Stability of Hydroxycinnamic Acid Derivatives, Flavonol Glycosides, and Anthocyanins in Black Currant Juice

Article abstract of DOI:10.1021/acs.jafc.6b01005

The stability of phenolic compounds was followed in black currant juice at ambient temperatures (in light and in dark conditions) and at +4 °C for a year. Analyses were based on high-performance liquid chromatography-diode-array detection-electrospray ionization-mass spectrometry (or tandem mass spectrometry) and high-performance liquid chromatography-diode-array detection-electrospray ionization-quadrupole time-of-flight mass spectrometry methods supported by nuclear magnetic resonance after selective high-performance liquid chromatography isolation. Altogether, 43 metabolites were identified, of which 2-(Z)-p-coumaroyloxymethylene-4-β-d-glucopyranosyloxy-2-(Z)-butenenitrile, 2-(E)-caffeoyloxymethylene-4-β-d-glucopyranosyloxy-2-(Z)-butenenitrile, 1-O-(Z)-p-coumaroyl-β-d-glucopyranose, (Z)-p-coumaric acid 4-O-β-d-glucopyranoside, and (Z)-p-coumaric acid were novel findings in black currant juice. Hydroxycinnamic acid derivatives degraded 20-40% at room temperature during one year of storage, releasing free hydroxycinnamic acids. O-Glucosides of hydroxycinnamic acid compounds were the most stable, followed by O-acylquinic acids, acyloxymethyleneglucosyloxybutenenitriles, and O-acylglucoses. Light induced the isomerization of (E)-coumaric acid compounds into corresponding Z-isomers. Flavonol glycosides stayed fairly stable. Flavonol aglycones were derived mainly from malonylglucosides. Over 90% of anthocyanins were lost at room temperature in a year, practically independent of light. Storage at low temperatures, preferably excluding light, is necessary to retain the original composition of phenolic compounds.

Full text of DOI:10.1021/acs.jafc.6b01005

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Article
Stability of Hydroxycinnamic Acid Derivatives, Flavonol
Glycosides and Anthocyanins in Black Currant Juice
Leenamaija Mäkilä, Oskar A. Laaksonen, Aino-Liisa Maria Alanne,
Maaria Katariina Kortesniemi, Heikki P. T. Kallio, and Baoru Yang
J. Agric. Food Chem., Just Accepted Manuscript • DOI: 10.1021/acs.jafc.6b01005 • Publication Date (Web): 04 May 2016
Downloaded from http://pubs.acs.org on May 11, 2016
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Stability of Hydroxycinnamic Acid Derivatives, Flavonol Glycosides and Anthocyanins in Black
Currant Juice
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Leenamaija Mäkilä †,*, Oskar Laaksonen †, AinoꢀLiisa Alanne ‡, Maaria Kortesniemi †, Heikki Kallio
†,§, Baoru Yang †
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† Food Chemistry and Food Development, Department of Biochemistry, University of Turku, FIꢀ20014
Turku, Finland
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‡ Instrument Centre, Department of Chemistry, University of Turku, FIꢀ20014, Turku, Finland
§ Department of Food Science and Engineering, Jinan University, 510632 Guangzhou, China
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* Corresponding author (Tel: + 358 2 333 6871; Fax: +358 2 333 6860: Eꢀmail:
leenamaija.makila@utu.fi)
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Abstract
Stability of phenolic compounds was followed in black currant juice at ambient temperature (in light
and in dark) and at + 4 °C for a year. Analyses were based on HPLCꢀDADꢀESIꢀMS(ꢀMS2) and HPLCꢀ
DADꢀESIꢀQꢀTOFꢀMS methods supported by NMR after selective HPLC isolation. Altogether 43
phenolic compounds were identified, of which 2ꢀ(Z)ꢀpꢀcoumaroyloxymethyleneꢀ4ꢀβꢀ
glucopyranosyloxyꢀ2ꢀ(Z)ꢀbutenenitrile, 2ꢀ(E)ꢀcaffeoyloxymethyleneꢀ4ꢀβꢀ ꢀglucopyranosyloxyꢀ2ꢀ(Z)ꢀ
butenenitrile, 1ꢀOꢀ(Z)ꢀpꢀcoumaroylꢀβꢀ ꢀglucopyranose, (Z)ꢀpꢀcoumaric acid 4ꢀOꢀβꢀ ꢀglucopyranoside
Dꢀ
D
D
D
and (Z)ꢀpꢀcoumaric acid were novel findings in black currant juice. Hydroxycinnamic acid derivatives
degraded 20–40% at room temperature during oneꢀyear storage, releasing free hydroxycinnamic acids.
OꢀGlucosides of hydroxycinnamic acid compounds were the most stable followed by Oꢀacylquinic
acids, acyloxymethyleneglucosyloxybutenenitriles and Oꢀacylglucoses. Light induced isomerization of
(E)ꢀcoumaric acid compounds into corresponding Zꢀisomers. Flavonol glycosides stayed fairly stable.
Flavonol aglycones derived mainly from malonylglucosides. Over 90% of anthocyanins were lost at
room temperature in a year, practically independent on light. Storage at low temperatures, preferably
excluding light, is necessary to retain the original composition of phenolic compounds.
