Oxidation of Procyanidins with Various Degrees of Condensation: Influence on the Color-Deepening Phenomenon

Article abstract of DOI:10.1021/acs.jafc.9b02085

Color deepening in red rice is a known phenomenon that occurs during postharvest storage. As procyanidins potentially causes this color deepening, we previously subjected synthetic procyanidin B3, a model procyanidin, to chemical oxidation and determined the structures of the products. To further elucidate the effects of various degrees of polymerization on color deepening, in this study we oxidized synthetic procyanidin C2. The intensity of the UV-vis absorption between 300 and 500 nm in the spectrum of the reaction mixture increased with increasing oxidation time. This absorption following the oxidation of procyanidin C2 was more intense than that of procyanidin B3. HPLC-ESI-MS of the oxidized reaction mixture revealed several new peaks that suggested the formation of products with new intramolecular rather than intermolecular bonds. The major product was structurally identified by comparison with the UPLC-ESI-MS/MS data for a synthetic procyanidin C2 oxide, independently prepared by condensing procyanidin B3 oxide and taxifolin.

Full text of DOI:10.1021/acs.jafc.9b02085

Subscriber access provided by UNIV OF LOUISIANA
Food and Beverage Chemistry/Biochemistry
Oxidation of procyanidins with various degrees of
condensation: influence on the color deepening phenomenon
Yoshihide Hibi, and Emiko Yanase
J. Agric. Food Chem., Just Accepted Manuscript • DOI: 10.1021/acs.jafc.9b02085 • Publication Date (Web): 16 Apr 2019
Downloaded from http://pubs.acs.org on April 16, 2019
Just Accepted
“Just Accepted” manuscripts have been peer-reviewed and accepted for publication. They are posted
online prior to technical editing, formatting for publication and author proofing. The American Chemical
Society provides “Just Accepted” as a service to the research community to expedite the dissemination
of scientific material as soon as possible after acceptance. “Just Accepted” manuscripts appear in
full in PDF format accompanied by an HTML abstract. “Just Accepted” manuscripts have been fully
peer reviewed, but should not be considered the official version of record. They are citable by the
Digital Object Identifier (DOI®). “Just Accepted” is an optional service offered to authors. Therefore,
the “Just Accepted” Web site may not include all articles that will be published in the journal. After
a manuscript is technically edited and formatted, it will be removed from the “Just Accepted” Web
site and published as an ASAP article. Note that technical editing may introduce minor changes
to the manuscript text and/or graphics which could affect content, and all legal disclaimers and
ethical guidelines that apply to the journal pertain. ACS cannot be held responsible for errors or
consequences arising from the use of information contained in these “Just Accepted” manuscripts.
is published by the American Chemical Society. 1155 Sixteenth Street N.W.,
Washington, DC 20036
Published by American Chemical Society. Copyright © American Chemical Society.
However, no copyright claim is made to original U.S. Government works, or works
produced by employees of any Commonwealth realm Crown government in the
course of their duties.

Page 1 of 31
Journal of Agricultural and Food Chemistry
Oxidation of procyanidins with various degrees of condensation: influence on the
color deepening phenomenon
Yoshihide Hibi, Emiko Yanase*
Faculty of Applied Biological Sciences, Gifu University, 1-1 Yanagido, Gifu 501-1193,
Japan
*Corresponding author:
Emiko Yanase
TEL/FAX +08-58-293-2914
E-mail e-yanase@gifu-u.ac.jp
1
ACS Paragon Plus Environment

Journal of Agricultural and Food Chemistry
Page 2 of 31
1
ABSTRACT
2
3
Color deepening in red rice is a known phenomenon that occurs during post-harvest
storage. As procyanidins potentially causes this color deepening, we previously
subjected synthetic procyanidin B3, a model procyanidin, to chemical oxidation and
determined the structures of the products. To further elucidate the effects of various
degrees of polymerization on color deepening, in this study we oxidized synthetic
procyanidin C2. The intensity of the UV-Vis absorption between 300 and 500 nm in the
spectrum of the reaction mixture increased with increasing oxidation time. This
absorption following the oxidation of procyanidin C2 was more intense than that of
procyanidin B3. HPLC-ESI-MS of the oxidized reaction mixture revealed several new
peaks that suggested the formation of products with new intramolecular rather than
intermolecular bonds. The major product was structurally identified by comparison with
the UPLC-ESI-MS/MS data for a synthetic procyanidin C2 oxide, independently
prepared by condensing procyanidin B3 oxide and taxifolin.
4
5
6
7
8
9
10
11
12
13
14
15
16
KEYWORDS: Color deepening, polymerization degree, catechin, procyanidin, HPLC-
ESI-MS
17
2
ACS Paragon Plus Environment

