the alkaline hydrolysis.
Although, the sinapic acid yield after alkaline hydrolysis
was significantly higher compared to the enzymatic
hydrolysis, enzyme hydrolysis remains an efficient and an
environmentally friendly approach. Despite being lower
than that of the alkaline hydrolysis, the conversion rate
using enzyme hydrolysis was substantially high.
allowing quick progress through the hypotheses–experi-
ment–results iterations, resulting in more decision points
per unit of time [32].
In this study the decarboxylation reaction was carried
out under microwave irradiation in the presence of 1,8-
diazabicyclo[5.4.0]undec-7-ene (DBU) as a base reagent
and hydroquinone as a polymerization inhibitor which
were both pre-adsorbed onto the basic aluminum oxide—as
a microwave transparent inorganic solid support—prior to
microwave heating. This environmentally friendly solvent-
free approach in the MAOS has become more prevalent
since it allows the safe use of domestic microwave ovens
and standard open vessel technology while eliminating the
danger of possible explosions [33]. Gedye et al. [34] car-
ried out different types of organic reactions both in sealed
Teflon vessels in a microwave oven and under traditional
reflux conditions and approximate rate enhancement of
between 5 and 1,240 times were recorded. Derivatives
which normally required 2–3 h of preparation were effec-
tively synthesized in 2–3 min in microwave oven.
Thus, if the meal is used only for obtaining canolol,
alkaline hydrolysis is recommended as the yield is higher.
After alkaline hydrolysis, the left-over meal may be used as
a fertilizer. However, if the meal is proposed to be used in
the feed industry and production of protein isolates or
hydrolysates, enzymatic hydrolysis may be a suitable
alternative. Enzymatic hydrolysis is a milder method,
despite the lower yield, to produce a bio residue.
Different experimental conditions were examined to
achieve the highest canolol yield from the hydrolyzed canola
extracts. To decarboxylate the sinapic acid content in 2 mL
of the extract the optimum concentration of the reactants
were found to be 100 lL of DBU, 5 mg of hydroquinone and
Decarboxylation of Sinapic Acid to 4-Vinylsyringol
(
Canolol)
1
g of aluminum oxide (Fig. 2). The DBU reagent was
The use of microwave technology in organic chemistry has
been rapidly increasing since the mid-1980s for the syn-
thesis of new compounds. The most recent applications of
microwave assisted organic synthesis (MAOS) include
N-acylation, alkylation, aromatic and nucleophilic substi-
tution, condensation, cycloaddition, deprotection, protec-
tion, esterification, transesterification, heterocycles,
olefination, and metathesis [16]. Unlike conventional
heating, microwave penetrates inside the material and heat
is generated through direct microwave-material interaction.
Moreover, volumetric heating, reaction rate acceleration,
higher chemical yield, lower energy usage and different
reaction selectivity are advantages microwave heating has
over conventional methods. The short reaction times pro-
vided by microwave synthesis make it ideal for rapid
reaction scouting and optimization of reaction conditions,
chosen as diazabicycloalkenes are efficient bases for the
decarboxylation of unsaturated acids [35]. Furthermore,
2
4
DBU has a pKa = 12 and it was reported that the decar-
boxylation of cinnamic acids proceeded in better yields with
bases having higher basicity (pKa [ 7) [23].
The effect of microwave power level and heating times
is shown in Table 5. The microwave heating was applied at
different power levels; power 1 (100 W) to power 7
(700 W) for 1–25 min. However, the results shown in
Table 5 are limited to power 3 (12–15 min), power 4 (4
and 8 min) and power 5 (5 and 7 min) in which the
decarboxylation could be achieved and the canolol could
be detected. Decarboxylation did not occur at both power 1
and power 2. Furthermore, after 15 min at power 3, 8 min
at power 4, and 7 min at power 7 of microwave power, the
samples were visibly burnt and no further extraction could
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95
Table 5 Decarboxylation of the hydrolyzed canola extracts under
microwave irradiation as affected by different experimental
conditions
associated costs especially for industrial use. The major
advantage of the present work is the use of the under-
utilized meal as the substrate rather than the oil which is
the valuable product in the canola oil industry. More work
is needed to enhance the conversion rate, extractability and
final yield of canolol based on modifying the experimental
conditions given our first novel findings reported here. The
HPLC chromatograms of the original, hydrolyzed and
decarboxylated extracts are shown in Fig. 3. Around
Canolol occurrence
Content (mg/g)
% of sinapic acid
b
b
Power 3/12 min
Power 3/13 min
Power 3/15 min
Power 4/4 min
Power 4/8 min
Power 5/5 min
Power 5/6 min
Power 5/7 min
3.28 ± 0.04
43.50
a
a
4.16 ± 0.08
58.26
d
d
2.18 ± 0.04
28.91
b
b
3.33 ± 0.04
44.16
96.1 % of the total phenolics in canola meal was converted
c
c
2.86 ± 0.04
37.93
to sinapic acid of which 58.3 % was decarboxylated to
canolol. This yield of 4.2 g of canolol for each kg of canola
meal represents a significantly higher yield of canolol
compared to other reported values [22, 38]. Niu et al. [39]
found that the concentration of canolol in rapeseed
increased nearly six times after 7 min of microwave
toasting accompanied by a decrease in both sinapine and
sinapic acid suggesting that these compounds are chemi-
cally degraded by microwave heating. Herstad and Benn-
eche [21] used the decarboxylation in the synthesis of
4-alkyl-, 4-alkenyl- and 4-acylpyrimidines through both
pure thermally and microwave-induction. They concluded
that the time needed to get complete decarboxylation was
greatly reduced using the microwave irradiation. The pure
thermal decarboxylation at 111 °C took from 48 to 72 h
compared to 20 min only under microwave irradiation.