Keywords: anthocyanins, black currant, flavonols, hydroxycinnamic acids, storage
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INTRODUCTION
Fruits and berries are rich in phenolic compounds, such as anthocyanins, flavonols and phenolic
acids. The compounds have been found to influence positively on human health.1 They are synthesized
via the phenylpropanoid pathway as secondary metabolites during development of plants, especially in
response to UV radiation and other stress factors.2 Anthocyanins are water soluble pigments that
deliver blue and magenta colors to various fruits, berries and vegetables.3 Flavonols are accumulated
mainly in the skin of fruits due to the biosynthesis stimulated by light.4 The highest concentrations of
phenolic acids are also found in the skin of fruits, where they are esterified with pectins and
arabinoxylans constructing the plant cell wall, with decreasing trend as the fruit ripens.3
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In nature, phenolic compounds exist as glycosides and other derivatives. The most common
anthocyanins and flavonols occur as Oꢀglycosides at the carbon positions 3ꢀ, 5ꢀ or 7ꢀ.5 A less studied
group of phenolics, hydroxyphenylpropenoic acids, trivially named as hydroxycinnamic acids (HCAs),
exist commonly as Oꢀacylglucoses, Oꢀacylquinic acids or as Oꢀglucosides.2 The most common HCAs
found in nature are coumaric (compound 10, Figure 1), caffeic (8) and ferulic acids (12). The acids
rarely exist in free form unless the derivatives have undergone various processing steps6 or storage.7
Both Eꢀ and Zꢀforms of HCAs occur in plants, but the Eꢀform is predominant, and also found to be
more stable.8 Other types of HCA derivatives are nitrile containing glucosides. This type of compounds
were identified as coumaric and ferulic acid derivatives from black currant seed.9 The nitriles are nonꢀ
cyanogenic βꢀglucosides of βꢀ or γꢀhydroxynitriles, which do not release toxic hydrogen cyanide upon
hydrolysis as a plant defense mechanism, as opposed to αꢀhydroxynitrile βꢀglucosides.10
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HCAs have been demonstrated to possess various biological activities in vitro and in vivo, such as
antioxidative, antiꢀinflammatory, antiꢀmutagenic, antiꢀbacterial, and antiꢀcancer properties.2,11 The
antioxidant capacity of free HCAs is shown to be higher than their corresponding glucose esters.12
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Similarly, flavonol aglycones were found with stronger antioxidant capacity than their glycosides.13
Anthocyanins and anthocyanidins vary in their antioxidant power depending on the structures.14
Phenolic compounds contribute to sensory quality of products rich in phenolic compounds, such as
fruit and berry juices. Especially hydroxycinnamic acids and flavonols are responsible of bitter and
astringent sensations of the juices.15ꢀ17 Storage often leads to changes in phenolic compounds due to
temperature, light, pH, enzymes and oxygen. These factors induce hydrolysis of derivatives, releasing
free HCAs and flavonol aglycones to the food matrix. 7,18,19 Hydrolysis of anthocyanidin glycosides
releases free anthocyanidins, which are unstable, especially at pH values higher than 2. Thus,
anthocyanidins are usually degraded further to hydroxybenzoic acids by thermal degradation20,21, or
they may polymerize or become coꢀpigmented with e.g. other phenolic compounds, forming more
stable stuctures.7,22,23 Long shelfꢀlife is often applied to the fruit and berry products, which may cause
drastic changes in phenolic compounds altering the sensory qualities. It is of high importance to study
the impact of storage on phenolics at molecular level. The degradation of anthocyanins is fairly widely
known. There is a severe lack of knowledge on the fate of individual HCA derivatives during storage,
since earlier studies have mainly concentrated on the changes in the content of free HCAs.7,18,19 Also,
the structural stability of flavonols and their derivatives has not been sufficiently investigated in
natural products.
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The objective of this research was to investigate the structural stability of glycosides and other
derivatives of phenolic compounds in black currant juice, pressed without macerating enzymes.
Thorough identification of various derivatives of HCAs, flavonol and anthocyanidin glycosides as well
as their degradation products, including aglycones, was conducted. The analytical tools for
identification were HPLC coupled with DAD, ESIꢀMS(ꢀMS2), QꢀTOFꢀMS, and NMRꢀanalysis after
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preparative HPLC fractionation. The main purpose of the study was to reveal the compositional
changes of the less investigated HCA derivatives during storage.
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MATERIALS AND METHODS
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Materials. Black currants (Ribes nigrum L.) of cultivar 'Mortti' were cultivated in Southern
Finland [Natural Resources Institute Finland (Luke) Piikkiö, Finland] and harvested in August 2010.
Berries were frozen right after harvesting and kept at – 20 °C before juice pressing. Reference
compounds of anthocyanins, delphinidin and cyanidin chlorides and flavonols were obtained from
Extrasynthese (Genay, France) and the phenolic acids from Sigma (St. Louis, MO).
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Sample preparation. Berries were pressed with a hydraulic laboratoryꢀscale juice extractor
without enzyme supplementation.15 The juice yield was fixed to 60% using a pressure of 14 MPa. The
juice was degassed in vacuum for three minutes. Juice was pasteurized in 50 mL transparent glass test
tubes in a water bath at 100 °C. The juice temperature in the central part of the test tubes was
monitored with a thermoꢀsensor Testo 177ꢀT4 (Testo AG, Lenzkirch, Germany). Temperature was
raised to 95 °C in 20–30 min, and kept at 95–97 °C for 30 seconds. Thereafter, the juiceꢀtubes were
transferred into an ice bath to cool down to approximately 20 °C in 30 min. The samples were stored
for 3, 6, 9 or 12 months in refrigerator (+ 4 °C) or at ambient room temperature (the temperature
varying from 19–27 °C). The samples stored at room temperature were kept on the same laboratory
shelf either in dark (RT in dark) or exposed to surrounding light (RT in light). A juice sample was
taken right after pasteurization to serve as the “baseline”, to which other storage samples were
compared. The baseline sample and the juice from each storage time point were stored in 10 mL falcon
tubes at –20 °C until further analyses.
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Before juice was transferred into falcon tubes, the microbial quality was tested. Cultivations were
carried out using potato dextrose agar (LAB 98) and tryptone glucose extract agar (LAB 63) by the
poured plate method (Lab M Limited, Lancashire, UK). No growth was detected in any samples in any
storage time points.
Anthocyanins were extracted from the juices with acidified MeOH, as described in our previous
studies.15,24,25 The rest of the compounds were extracted with ethyl acetate by modifying an earlier
method.25 A portion of 15 g of juice was weighed and extracted four times with 10 mL of ethyl acetate.
The extracts were combined and the evaporated product was diluted in 2 mL of MeOH and filtered (0.2
µm) for further analysis.