Page 3 of 31
Journal of Agricultural and Food Chemistry
18
INTRODUCTION
19
20
21
22
23
24
25
26
27
28
29
30
31
32
33
34
35
Many varieties of rice (Oryza sativa) are cultivated around the world, and
those with pigments are commonly known as colored rice varieties. Almost all pigments
are contained within the bran layers, and colored rice is classified as red or black based
on its color. The main pigment in black rice is anthocyanin. Although the pigment of
red rice has been presumed to be an anthocyanin, it is not observed in its extract; hence,
the major pigment has not yet been clearly identified.
Procyanidin is present in the bran layers of red rice. It is condensed tannin
composed of flavan-3-ol units, such as catechin, epicatechin, gallocatechin, and
epigallocatechin, which are widely distributed in plants, particularly in fruits. They are
classified into two types (A and B) according to their bonding systems. These bioactive
constituents exhibit beneficial biological properties, including strong anti-oxidative,
anti-inflammatory, and anti-cancer activities.1-4 The color of red rice intensifies during
ripening after flowering, and becomes darker in the post-harvest storage period. This
phenomenon is the result of chemical changes undergone by the preexisting components
of the red rice; however, the detailed mechanism associated with these changes is not
clear. We presume that color-deepening is related to the oxidation of the procyanidins in
the red rice and previously studied the oxidation of procyanidin B3, a model
3
ACS Paragon Plus Environment

Journal of Agricultural and Food Chemistry
Page 4 of 31
36
37
38
39
40
41
42
43
44
45
46
47
48
49
50
51
52
procyanidin, and isolated several novel oxidized products with spiro-type skeletons that
contain new internal bonds.5,6
The procyanidins present in red rice are generally assumed to have polymeric
structures that range from 2-mers to 14-mers, depending on the cultivar. Therefore, it is
difficult to explain the color-deepening phenomenon from the knowledge gained
through the oxidation of the model procyanidin B3 alone. In order to elucidate the
effects of the degree of catechin polymerization on the oxidation reaction and the color
change, in this study we synthesized procyanidin C2, as a model procyanidin, and
subjected it to chemical oxidation.
MATERIALS AND METHODS
Chemicals (+)-Catechin (1) was purchased from Sigma-Aldrich Co. (St.
Louis, MO, USA) and (+)-taxifolin (2) was isolated and purified from larch.7 Copper(II)
chloride dihydrate (CuCl2·2H2O), 2-mercaptoethanol, and all solvents were purchased
from Nacalai Tesque Co. (Kyoto, Japan).
Instrumentation Column chromatography was performed on Diaion
HP20SS resin (Mitsubishi Chemical Co., Tokyo, Japan). HPLC was performed using a
4
ACS Paragon Plus Environment

Page 5 of 31
Journal of Agricultural and Food Chemistry
53
JASCO PU2089 intelligent pump equipped with a JASCO MD-2010 detector and a
JASCO CO-2065 column oven (Tokyo, Japan).
54
55
56
57
58
59
60
61
62
63
64
65
66
67
68
69
70
Preparation of the red rice procyanidin fraction. Red rice (20 g) was
soaked for 6 h in water (70 mL) at room temperature. After filtration, the solid was
extracted for 1 day with acetone (50 mL) at room temperature. After removal of the
solid by filtration, the acetone was removed by rotary evaporation and the extract was
subjected to column chromatography. To obtain the procyanidin rich fraction,
fractionation was performed using a Diaion HP20SS column (20 mm I.D. × 200 mm).
The extract reside was dissolved in water and then adsorbed on the column. The sample
was eluted sequentially with 0%, 20%, and 50% aqueous MeOH. The fraction that
eluted with 50% aqueous MeOH was evaporated under reduced pressure to leave a
residue (100.3 mg). The sample was subjected to HPLC with a HILIC column (Inertsil
Amide 4.6 mm I.D. × 250 mm). The column temperature was maintained at 35°C. The
mobile phase consisted of H2O (A) and MeCN (B) at a flow rate of 1.0 mL/min. The
following linear gradient was used for analysis: 85% to 65% B over 15 min.
Synthesis of procyanidin C2 (4). Procyanidin B3 was prepared from
catechin and taxifolin following a literature procedure.6,8 (+)-Taxifolin (2, 50 mg) was
dissolved in EtOH (1.0 mL). Sodium borohydride (75 mg) was added under argon and
5
ACS Paragon Plus Environment