Similarly, microwave-enhanced decarboxylation of aro-
matic carboxylic acids was found to occur very rapidly and
complete within 4 min at 300 W microwave power [18].
a
a
3.99 ± 0.06
55.88
c
c
2.86 ± 0.04
37.93
c
c
2.92 ± 0.04
40.90
Results are means ± standard deviations (SD). Values in the same
column followed by same superscript letters are not significantly
different (p B 0.05)
be done. Canolol was identified based on its absorbance
properties according to the literature. Wijesundera et al.
[
36] found that canolol had a UV spectrum with maxima at
20 and 270 nm which agree with our findings. Similarly,
2
Koski et al. [1] reported that canolol has an absorption
maximum occurring at 275 nm while our study indicated
an absorbance maximum at 270 nm.
It was concluded that using the microwave at power 3 of
microwave power for 13 min achieved the highest con-
version rate (58.4 % of sinapic acid) and canolol yield
(
4.2 mg/g of canola meal). This canolol yield (4.2 g from
each kilogram of canola meal) is outstanding from an
economical point of view. Lower power levels were more
efficient in decarboxylation which suggests that the con-
version of sinapic acid to canolol is more effective under
these conditions.
Conclusions and Future Perspectives
The present work discloses a simple technique for obtain-
ing canolol from canola meal using microwave irradiation
as an environmentally friendly process for the first time
according to the best of our knowledge. The optimum
condition for microwave treatment is power 3 (300 W of
microwave power) for a total treatment of 13 min.
Hydrolysis is best acheived using FAE. Parts of this study
were reported in the Annual Meeting of the American Oil
Chemist’s Society in 2009 and 2010. Further studies to
scale up and commercialize this technology are essential to
pursue value addition to both canola oil and canola meal.
The canolol obtained could be added back to vegetable
oils, oil-containing systems, snack foods, and other sup-
plementary foods, for better nutritional/nutraceutical qual-
ity. Utilizing canola meal could provide additional
economic benefits to canola growers and crushers. The
lower chemical and solvent consumption involved in the
extraction, hydrolysis and decarboxylation makes it a more
environmentally friendly process. Results of this research
may create new integrated techniques for processing canola
Since the early days of microwave synthesis, the
observed rate-accelerations and sometimes altered product
distributions compared to conventional heating led to
speculation on the existence of so-called ‘‘specific’’ or
‘‘non-thermal’’ microwave effects [37]. Such effects were
claimed when the outcome of a synthesis performed under
microwave conditions was different from the convention-
ally heated counterpart at the same apparent temperature.
Scientists agree that the reason for the observed rate
enhancements is a purely thermal/kinetic effect, that is, a
consequence of the high reaction temperatures that can
rapidly be attained when irradiating polar materials in a
microwave field, although clearly effects that are caused by
the uniqueness of the microwave dielectric heating mech-
anism (specific microwave effects) must also be considered
[
32].
Using the lower power level (300 W) of microwave
power might prove to be an asset in energy saving and
1
23

9
6
J Am Oil Chem Soc (2014) 91:89–97
Fig. 3 HPLC-DAD
chromatograms of canola
extracts before hydrolysis (a),
after hydrolysis (b) and after
decarboxylation (c)
seed and canola meal that might include microwave
treatments and hydrolysis for the extraction of this
nutraceutical.
Conflict of interest The authors have declared no conflict of
interest.
Acknowledgments Dr. Amy Logan, CSIRO, Food and Nutrition
Sciences Australia and Dr Bertrand Mathaus, Max Rubner-Institute
Germany are kindly acknowledged for the authentic canolol standard
synthesized in their laboratories. The authors would like to
acknowledge the technical assistance of Yougui Chen and Ayyapan
Appukutan in this manuscript. This work was funded by the Natural
Sciences and Engineering Research Council of Canada (NSERC), the
Canola Council of Canada, and Syngenta Crop Protection Inc, Can-
ada. The Post-doctoral fellowship of Dr. R. Khattab at the University
of Manitoba is sincerely acknowledged.
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