HPLC-DAD-ESI-MS(-MS2) analysis. All the phenolic compounds and peaks 17 and 18 (Figure
1, Table 1) were analyzed with HPLCꢀDADꢀESIꢀMS and hydroxycinnamic acid compounds in
addition by MS2. The apparatus was a Waters Acquity Ultra Performance LC system in combination
with a Waters 2996 DAD detector and a Waters Quattro Premier mass spectrometer (Waters Corp.,
Milford, MA) equipped with an ionꢀspray interface. An analytical column used for hydroxycinnamic
acid and flavonol compounds, degradation products of anthocyanins (except anthocyanidins), and
peaks 17 and 18, was a 150 mm × 4.60 mm i.d., 3.6 µm, Aeris PEPTIDE XBꢀC18, with a 2.00 mm ×
4.60 mm i.d. guard column of the same material (Phenomenex, Torrance, CA). The elution was carried
out with a binary solvent system. Solvent A was 0.1% formic acid in MilliꢀQ water and solvent B was
0.1% formic acid in acetonitrile. The flow rate was 1.0 mL / min and the injection volume 10 µL. The
gradient program for solvent B was 0–15 min, 2–18%; 15–20 min, 18%; 20–30 min, 18–20%; 30–35
min, 20–60%; 35–40 min, 60–2%; 40–45 min, 2%. Chromatograms were recorded by scanning the
absorption at 190–600 nm. Hydroxycinnamic acids, as well as peak 18 were monitored at 320 nm,
flavonols at 360 nm, and the anthocyanin degradation products, protocatechuic acid (34, Table 1) and
4ꢀhydroxybenzoic acid (35) at 250 nm, phloroglucinaldehyde (36) at 290 nm and peak 17 at 290 nm.
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After the DAD, a flow of 0.4 mL / min was directed to the mass spectrometer. The MSꢀdata was
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acquired simultaneously in positive and negative ion modes, separately for MS2ꢀdata, by applying the
methods of Liu et al.26 with modifications. The capillary voltage was 3.5 kV and 3.6 kV, the cone
voltage 15 V and 22 V and the extractor voltage 3 V and 4 V, respectively, for the positive and
negative ion mode. For the MS2 analysis the capillary voltage was 3 kV and 3.6 kV, cone voltage 22 V
and 22 V and extractor voltage 5 V and 4 V, respectively, for the positive and negative ion mode. The
source temperature was 120 °C, and the desolvation temperature 300 °C. The desolvation gas flow was
600 L / h, and the cone gas flow 97 L / h. The collision energy for the daughter ion scan was 30 eV.
Mass spectra (MS) were obtained by scanning larger ions between m/z 220 and 1000 and smaller ions
between m/z 80 and 600 (full scan). MassLynx 4.1 software was used to operate the HPLCꢀESIꢀMS(ꢀ
MS2) system.
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In MS analysis of anthocyanins, the analytical column used was a 100 mm × 4.6 mm i.d., 2.6 ꢁm,
Kinetex C18 100A, with a guard column of the same support material (Phenomenex, Torrance, CA).
Solvent A was 5% formic acid in MilliꢀQ water and solvent B was 100% acetonitrile. The flow rate
was 1.0 mL / min and the injection volume 10 µL. The gradient program for solvent B was 0–1 min, 4–
6%; 1–2 min, 6–8%; 2–6 min, 8–9%; 6–7 min, 9%; 7–10 min, 9–10%; 10–14 min, 10–11%; 14–18
min, 11%; 18–20 min, 11–18%; 20–21 min, 18–24%; 21–28 min, 24–80%; 28–29 min, 80–20%; 29–
31 min, 20–4%; 31–35 min, 4%. Chromatograms were recorded by scanning absorption at 190–600
nm. Anthocyanins were recorded at 520 nm. After the DAD, a flow of 0.2 mL / min was directed to the
mass spectrometer operated in the positive ion mode. The capillary voltage was 3.25 kV, the cone
voltage 30 V and the extractor voltage 2.5 V. The source temperature was 150 °C, and the desolvation
temperature 400 °C. The desolvation gas flow was 800 L / h, and the cone gas flow 47 L / h. Mass
spectra were obtained by scanning ions between m/z 250 and 1000 (full scan).
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HPLC-DAD-ESI-Q-TOF-MS analysis. The structures of hydroxycinnamic acid and flavonol
metabolites and compounds 17 and 18 were further verified with LCꢀDADꢀESIꢀQꢀTOFꢀMS system,
which consisted of an Agilent HPLC 1200 Series instrument equipped with an Agilent G1315C
Starlight diode array detector (Agilent Technologies, Waldbronn, Germany) and micrOTOFQ ESIꢀmass
spectrometer (Bruker Daltonics, Bremen, Germany). The column and the HPLC method were the same
as applied for HPLCꢀDADꢀESIꢀMS(ꢀMS2) analysis of HCA and flavonol compounds. After the DAD,
a flow of 0.3 mL / min was directed to the ion source. The HPLC system was controlled by Hystar
software version 3.2 (Bruker Biospin, Rheinstetten, Germany). The mass spectrometer was controlled
by Bruker Compass 1.3 for micrOTOF control software (Bruker Daltonics, Bremen, Germany) and
operated in the positive and negative ion mode by applying the earlier method with modifications.27
The capillary voltage was 4.5 kV in positive ion mode and 4 kV in negative ion mode. The end plate
offset was at –500 V. The pressure of the nebulizer gas (N2) was set at 1.6 bar. The drying gas (N2)
flow was 12 L / min, and the drying gas temperature 200 °C. The mass range was from m/z 50 up to
2000 (full scan). The calibration was carried out with 5 mM sodium formate injected via a sixꢀportꢀ
valve at the end of LCꢀMS experiment in order to provide highꢀaccuracy mass measurements. The
internal mass spectrum calibration was performed with HPC mode. The minimal number of calibration
points was seven, and the standard deviation was below 1.5 ppm. The data were handled by Bruker
Compass DataAnalysis (version 4.0 SP2; Bruker Daltonics, Bremen, Germany).