Journal of Agricultural and Food Chemistry
Page 6 of 31
71
72
73
74
75
76
77
78
79
80
81
82
83
84
85
86
87
88
stirred 30min. Water (10 mL) was added to the mixture to decompose the excess
sodium borohydride, and the pH was adjusted to below 5.0 with acetic acid.
Procyanidin B3 (3, 200 mg) in 50% aqueous EtOH was then added to the mixture at
40°C and stirred for 3 h.8 The reaction mixture was extracted with ethyl acetate and
concentrated. The ethyl acetate extract was purified by HPLC using a HILIC column
(Inertsil Amide 10 mm I.D. × 250 mm, 5 µm (GL Sciences Inc., Tokyo, Japan)). The
mobile phase consisted of 90% MeCN in H2O, at a flow rate of 5.0 mL/min, and an
injection volume of 50 L. Procyanidin C2 (4, 41 mg, purity > 95% by HPLC) was
obtained in this manner. The chemical structure of 4 was confirmed by comparing its
NMR and MS data with those in the literature.8 The structures of catechin (1), taxifolin
(2), procyanidin B3 (3), procyanidin C2 (4), and oxidative procyanidin B3 (5) are
shown in Fig. 1.
Oxidation of 4 and the procyanidin fraction of red rice. An aqueous
solution of 4 (2.0 mg) and CuCl2·2H2O (6.0 mg) was vigorously stirred at room
temperature. An aliquot of the reaction mixture at each reaction period (0.5, 1, 2, and 4
h) was subjected to Diaion HP20SS column chromatography. After washing with water
to remove CuCl2·2H2O, the column was eluted with MeOH. Each MeOH fraction was
concentrated and the residue (2.1 mg) examined by reverse-phase HPLC (NB-ODS-9
6
ACS Paragon Plus Environment

Page 7 of 31
Journal of Agricultural and Food Chemistry
89
4.6 mm I.D. × 250 mm). The column temperature was maintained at 35°C. The mobile
phase consisted of 10% MeCN / 0.5% HCOOH in H2O, at a flow rate of 1.0 mL/min.
An injection volume of 10 L was used.
90
91
92
UV-Vis spectroscopy. The mixtures of oxidation products were dissolved in
MeOH to a concentration of 0.10 g/L. UV/Vis spectra were recorded on a Perkin Elmer
Lambda 950 UV/VIS/NIR spectrophotometer (Kanagawa, Japan). The spectrum of each
sample was acquired in the 200–600 nm wavelength range at a spectral resolution of 1
nm. The spectra were recorded against the corresponding solvent as the baseline.
Thiolysis of the mixture obtained by the oxidation of 4. Thiolysis was
conducted according to the procedure reported by Tanaka et al., with slight
modifications.9 The oxidation-product mixture containing the trimer (1 mg) was
dissolved in EtOH (1.6 µL), and 0.2 N aqueous HCl (1.6 µL) and 2-mercaptoethanol
(2.0 µL) were added; the mixture was then stirred at room temperature for 24 h, after
which the EtOH was removed under reduced pressure. The aqueous solution was
separated by Diaion HP20SS column chromatography and monitored by HPLC with a
HILIC column (Inertsil Amide 4.6 mm I.D. × 250 mm). The column temperature was
maintained at 35°C. The mobile phase consisted of 90% MeCN in H2O at a flow rate of
1.0 mL/min; the injection volume was 10 L.
93
94
95
96
97
98
99
100
101
102
103
104
105
106
7
ACS Paragon Plus Environment

Journal of Agricultural and Food Chemistry
Page 8 of 31
107
108
109
110
111
112
113
114
115
116
117
118
119
120
121
122
123
124
UPLC-ESI-MS. Ultra-high-performance liquid chromatography–time-of-
flight mass spectrometry (UPLC-TOF-MS) was performed using a Waters Xevo G2
QTof (Waters, USA) mass spectrometer, equipped with a C18 analytical column
(ACQUITY UPLC BEH C18 2.1 mm I.D. × 100 mm; Waters, USA) and a HILIC
column (Inert Sustain Amide HP 3 µm, 2.1 mm I.D. × 150 mm; GL Science, Japan).
The mobile phase consisted of 1.0% HCOOH in H2O (A) and MeCN (B), with a flow
rate of 0.2 mL/min and an injection volume of 1.0 µL used. The column temperature
was maintained at 35°C. The mobile phase for analysis of the trimer oxidation products
was 10% B for 20 min for the C18 analytical column, or 90% B for 30 min for the
HILIC column. The following linear gradient was used to analyze the thiolysis reaction
mixture: 90% to 85% B for 15 min using the HILIC column. The ESI interface was
operated in negative mode. MS parameters for analysis were: capillary potential, 2.5
kV; sampling cone, 30 V for in-source collision-induced dissociation (CID); desolvation
temperature, 500°C; and source temperature, 150°C. The mass spectrometer was
operated in MS/MS mode with a collision cell energy of 25 V for the oxide of 4.
RESULTS AND DISCUSSION
UV/Vis Spectra Measurements. The UV-Vis spectra of the oxidation
8
ACS Paragon Plus Environment

Page 9 of 31
Journal of Agricultural and Food Chemistry
125
mixtures of both 3 and 4 were acquired in order to study the relationship between
chemical structure and color. The absorption band between 300 and 500 nm for each
sample was more intense following oxidation (Figs. 2-A and B). The relationship
between the absorbance at 330 nm of each sample and time is shown in Fig. 2-C. The
increase in the absorbance at 330 nm following oxidation was four times greater for 4
than that it was for 3. In addition, although moderate changes in absorption were
observed for the reaction mixture of 3 beyond ~1 h, 4 exhibited an absorption that
continued to increase even after 2 h.
126
127
128
129
130
131
132
133
134
135
136
137
138
139
140
141
142
The procyanidin fraction of the red rice extract was next subjected to oxidation
for 2 h and examined by UV-Vis spectroscopy (Fig. S2). 2-Mer-to-8-mer procyanidins
were observed in this sample by HPLC (Fig. S3), with the intensity of the absorption band
in the 300–500 nm range observed to increase, in addition to the absorption maximum
near 280 nm. This trend is similar to that observed for the oxidations of 3 and 4; therefore,
we assume that the oxidation systems in this study, namely 3 and 4, are good models for
assessing the effects of procyanidin oxidation on color change. Moreover, we assume that
the degree of procyanidin polymerization significantly influences the color-deepening
phenomenon.
HPLC. The oxidized mixtures of 3 and 4 were subjected to HPLC. The
9
ACS Paragon Plus Environment

Journal of Agricultural and Food Chemistry
Page 10 of 31
143
144
145
146
147
148
149
150
151
152
153
154
155
156
157
158
159
160
time-dependent decreases in the contents of 3 and 4 under oxidative conditions are
shown in Figs. S4 and S5. The contents of both model compounds 3 and 4 decrease
sharply after commencement of oxidation. Trimer 4 was observed to react faster than
dimer 3, and required only 1 h to disappear completely from the mixture (Fig. 3).
The major oxidation products of 3 appear to be spiro-type compounds, as
represented by compound 5, among others, (Figs. 1 and S1) and have been reported
previously.6 In contrast, a complicated assortment of peaks (a–k) was observed in the
HPLC trace following the oxidation of 4; reactions continued following the
disappearance of compound 4, as evidenced by changes in the intensities of these peaks
(Figs. S4 and S5). Several peaks (e.g., peak j) were observed to increase in intensity,
whereas others (e.g., peak k) decreased. The UV-Vis spectra of the fractions
corresponding to peaks a–k were acquired using an HPLC-photodiode array (PDA)
detector (Fig. S6). Although the maximum absorption wavelengths of these fractions
were similar to that of compound 4, the ratios of the absorption intensities in the 300–
400 nm region to the that of maximum absorption peak were higher, suggesting that
these peaks correspond to oxidation products involved in the color deepening process.
UPLC-MS. The mixture resulting from the oxidation from 4 was subjected to
UPLC-MS. Molecular ion peaks were observed at m/z 861 or 863 [M-H]−, although a
10
ACS Paragon Plus Environment

Page 11 of 31
Journal of Agricultural and Food Chemistry
161
162
163
164
165
166
167
168
169
170
171
172
173
174
175
176
177
178
variety of new peaks were observed by HPLC (Figs. 4, S7, and S8). These data suggest
that the oxidation products contain new intramolecular bonds rather than intermolecular
bonds, which is similar to the trend observed for the oxidation of 3. UPLC-MS traces
for each reaction period are shown in Figs. 4 and S7. Many complex overlapping peaks
are observed in the total ion chromatograms (TICs) and the chromatograms at 280 nm.
The reconstructed ion chromatograms for m/z 863 and 861 facilitated the resolution of
multiple well-separated peaks with the same mass number that correspond to positional-
or stereo-isomers. De Taeye et al. reported that Procyanidin is chemically modified by
food processes;10, 11 in their research, procyanidin B2 and C1, which are the dimer and
trimer of epicatechin, were epimerized by heating in a model aqueous system, to
generate dehydrodicatechin A; however, this was not observed in our experiment. It is
well known that catechin rapidly epimerizes at the 2-position at high temperatures in an
aqueous medium.12 In addition, De Taeye et al.10 concluded that dehydrodicatechin A13-
16 is formed from procyanidin B2 only at high temperatures because cleavage of the
interflavan linkage is required in order to generate the catechin monomer. Because our
oxidation experiments were performed at room temperature, we believe that
procyanidin C2 does not epimerize to generate a dehydrodicatechin A-like component.
We conclude that many peaks are observed because several potential positions for
11
ACS Paragon Plus Environment