Quantitative analysis. For quantitative analysis the stored samples were analyzed in duplicates
and the baseline juice five times. The duplicate samples mainly from separate pasteurization batches
were prepared identically. All the compounds were quantitated with a Shimadzu LCꢀ10AVP HPLC
(Shimadzu, Kyoto, Japan) with a LCꢀ10AT pump, a SILꢀ10A autosampler and a SDPꢀM10AVP diode
array detector linked to the SCLꢀM10AVP data handling station. The column temperature was set to 30
°C. For hydroxycinnamic acid and flavonol compounds, degradation products of anthocyanins (except
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anthocyanidins), and compounds 17 and 18, the column, the binary gradient system, the injection
volume and the wavelengths for monitoring the UV absorption spectra were the same as described
earlier. For anthocyanin analyses, our earlier methods were used.15,24,25 The analysis was carried out
with an external standard method based on calibration curves. The total content of anthocyanins stored
at various conditions was calculated by defining the descending curve y = A × e(ꢀx/t) + y0 and using
Origin 8 software (Originlab Corp., Northampton, MA).16
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Purification and NMR analysis of selected compounds. Juice (4 kg) pressed in larger scale28
was extracted with ethyl acetate following the procedure described above. A portion of 1ꢀ4 g of the
evaporated extract was diluted into MilliꢀQ water (40 °C) to reach a final concentration of 100 mg /
mL. The diluted sample was applied in a glass column (230 mm × 26 mm i.d.) packed with 34 g of
Sephadex LHꢀ20 (average dry particle size 70 µm) swollen in MilliꢀQ water overnight. The extract was
purified with EtOH –MilliꢀQ water according to the elution gradient29 as follows: F1A (50 mL), 100%
water; F1B (200 mL), 100% water; F2 (200–300 mL), 10% EtOH in water; F3 (200–300 mL), 20%
EtOH in water; F4 (200–300 mL), 30% EtOH in water; F5 (200–300 mL), 40% EtOH in water. An
elution volume of 200 mL was used, when the amount of sample applied into the column was 1–2 g. A
volume of 300 mL was used, when the sample amount was 3–4 g. Gradient was continued by stepwise
increasing ethanol content from 50% to 80%, followed by aqueous acetone of two concentrations of
30% and 70%. Selected unknown compounds 14, 15, 17 and 18 (Table 1, Figure 1) eluted in fractions:
F1B, 17; F2–3, 14 and 15 and F4–5, 18. The elution speed was maintained at 21 mL / min with a
peristaltic pump. The fractions were evaporated to dryness using a rotary evaporator at 39 °C. Each
dried fraction was recollected in 4–5 mL of MeOH. The fractions were analyzed with HPLCꢀDAD and
HPLCꢀDADꢀESIꢀMS, using the same methods.
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Compounds 14, 15, 17 and 18 in the Sephadex LHꢀ20 fractions (F1BꢀF5) were purified by a semiꢀ
preparative system consisting of a SILꢀ20A autosampler, an LCꢀ20AB pump, a CTOꢀ10AC column
oven, an SPDꢀ20A UVꢀdetector and a DGUꢀ20A5 degasser. The column was a 250 mm × 10.0 mm
i.d., 5 µm, Aeris PEPTIDE XBꢀC18, with a 10 mm x 10 mm i.d. guard column of the same material
(Phenomenex, Torrance, CA). The fractions obtained from Sephadex purification were dissolved in
MeOH:H2O (1:1) and filtered (0.2 µm) before injection. Elution was carried out with a binary solvent
system. Solvent A was MilliꢀQ water for isolation of the peak 17. For the rest of the compounds 14, 15
and 18 solvent A was 0.1% formic acid in water. Solvent B was 100% acetonitrile for the isolation of
all the analytes. For 17, the gradient program for solvent B was 0–9 min, 2%; 9–20 min, 2–20%; 20–30
min, 20–35%; 30–35 min, 35–68%; 35–40 min, 68–2%; 40–45 min, 2%. For isolating 14 and 15, the
gradient program for solvent B was 0–15 min, 2–18%; 15–24 min, 18%; 24–25 min, 18–14%; 25–34
min, 14–17%; 34–38 min, 17–65%; 38–40 min, 65–2%; 40–45 min, 2%. For 18, the gradient program
for solvent B was 0–15 min, 2–18%; 15–20 min, 18%; 20–30 min, 18–20%; 30–35 min, 20–60%; 35–
40 min, 60–2%; 40–45 min, 2%. The flow rate was 4.9 mL / min. The column oven was kept at 40 °C.
The chromatograms were recorded at 320 nm. Injections, ranging from 500–1500 µL, were repeated
25–50 times for each peak.
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For confirmation of successful collections, the fractions were reꢀanalyzed with HPLCꢀDAD and
HPLCꢀDADꢀESIꢀMS. Fractions of the same peaks were combined and the solvents evaporated using a
rotary evaporator at 30 °C, approximately for 45 min. After evaporation, the samples were dissolved in
MilliꢀQ water and freezeꢀdried.
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1H and 13C NMR spectra were obtained with an Avance Bruker 500 spectrometer (Bruker BioSpin
AG, Fällanden, Switzerland) operating at 500.13 and 125.77 MHz, respectively. Spectra were recorded
at 25 °C using CD3OD as the solvent with a nonꢀspinning sample in a 5 mm NMRꢀtube. The spectra
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were calibrated on the solvent residual signal of CD3OD at 3.31 ppm for 1H and 49.15 ppm for 13C and
processed by TopSpin 3.2 software.
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2ꢀ(Z)ꢀpꢀcoumaroyloxymethyleneꢀ4ꢀβꢀ
D
ꢀglucopyranosyloxyꢀ2ꢀ(Z)ꢀbutenenitrile (15, Figure 1). H
NMR (CD3OD, 500.13 MHz): δ 7.67 (2ʹʹ, 6ʹʹ, d, J = 8.7 Hz), 6.95 (7ʹʹ, d, J = 12.7 Hz), 6.81 (3, dd, J =
6.5 Hz, J = 5.9 Hz), 6.76 (3ʹʹ, 5ʹʹ, d, J = 8.7 Hz), 5.81 (8ʹʹ, d, J = 12.7 Hz), 4.77 (10ʹʹ, s), 4.64 (4a, dd, J
= 14.5, J = 5.9), 4.51 (4b, dd, J = 14.5, J = 6.5), 4.33 (1ʹ, d, J = 7.8 Hz), 3.88 (6ʹa, dd, J = 12.0 Hz, J =
2.1 Hz), 3.68 (6ʹb, dd, J = 12.0 Hz, J = 5.2 Hz), 3.39–3.26 (3ʹ, 4ʹ, 5ʹ, m), 3.21 (2ʹ, dd, J = 7.8 Hz, J =
13
8.9 Hz). C NMR (CD3OD, 125.77 MHz): 167.2 (Cꢀ9ʹʹ), 160.5 (Cꢀ4ʹʹ), 149.3 (Cꢀ3), 146.9 (Cꢀ7ʹʹ),
134.1 (Cꢀ2ʹʹ, Cꢀ6ʹʹ), 127.1 (Cꢀ1ʹʹ), 116.3 (Cꢀ1), 116.1 (Cꢀ3ʹʹ, Cꢀ5ʹʹ), 115.4 (Cꢀ8ʹʹ), 112.9 (Cꢀ2), 104.4
(Cꢀ1ʹ), 78.2, 78.1, 71.6 (Cꢀ3ʹ, Cꢀ4ʹ, Cꢀ5ʹ), 75.1 (Cꢀ2ʹ), 68.4 (Cꢀ4), 64.3 (Cꢀ10ʹʹ), 62.7 (Cꢀ6ʹ).