Journal of Agricultural and Food Chemistry
Page 12 of 31
179
180
181
182
183
184
185
186
187
188
189
190
191
192
193
194
195
196
reaction exist in the trimer molecule. In addition, we assume that the trimer oxide
epimerizes during the reaction. In fact, three isomers of the dimer oxide 5 were
observed in the reaction solution following the oxidation of 3 (Fig. S1).6 Many large
peaks are observed in the initial stages of the reconstructed m/z 863 ion chromatogram,
h
which is 2 Da smaller than the m/z value of 4; these peaks decrease in size with
increasing reaction time. In addition, the intensities of the peaks corresponding to m/z
861, which is 4 Da smaller than that of 4, increase as the reaction progresses. These
results suggest that the oxidation process involves multiple steps rather than a single
step.
UPLC-MS/MS. Almost all of the peaks observed by UPLC-MS following
the oxidation of 4 were observed to have m/z 861 [M-H] − or 863 [M-H] – peaks (Fig.
S7); hence, each peak was subjected to MS/MS analysis (Fig. S9). MS/MS of the m/z
863 peak, with a retention time of 15.9 min, produced fragment ion peaks at m/z 287,
407, 449, 455, 575, 693, 711, and 737. The fragments at m/z 711 and 737 are assumed
to be formed by the heterocyclic ring fission and retro Diels–Alder cleavage of the
catechin unit of the trimer oxide,17, 18 while quinone-methide cleavage of the interflavan
bond generates the fragment ion at m/z 287. The fragment ion at m/z 693 is produced by
the loss of molecular water from the ion corresponding to m/z 711; these peaks are also
12
ACS Paragon Plus Environment

Page 13 of 31
Journal of Agricultural and Food Chemistry
197
198
199
200
201
202
203
204
205
206
207
208
209
210
211
212
213
214
observed by the MS/MS of 4 prior to oxidation. On the other hand, m/z 575, 455, and
449 correspond to product peaks, and are newly observed in the MS/MS spectrum of the
m/z 863 ion. The fragment ion peak at m/z 575 corresponds to the product of catechin-
dimer oxidation. Some of the fragment ion peaks (m/z 125, 167, 287, 407, and 449) are
also observed in the MS/MS spectrum of compound 5. These results suggest that
products exhibiting peaks at m/z 863 have spiro-type skeletons, such as 5, as partial
structures in their molecular frameworks. The possibility exists that one of the m/z 863-
peak products is A-type procyanidin formed through an additional C2-O-C7 ether bond.
The transformations of B-type procyanidins to the A-type under alkaline/H2O2,19 DPPH
(2,2-diphenyl-1-picrylhydrazyl)-radical,20 and enzymatic conditions,21 have been
previously reported. Although these oxidation reactions are similar to those used in our
experiments, we believe that the A-type structure is not formed in our experiments due
to steric factors. The C4 position of the catechin trimer prepared from (+)-catechin
mainly exists in the  form. Procyanidin C2, which is -configured at the C4 position,
is produced as the major product through condensation reactions from (+)-catechin. In
practice, the / ratio at the C4 position was observed to be 7/3 by NMR spectroscopy.
Condensed (+)-catechin in the  form is incapable of forming the C2-O-C7 ether bond;
therefore, we conclude that A-type structures are hardly produced in this study.
13
ACS Paragon Plus Environment

Journal of Agricultural and Food Chemistry
Page 14 of 31
215
216
217
218
219
220
221
222
223
224
225
226
227
228
229
230
231
232
MS/MS of the m/z 861 [M-H]− peak, with a retention time 8.2 min, reveals
peaks at m/z 407, 453, and 709, which are related to the fragment peaks observed in the
MS/MS spectrum of the m/z 863 ion. Although the presence of two spiro skeletons in
the molecule is assumed on the basis of the peak at m/z 861, the expected fragment
peaks corresponding to the catechin unit (m/z 575) and the remaining component (m/z
287) were not observed, suggesting that different types of intramolecular bonds were
generated in this case.
Thiolysis degradation reactions. Thiolysis is a known method for the
analysis procyanidins. In this method, procyanidins are fragmented into their constituent
units in the presence of a thiol, such as thiophenol or 2-mercaptoethanol. Although
thiolysis can release the extension unit-thiol adducts and the terminal units from
procyanidin B-type polymers, which contain C4–C8 or C4–C6 bonds, this method does
not degrade procyanidins possessing other types of bond. We assume that the C4–C8/6
bond in 4 are cleaved by this reaction; however, the new bonds produced by the
oxidation of 4 cannot be cleaved; consequently, any novel unit is retained, which
facilitates the elucidation of the novel bonding pattern produced through oxidation.
Hence, compound 4 and its oxidation mixtures were treated with 2-mercaptoethanol and
the reaction solution was subjected to HPLC. Due to thiolytic degradation, the
14
ACS Paragon Plus Environment

Page 15 of 31
Journal of Agricultural and Food Chemistry
233
234
235
236
237
238
239
240
241
242
243
244
245
246
247
248
249
250
originally observed peaks were replaced by new peaks, with negative UPLC-ESI-MS of
the generated peaks exhibiting pseudo-molecular-ion peaks at m/z 575, 651, and 653
[M-H]-.
The thiolysis degradation products were next subjected to UPLC-ESI-MS/MS
(Fig. 5). The precursor ion peak at m/z 575 corresponds to 5, which is the oxidation
product of procyanidin B3. The m/z 575 peak exhibited major fragment ions at m/z 449
and 287, which are in agreement with the fragment pattern of 5. Fragment ions that are
78 Da less than the precursor ions were observed at m/z 651 and 653. The loss of 78 Da
suggests the presence of the mercaptoethanol residue in these molecules, and these
compounds are presumed to be derived from the extended units of 4 or its oxidized
product, rather than the terminal unit.6 The m/z 651 peak exhibited major fragment ions
at m/z 573, 449, and 285; this fragment pattern is similar (each 2 Da less) to that of 5.
Based on this result, the peak with the m/z 651 precursor ion is assigned to be the
mercaptoethanol adduct of 5.
Fragment ions from the precursor ion peak at m/z 653 were observed at m/z
575, 449, and 287, which is same fragment pattern observed for 5. The spiro ring
system formed through the oxidation reaction retains the C4-C6/8 bond; hence we
propose that the precursor ion peak at m/z 653 is produced by the ring-opening reaction
15
ACS Paragon Plus Environment

Journal of Agricultural and Food Chemistry
Page 16 of 31
251
252
253
254
255
256
257
258
259
260
261
262
263
264
265
266
267
268
of the spiro ring system following the formation of the thiol adduct (Fig. S10). These
results suggested that the product formed by the oxidation of procyanidin C2 contains a
spiro ring system as a main partial structure.
Determining the structure of peak k. As detailed above, the UPLC-MS
spectrum of the oxidation mixture of 4 exhibited peaks that increased and decreased in
intensity depending on the duration of oxidation (Fig. 3). This is especially evident for
peak k, with a retention time of 8 min, which was most intense at 0.5 h and decreased in
intensity thereafter. The molecular weight corresponding to peak k was determined to
be 864 Da from the mass chromatogram (2 Da less than that of 4), suggesting that
intramolecular bonds are formed during oxidation, as was observed for 5 (the oxidation
product of 3). Considering the time required to generate peak k, we assume that the
reaction and bond formation occur in a manner similar to that of compound 5. In other
words, the chemical structure corresponding to peak k is presumed to contain the same
spiro skeleton as in compound 5. Therefore, we postulate that peak k corresponds to
structure 6, which is obtained through the condensation of the dimer oxide 5 and 2 (Fig.
6). Hence, we separately condensed 5 with 2 and subjected the resulting mixture to
HPLC. The retention time of 6 matched that of peak k. UPLC-MS of this synthesized
trimer revealed a pseudo-molecular ion peak at m/z 863 for [M−H]−, which is the same
16
ACS Paragon Plus Environment

Page 17 of 31
Journal of Agricultural and Food Chemistry
269
270
271
272
273
274
275
276
277
278
279
280
281
282
283
284
285
286
as that observed for peak k. In addition, the MS/MS data are in good agreement with
those of peak k (Fig. S11). On the basis of these result, 6 was confirmed as the chemical
structure corresponding to peak k.
As described above, although various peaks were observed in the oxidation
mixture in this LC-MS study, their molecular ion peaks were observed at either m/z 861
[M-H]− or 863 [M-H]−, some of which are possibly stereo- or positional isomers of 6. In
particular, spiro-type compounds, such as 5, and their derivatives were identified in red
rice extracts through thiolysis reactions and HPLC-MS/MS multiple reaction monitoring
(MRM) analyses in our previous study.6 These results reveal that the color deepening in
red rice proceeds through the oxidative formation of intramolecular bonds in procyanidins.
In contrast, the UV absorptions of compound 6 and other oxidants are insufficient to
explanation the color deepening of red rice (Fig. 5). We believe that the internal bonding
system formed in procyanidin is one of factors that causes color deepening, but additional
other mechanisms might also operate. Further studies are needed in order to clarify and
fully understand this phenomenon.
The term “oxidation” is often used to describe the degradation of food quality. It
is possible that the UV absorption changes and oxidation products clarified in this study
serve as the basis for evaluating the freshness of red rice. In addition, this study will aid
17
ACS Paragon Plus Environment

Journal of Agricultural and Food Chemistry
Page 18 of 31
287
288
289
290
291
292
293
294
295
296
297
298
299
300
301
302
303
304
in the further elucidation of the color-deepening phenomenon of red rice as well as
various plants containing procyanidins.
ABBREVIATIONS
ESI-MS, electrospray ionization mass spectroscopy; HPLC, high-performance liquid
chromatography; NMR, nuclear magnetic resonance; TIC, total ion chromatogram; UPLC, ultra-
high-performance liquid chromatography; MRM, multiple reaction monitoring
ACKNOWLEDGMENTS
We thank Dr. Yoshihiro Miwa (Professor Emeritus at Gifu University) for providing the red rice
samples, and Dr. Tohru Mitsunaga (Professor at Gifu University) for technical assistance in the
preparation of the taxifolin sample.
Supporting Information: Dimer oxidation product structures; additional UV, HPLC, LC/MS,
and MS/MS data for mixtures of oxidation products. Supporting Information is available free of
charge on the ACS Publications website.
REFERENCES
(1) Chen, M. H.; Choi, S. H.; Kozukue, N.; Kim, H. J.; Friedman, M. Growth-inhibitory
effects of pigmented rice bran extracts and three red bran fractions against human
cancer cells: Relationships with composition and antioxidative activities. J. Agric.
Food Chem. 2012, 60, 9151-9161.
18
ACS Paragon Plus Environment