242
243
244
245
246
247
248
249
250
251
252
253
254
UV-A and sunlight irradiation of HCAs. Each reference compound of Eꢀisomers of HCAs, i.e.
pꢀcoumaric acid, ferulic acid, caffeic acid, chlorogenic acid (3ꢀOꢀcaffeoylquinic acid) and
neochlorogenic acid (5ꢀOꢀcaffeoylquinic acid) was irradiated separately in water solutions by UVꢀA
and by sunlight. Concentrations of the reference compound in water solutions were 0.8 mg / mL,
except of neochlorogenic acid was 0.1 mg / mL due to shortage of the compound. The solutions were
divided in two batches of 25 mL volume each, one batch to be irradiated with UVꢀA and the other with
direct sunlight. The solutions were transferred into 50 mL transparent glass test tubes, same as used in
the storage of juice samples. The solutions of reference compounds were irradiated with a ledꢀlight
emitting UVꢀA radiation (360–370 nm) with intensity of 41 W/m2. The intensity was measured with
photo/radiometer HD 2102.1 using a probe LP 471 UVA (Delta OHM, Padua, Italy). The distance of
the probe to ledꢀlight was the same as distance from the bottom of the test tube to ledꢀlight (1.0 cm).
Height of the spout irradiated was 6.5 cm and duration of irradiation 1.0 hour. The purified peak 14
(concentration 1.8 mg / mL) was also irradiated with UVꢀA, but in 3 mL MeOH with a spout of 1.2 cm.
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The reference compounds of (E)ꢀpꢀcoumaric, (E)ꢀferulic and (E)ꢀcaffeic acid were kept in direct
sunlight for 7 days in the middle of August 2015 and chlorogenic and neochlorogenic acid for 6 days.
Statistical analyses. Influence of storage on the metabolites was determined by analyses of
variance (ANOVA) together with Tukey’s post hoc and LSD test. Oneꢀway ANOVA was used for
comparing the storage conditions at each timeꢀpoint, and for comparing the samples in different
conditions at all the time points (3–12 months of storage samples) with the baseline sample. Oneꢀway
ANOVA was used also to compare the changes in the content of various groups of compounds.
Student’s tꢀtest (equal variances assumed) was applied with less than three variables. The criterion for
statistical significances was p < 0.05. ANOVA analysis and tꢀtests were performed using SPSS 21.0
(SPSS Inc. Chicago, IL).
RESULTS AND DISCUSSION
Identification of the metabolites. Identification of the analytes extracted with acidified methanol
and ethyl acetate from black currant juices were based on the HPLC retention behavior, DAD spectra,
1
ESIꢀMS(ꢀMS2) spectra, ESIꢀQꢀTOFꢀMS spectra, and H and 13C NMR analysis of the isolated
compounds, as well as analyses of reference compounds and literature comparisons.9,15ꢀ17,20ꢀ22,24,25,30ꢀ43
For NMR analyses the extracts were preꢀfractionated before HPLC isolation. Based on the results, the
compounds were classified either as identified or tentatively identified (Table 1, Figure 1). The
calculations to determine the probable origin of positive ion fragments in MS2 ꢀdata in Table 1 were
based on previous reports.30,32
275
276
277
The less investigated group of phenolic compounds, hydroxycinnamic acids (HCAs), was of
special interest in our study. The HCAs were identified as caffeic, coumaric and ferulic acid
derivatives (Figure 2, Table 1). Previously identified 1ꢀOꢀ(E)ꢀcaffeoylꢀ (peak 3), 1ꢀOꢀ(E)ꢀpꢀ
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coumaroylꢀ (6) and 1ꢀOꢀ(E)ꢀferuloylꢀβꢀ
D
ꢀglucopyranoses (9)24,25,36 (Figure 1) were confirmed based
on the bathochromic shift compared to the corresponding free HCA reference compound.32 These
Oꢀacylated glucoses of HCAs shared a similar MS2 fragmentation pattern of [M + H – Glc – H2O]+
in the positive ion mode. Tentatively identified HCA Oꢀglucosides, presumably 4ꢀOꢀβꢀDꢀ
glucopyranosides 17,32,33 of (E)ꢀcaffeic and (E)ꢀpꢀcoumaric acids were found to coꢀelute (peak 2). 4ꢀ
OꢀGlucosides were identified based on similar MS and MS2 spectra as the corresponding 1ꢀOꢀ
acylglucoses, and on elution before the corresponding 1ꢀOꢀacylglucose (Table 1).34 UV spectra of
the 4ꢀOꢀglucosides also showed the indicative hypsochromic shift compared to the free HCA
reference compound.34 The 4ꢀOꢀglucoside of (E)ꢀpꢀcoumaric acid eluted before (E)ꢀ
caffeoylglucose.32 Three forms of Oꢀacylquinic acids were detected: neochlorogenic acid31 [5ꢀOꢀ
(E)ꢀcaffeoylquinic acid, peak 1], 3ꢀOꢀ(E)ꢀpꢀcoumaroylquinic acid34 (peak 4) and chlorogenic acid34
[3ꢀOꢀ(E)ꢀcaffeoylquinic acid, peak 6]. Chlorogenic acid coꢀeluted with (E)ꢀpꢀcoumaroylglucose
showing low signal intensity in MS spectra compared to (E)ꢀpꢀcoumaroylglucose (Table 1).
291
292
293
294
295
296
297
298
299
300
Tentatively identified derivatives and free (Z)ꢀpꢀcoumaric acid (peaks 5, 7, 11 and 15) were
more abundant in the chromatogram of Figure 2B compared to 2A. Zꢀisomers of pꢀcoumaric acid
compounds eluted 0.5–2 minutes after the corresponding Eꢀisomers37, and had similar MS and MS2
spectra, but the UV spectra of the Zꢀisomers showed hypsochromic shift compared to (E)ꢀisomers. 35
From previous studies,17,35 the 4ꢀOꢀglucoside of (Z)ꢀpꢀcoumaric acid was also found to elute before
the corresponding Eꢀisomer. (Z)ꢀpꢀcoumaric acid was identified with a reference compound;
irradiation with UVꢀA light and sunlight led to partial conversion of (E)ꢀpꢀcoumaric acid to the Zꢀ
isomer (Figure 3A). To our knowledge, this was the first time (Z)ꢀpꢀcoumaric acid and its
derivatives were reported in black currant juice. Zꢀconfigurations of caffeic and ferulic acids as free
form or derivatives were not detected.
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pꢀCoumaric acid derivatives dominated in black currant juice whereas ferulic acid derivatives
were not common. Feruloylquinic acid and 4ꢀOꢀglucoside of ferulic acid were either not found, or
they existed in trace amounts only. HCA derivatives of special interest were the four compounds
13–16 with higher molecular weight (Table 1, Figure 1 and 2), of which peaks 14 and 16 have
earlier been identified in black currant seeds.9 Odd number nominal masses [M] of m/z 437, 421 and
451 indicated the presence of odd number of nitrogen atoms in the molecules. QꢀTOFꢀMSꢀdata
confirmed this preliminary identification giving mass error: |error ppm| < 5.8 (Table 1). These
nitrogen containing compounds shared a similar fragmentation pattern, leaving an unknown residue
of m/z 95 in the mass spectra of every compound (caffeic / coumaric / ferulic acid + m/z 162 (Glc) +
m/z 95). Also MS2 showed a positive ion fragment with m/z value of 114 [M + H – (HCA – H2O) –
Glc]+ for every peak 13–16. Thus, determining the m/z 95 residue for one compound would establish
the structures for the other compounds as well. The peaks 14 and 15 gave similar UV, MS and MS2
spectra, suggesting that these compounds were geometric isomers.
314
315
316
317
318
319
Peak 14 was selected to determine the structure of the nitrogen containing m/z 95 residue. Peak
15 was selected to possibly reveal the Zꢀisomer of the peak 14. Peaks 17 and 18 were isolated for
identification, due to their greatly increased contents during storage (Figure 2B). The UV spectra of
these compounds did not correspond to HCA. After isolation with preparative HPLC, the structures
1
13
of peaks 14, 15, 17 and 18 were further studied by H and C NMR using 1DꢀTOCSY, COSY,
HSQC and HMBC measurements as additional tools.
320
321
322
323
Peaks 14 and 15 (Figure 1) were identified as geometric isomers: 2ꢀ(E)ꢀpꢀ
coumaroyloxymethyleneꢀ4ꢀβꢀ
coumaroyloxymethyleneꢀ4ꢀβꢀ
D
D
ꢀglucopyranosyloxyꢀ2ꢀ(Z)ꢀbutenenitrile
and
2ꢀ(Z)ꢀpꢀ
ꢀglucopyranosyloxyꢀ2ꢀ(Z)ꢀbutenenitrile respectively. The NMR data
of the peak 14 was published earlier.9 The NMR data for the laterꢀeluting peak 15 has not been
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328
published previously and is now reported. Peak 17 was found to contain two forms of citric acid
monomethyl esters: 2ꢀhydroxyꢀ1,2,3ꢀpropanetricarboxylic acidꢀ2ꢀmethyl ester (symmetric, 16%)
and 2ꢀhydroxyꢀ1,2,3ꢀpropanetricarboxylic acidꢀ1ꢀmethyl ester (nonꢀsymmetric, 84%).41 The
structure of peak 18 was elucidated by NMR, and the data were consistent with those of flazin: 1ꢀ(5ꢀ
hydroxymethylꢀ2ꢀfuryl)ꢀβꢀcarbolineꢀ3ꢀcarboxylic acid.42
329
330
331
332
333
334
335
336
337
338
339
340
Compounds 13 and 16 (Figure 1) were identified as 2ꢀ(E)ꢀcaffeoyloxymethyleneꢀ4ꢀβꢀDꢀ
glucopyranosyloxyꢀ2ꢀ(Z)ꢀbutenenitrile and 2ꢀ(E)ꢀferuloyloxymethyleneꢀ4ꢀβꢀ
Dꢀglucopyranosyloxyꢀ
2ꢀ(Z)ꢀbutenenitrile, respectively. Identification was based on similar patterns in the MS, MS2 and Qꢀ
TOFꢀMS ꢀdata (Table 1) with the peak 14. Peak 15, 2ꢀ(Z)ꢀcoumaroyloxymethyleneꢀ4ꢀβꢀDꢀ
glucopyranosyloxyꢀ2ꢀ(Z)ꢀbutenenitrile was succeeded to be purified only up to 65% (Figure 4A).
The rest of the isolate, 35%, was converted into the corresponding, more stable Eꢀisomer (14). When
optimizing the chromatographic separation of the peaks 14 and 15, the difference in the retention
times of the two isomers reached even 10 minutes. Thus, the presence of Eꢀisomer was not due to
the lack of chromatographic resolution, but due to spontaneous conversion of the isomers. Previous
research17 has pointed out the difficulties in purifying Eꢀ and Zꢀisomers of coumaric acid 4ꢀOꢀ
glucosides. The isolated peak 14 had also partially (5%) converted into the corresponding Zꢀform
(Figure 4A). The purity of both isomers was enough for the structure determination with NMR.
341
342
343
344
345
346
The chromatograms of flavonol compounds and anthocyanins are shown in Figures 2C and 2E.