Page 19 of 31
Journal of Agricultural and Food Chemistry
305
306
307
308
309
310
311
312
313
314
315
316
317
318
319
320
(2) Itani, T.; Tatemoto, H.; Okamoto, M.; Fujii, K.; Muto, N. A comparative study on
antioxidative activity and polyphenol content of colored kernel rice. J. Jpn. Soc.
Food Sci. 2002, 49, 540-543.
(3) Min, B.; McClung, AM.; Chen, MH. Phytochemicals and antioxidant capacities in
rice brans of different color. J. Food Sci. 2011, 76, C117-C126.
(4) Oki, T.; Masuda, M.; Kobayashi, M.; Nishiba, Y.; Furuta, S.; Suda, I.; Sato, T.
Polymeric procyanidins as radical-scavenging components in red-hulled rice. J.
Agric. Food Chem. 2002, 50, 7524-7529.
(5) Hayashi, S.; Yanase, E. A study on the color deepening in red rice during storage.
Food Chem. 2016, 199, 457-462.
(6) Hayashi, S.; Kohei, N.; Yanase, E. Investigation of color-deepening phenomenon in
catechin - (4 → 8) - dimer as a proanthocyanidin model and structural
determination of its derivatives by oxidation. Food Chem. 2018, 239, 1126-1133.
(7) Mitsunaga, T.; Abe, I.; Kontani, M.; Ono, H.; Tanaka, T. Inhibitory effects of bark
proanthocyanidins on the activities of glucosyltransferases of Streptococcus
sobrinus. J. Wood. Chem. Tech. 1997, 17, 323-340.
19
ACS Paragon Plus Environment

Journal of Agricultural and Food Chemistry
Page 20 of 31
321
322
323
324
325
326
327
328
329
330
331
332
333
334
335
336
337
(8) Delcour, J. A.; Ferreira, D.; Roux, D. G. Synthesis of condensed tannins. Part 9. The
condensation sequence of leucocyanidin with (+)-catechin and with the resultant
procyanidins. J. Chem. Soc. Perkin Trans. 1 1983, 0, 1711-1717.
(9) Tanaka, T.; Takahashi, R.; Kouno, I.; Nonaka, G. I. Chemical evidence for the de-
astringency (insolubilization of tannins) of persimmon fruit. J. Chem. Soc. Perkin
Trans. 1 1994, 0, 3013-3022.
(10)
Cédric, D. T.; Marie-Lucie, K. C.; Vesna, J.; Sonia, C. Degradation of (−)-
Epicatechin and Procyanidin B2 in Aqueous and Lipidic Model Systems. First
Evidence of “Chemical” Flavan-3-ol Oligomers in Processed Cocoa. J. Agric. Food
Chem., 2014, 62, 9002–9016.
(11)
Cédric, D. T.; Marie-Lucie, K. C.; Vesna, J.; Sonia, C. Occurrence and
Antioxidant Activity of C1 Degradation Products in Cocoa. Foods., 2017, 6, 18.
(12)
Ishino, N.; Yanase, E.; Nakatsuka, S. Epimerization of tea catechins under
weakly acidic and alkaline conditions. Biosci. Biotechnol. Biochem., 2010, 74, 875-
877.
(13)
Callemien, D.; Collin, S. Involvement of Flavanoids in Beer Color Instability
during Storage. J. Agric. Food Chem., 2007, 55, 9066-9073.
20
ACS Paragon Plus Environment

Page 21 of 31
Journal of Agricultural and Food Chemistry
338
339
340
341
342
343
344
345
346
347
348
349
350
351
352
353
354
(14)
Guyot, S.; Vercauteren, J.; Cheynier, V. Structural determination of colourless
and yellow dimers resulting from (+)-catechin coupling catalyzed by grape
polyphenoloxidase. Phytochem., 1996, 42, 1279-1288
(15)
Tanaka, T.; Matsuo, Y.; Kouno, I. Chemistry of Secondary Polyphenols
Produced during Processing of Tea and Selected Foods Int. J. Mol. Sci., 2010, 11(1),
14–40
(16)
Uchida, K.; Ogawa, K.; Yanase, E. Structure Determination of Novel
Oxidation Products from Epicatechin: Thearubigin-Like Molecules. Molecules,
2016, 21 (273), 12-14.
(17)
Hanen, F.; Samia, O.; Sylvain, G.; Alia, B. D.; Christian, M.; Chedly, A.;
Riadh, K. LC/ESI-MS/MS characterization of procyanidins and propelargonidins
responsible for the strong antioxidant activity of the edible halophyte
Mesembryanthemum edule L. Food Chem. 2011, 127, 1732-1738.
(18)
Gu, L.; Kelm, M. A.; Hammerstone, J. F.; Zhang, Z.; Beecher, G.; Holden, J.;
Haytowitz, D.; Prior, R. L. Liquid chromatographic/electrospray ionization mass
spectrometric studies of proanthocyanidins in foods. J. Mass Spectrom. 2003, 38,
1272-1280.
21
ACS Paragon Plus Environment