Flavonol glycosides were identified as flavonol 3ꢀOꢀglycosides based on reference compounds and
earlier studies.16,24 The tentatively identified aureusidin glucoside (peak 22) coꢀeluted with myricetin
3ꢀOꢀβꢀ ꢀ(6ʹʹꢀmalonyl)ꢀglucopyranoside. This was seen as a shoulder of UV absorption maximum of
D
400 nm, typical for aureusidin,36 in the spectrum of myricetin malonylglucoside (main absorption
maximum 350 nm). Aglycones myricetin (30), quercetin (31), kaempferol (32) and isorhamnetin
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(33) were seen in trace amounts in the samples after twelve months of storage (Figure 2C). The four
major anthocyanidin 3ꢀOꢀglycosides16,24 were identified as delphinidin 3ꢀOꢀβꢀ
D
ꢀglucopyranoside
(37), delphinidin 3ꢀOꢀαꢀ
Lꢀrhamnopyranosylꢀ(1→6)ꢀβꢀ
D
ꢀglucopyranoside (38), cyanidin 3ꢀOꢀβꢀ
Dꢀ
glucopyranoside (39) and cyanidin 3ꢀOꢀαꢀ
L
ꢀrhamnopyranosylꢀ(1→6)ꢀβꢀ ꢀglucopyranoside (40)
D
(Figure 2E, baseline sample).
352
353
354
355
356
357
358
359
360
361
362
Free delphinidin was detected in trace amounts and only in the baseline sample, even though the
decline of the corresponding glycosides was drastic during storage (Figure 2E, 12ꢀmonth
chromatogram). To some extent, the anthocyanins degraded to hydroxybenzoic acids. The pH of the
juice samples was ~2.9. In higher pH than 2.0, anthocyanins are thermally degraded through
formation of chalcone structure.20,21 Protocatechuic acid (peak 34) and 4ꢀhydroxybenzoic acid (35)
were two abundant peaks in the 12ꢀmonth samples (Figure 2D). They are known to be derived from
the Bꢀring of cyanidin and pelargonidin, respectively. Phloroglucinaldehyde (36) was detected in
minor concentrations from the possible cleavage of the Aꢀring of anthocyanidins.20,21 To our
knowledge, phloroglucinaldehyde has not been earlier identified from black currant. Gallic acid is
known to be derived via cleavage of the Bꢀring of delphinidin44, but was not found. Gallic acid
would have coꢀeluted with peak 17.
363
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365
366
367
368
369
Influence of storage. Table 2 summarizes the critical information related to stability of various
compound classes and individual metabolites during storage of the juice. The total content of HCA
compounds in black currant juice at baseline was 2.8 mg / 100 g of which only 6% were free acids
(Table 2). (E)ꢀcoumaroyloxymethyleneglucosyloxybutenenitrile (peak 14; 1.1 mg / 100 g juice) was
the major compound at baseline followed by (E)ꢀpꢀcoumaroylglucose / chlorogenic acid (6; 0.4),
(E)ꢀcaffeoyloxymethyleneglucosyloxybutenenitrile
(13;
(16;
0.3),
0.3),
(E)ꢀ
(Z)ꢀ
feruloyloxymethyleneglucosyloxybutenenitrile
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coumaroyloxymethyleneglucosyloxybutenenitrile (15; 0.2), (E)ꢀpꢀcoumaroylquinic acid (4; 0.1) and
(E)ꢀcaffeoylglucose (3; 0.1). Metabolites based on (E/Z)ꢀpꢀcoumaric acid (2 mg / 100 g juice)
formed the major group of HCAs followed by caffeic (0.5 mg / 100 g) and ferulic (0.4 mg / 100 g)
acid compounds.
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375
376
377
378
379
380
381
The total content of HCA compounds in juice stored at room temperature in light decreased in
three months by ca. 26%, and remained thereafter at lower level than in the other conditions. The
decrease was due to loss in the total HCA derivatives which became significant also in dark at room
temperature after nine months. Vice versa, the sum of free HCAs increased significantly during
storage at room temperature, both in light and in dark, even by a factor up to four, due to their
release via hydrolysis of their derivatives. Analogous decrease in the total HCA derivatives and
increase in the corresponding free acids in white wine have been reported earlier.19 The changes
were minimal and statistically insignificant when stored in refrigerator (+ 4 °C).
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384
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386
387
388
The most obvious change induced by storage among coumaric, caffeic and ferulic acid
compounds was increase in each of the corresponding free HCAs and decrease or minor increase in
the derivatives. The contents of free (E)ꢀcaffeic (8) and (E)ꢀferulic acids (12) increased at room
temperature by a factor 3–4 during 12 months, regardless of the light conditions. Free (E)ꢀ and (Z)ꢀ
coumaric acids (10, 11) were released in even higher degree. The very special feature was that
formation of Zꢀisomer was dominant. Especially when juice was stored in light the content increased
from 11–140 μg / 100 g (Table 2).
389
390
391
Analogously to increase in free HCAs, the contents of (E)ꢀcoumaroylꢀ (6), (E)ꢀcaffeoylꢀ (3) and
(E)ꢀferuloylglucoses
(9),
as
well
as
(E)ꢀcoumaroylꢀ
(14)
and
(E)ꢀ
feruloyloxymethyleneglucosyloxybutenenitriles (16) decreased significantly at room temperature
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but the caffeoyl derivative (13) to a lesser extent. Another special characteristic of black currant
juice was the lightꢀinduced conversion of (E)ꢀcoumaroylglucose and (E)ꢀ
coumaroyloxymethyleneglucosyloxybutenenitrile into the corresponding Zꢀforms. When stored at +
4 °C, the changes were minimal, except for some formation of (Z)ꢀcoumaric acid, which seemed to
be a sensitive indicator of the shelfꢀlife of the juice.
397
398
399
400
401
402
403
404
405
406
Stability of the sums of HCAs and their derivatives were compared to each other (Table 2).