Journal of Agricultural and Food Chemistry
Page 22 of 31
355
356
357
358
359
360
361
362
363
364
(19)
Morimoto, S.; Nonaka, G.; Nishioka, I. Tannins and Related Compounds.
LIX.Aesculitannins, Novel Proanthocyanidins with Doubly-Bonded Structures from
Aesculus hippocastanum L. Chem. Pharm. Bull. 1987, 35, 4717-4729.
(20)
Kondo, K.; Kurihara, M.; Fukuhara, K.; Tanaka, T.;Suzuki, T.; Miyata, N.;
Toyoda, M. Conversion of procyanidin B-type (catechin dimer) to A-type: evidence
for abstraction of C-2 hydrogen in catechin during radical oxidation. Tetrahedron
Lett. 2000, 41, 485–488.
(21)
Osman, A.M.; Wong, K.K.Y. Laccase (EC 1.10.3.2) catalyses the conversion
of procyanidin B-2 (epicatechin dimer) to type A-2. Tetrahedron Lett. 2007, 48,
1163–1167.
22
ACS Paragon Plus Environment

Page 23 of 31
Journal of Agricultural and Food Chemistry
365
366
FUNDING SOURCE
This work was partially supported by the Tojuro Iijima Foundation for Food Science and
Technology (Chiba, Japan).
367
23
ACS Paragon Plus Environment

Journal of Agricultural and Food Chemistry
Page 24 of 31
368
369
370
371
372
373
374
375
376
377
378
379
380
FIGURE CAPTIONS
Figure 1 Chemical structures of catechin (1), taxifolin (2), procyanidin B3 (3),
procyanidin C2 (4), and oxidized procyanidin B3 (5)
Figure 2 UV spectra of procyanidin B3 and procyanidin C2 after oxidation for various
times: (A) compound 3 (procyanidin B3), (B) compound 4 (procyanidin C2), and (C)
UV absorptions at 330 nm as functions of time. Values are expressed as means with
standard error bars (n=3).
Figure 3 Relative contents of compounds 3 and 4 during oxidation as functions of time.
Figure 4 UPLC/MS traces of the mixtures obtained following the oxidation of
procyanidin C2 (4) for: A) 0.5 h and B) 4 h.
Figure 5 UPLC-MS/MS spectra of thiolysis degradation products: (A) m/z 575, (B) m/z
653, and (C) m/z 651.
Figure 6 Proposed structure of 6 (peak k) and its fragmentations as determined by
MS/MS.
381
382
24
ACS Paragon Plus Environment

Page 25 of 31
Journal of Agricultural and Food Chemistry
Figure 1
OH
OH
HO
O
OH
OH
O
OH
OH
OH
HO
OH
OH
HO
O
O
HO
O
OH
+
OH
OH
OH
OH
OH
OH
n
1
2
Taxifolin ( )
(+)-Catechin ( )
3
n=1: Procyanidin B3 ( )
4
n=2: Procyanidin C2 ( )
OH
HO
O
OH
OH
OH
Oxidation
3
Compound
OH
O
O
OH
OH
etc.
O
5
Compound
25
ACS Paragon Plus Environment

Journal of Agricultural and Food Chemistry
Page 26 of 31
Figure 2
(A)
0.5
0.4
0.3
0.2
0.1
0
200
300
400
nm
500
600
(B)
0.5
0h
0.4
0.3
0.2
0.1
0
0.5h
1h
2h
4h
200
300
400
nm
500
600
(C)
0.5
0.4
0.3
0.2
0.1
0
3
4
0
1
2
time (h)
26
ACS Paragon Plus Environment

Page 27 of 31
Journal of Agricultural and Food Chemistry
Figure 3
100
90
80
70
60
50
40
30
20
10
0
3
4
0
10
20
30
40
50
60
time (min)
27
ACS Paragon Plus Environment

Journal of Agricultural and Food Chemistry
Page 28 of 31
Figure 4
(A)
25000
4
0.5 h
f
k
280 nm
g
e
i
j
20000
15000
10000
5000
0
861 [M-H]-
863 [M-H]-
TIC
10
0
2
4
6
8
time (min)
(B)
4 h
25000
b
f
e
h
d
a
c
280 nm
j
k
i
20000
15000
10000
5000
0
861 [M-H]-
863 [M-H]-
TIC
0
2
4
6
8
10
time (min)
28
ACS Paragon Plus Environment

Page 29 of 31
Journal of Agricultural and Food Chemistry
Figure 5
Δ 2 Da
(A)
(B)
Δ 78 Da
(C)
Δ 78 Da
29
ACS Paragon Plus Environment