Derivatives of (E/Z)ꢀpꢀcoumaric acids were commonly very sensitive to light, but at 12ꢀmonth time
point (E)ꢀferulic acids had degraded even more. In dark, (E)ꢀferulic acid derivatives degraded the
most, but (E)ꢀcaffeic acids stayed fairly stable. Therefore, the second OHꢀgroup in the HCA seemed
to stabilize the system. As a summary, at room temperature, Oꢀacylglucoses of HCAs were the most
labile type of derivatives followed by acyloxymethyleneglucosyloxybutenenitriles and Oꢀacylquinic
acids. HCA Oꢀglucosides were rather stable, showing even an increase in their contents. Oꢀ
Acylglucoses and Oꢀacylquinic acids of HCAs are degraded by the loss of sugar or quinic acid
moiety,
acyloxymethyleneglucosyloxybutenenitriles
by
the
loss
of
the
oxymethyleneglucosyloxybutenenitrile moiety, all resulting in free HCAs.
407
408
409
410
411
412
413
414
The pꢀcoumaric acid derivatives deserve special attention, as sunlight and UVꢀlight are known
to induce isomerization of (E)ꢀcinnamic acids into the corresponding Zꢀforms.45 Most of the
isomerization induced by UVꢀlight in nature is due to UVꢀA.46 Eꢀisomers of HCA reference
compounds and the purified (E)ꢀcoumaroyloxymethyleneglucosyloxybutenenitrile (14) were
irradiated by UVꢀA (360–370 nm) and direct sunlight (Figure 3). (E)ꢀpꢀcoumaric acid was converted
into the corresponding Zꢀisomer under UVꢀA (one hour) and sunlight (one week) by 25% and 34%,
respectively (Figure 3A). The corresponding conversions of ferulic acid were 15% and 19%,
respectively (Figure 3B). Caffeic acid, as free or Oꢀacylated to quinic acid (chlorogenic and
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417
418
419
420
neochlorogenic acids) (Figures 3CꢀE), showed only minimal conversion. The second OHꢀgroup of
caffeic acid might have stabilized the ring system. The minor peaks eluting after the Eꢀisomers of
caffeic and chlorogenic acids might have indicated the presence of the Zꢀisomers. The purified peak
of (E)ꢀcoumaroyloxymethyleneglucosyloxybutenenitrile (14) was found to contain 8% of the
corresponding Zꢀisomer (Figure 3F). After UVꢀA irradiation, proportion of the Zꢀisomer increased
up to 21%.
421
422
423
424
425
426
427
428
429
The total content of flavonol compounds in black currant juice was before storage 1.6 mg / 100
g of which 90% consisted of glycosides and the rest of aglycones (Table 2). Myricetin formed 60%
and quercetin 30% of the phenolic components, whereas kaempferol and isorhamnetin were minor
compounds. Changes of the total amount of flavonol compounds were small at + 4 °C and not
statistically significant. Table 2 indicates, however, that flavonol aglycones had a common trend to
be released at room temperature when compared with the samples in refrigerator. Free quercetin
(31) increased significantly during storage and about doubled in nine months at room temperature,
both in dark and in light. Myricetin was liberated more slowly reaching a significant change only
after nineꢀmonth storage.
430
431
432
433
434
435
436
Malonylglucosides of myricetin (22) and quercetin (26) decreased gradually during storage at
room temperature. Of quercetin malonylglucoside, only a fraction of the original quantity was left
after six months, but the corresponding myricetin compound decreased by ca. 60% only. In
refrigerator the changes did not reach the level of significance even in the more labile quercetin
derivatives. Formation of flavonol aglycones seemed to be derived from malonylglucosides, not
from other Oꢀglycosides. Bimpilas et al.18 found also an increase in flavonol aglycones during one
year storage in wines.
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438
439
440
441
442
443
444
445
446
447
448
449
Anthocyanins were clearly more affected by storage than the other phenolic compounds. In the
beginning of the trial, anthocyanins at the level of 30 mg / 100 g formed the major group of
phenolics and decreased by ca. 50% in one year in refrigerator (Table 2). The most drastic decline in
the total anthocyanins ended up to trace amounts only (0.1 – 0.2 mg / 100 g), when stored at room
temperature for one year. Reduction followed a descending curve (y = A × e(ꢀx/t) + y0) and was
practically independent of light, which was in contradiction to results of a previous study.47 By using
the equation above, it takes eight years for the anthocyanins to degrade in refrigerator to the same
extent as they are degraded at room temperature within a year. Wilkes et al.7 found that
anthocyanins decreased in a linear manner up to five months storage at 25 °C. Stability of cyanidin
and delphinidin glycosides was similar when stored at room temperature, but delphinidin glycosides
were more stable when stored in refrigerator (Table 2). The sugar moieties had no effect on the
stability of anthocyanins at room temperature. In refrigerator, glucosides seemed to be more stable
compared to rutinosides of one and the same anthocyanidin.
450
451
452
453
454
Protocatechuic acid (34), the dominant degradation product of cyanidin glycosides, showed a
fourꢀfold increase in one year at room temperature in both storage conditions. Minor degradation
products were 4ꢀhydroxybenzoic acid (35) and phloroglucinaldehyde (36). The characteristic UV
absorbance maximum of 4ꢀhydroxybenzoic acid at 253 nm was seen only after six months of storage
at room temperature. Before storage for six months the second absorbance maximum was at 217 nm.
455
456
457
458
459
Citric acid methyl esters (17, Figure 1, Figure 2B) eluted as a one broad peak. The peak 17
showed some minor increase at room temperature during storage (Table 2). Content of the peak rose
approximately 650%. Before pasteurization the second UV absorption maximum of the peak 17 was
at 270 nm, not at its characteristic 290 nm. Thus, the heat treatment (+ 95 °C) seemed to alter the
structure of the compound.
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461
462
463
464
465
466
467
468
469
Flazin (18, Figure 1), belonging to the group of βꢀcarboline alkaloids (Figure 2B) was found in
the black currant juice. Flazin is a highly fluorescent, yellowꢀcolored compound, which has been
previously isolated from fermented soy bean products.48,49 Deshydroxymethyl flazin and flazin
methyl ether have been previously revealed in fresh black currant juice, which were also formed
during storage by condensation of tryptophan and ascorbic acid or its degradation products, such as
furfural.50 Shaaban et al.49 postulated that flazin was a microbial metabolite, since a sterile nutrient
broth did not contain flazin, but the concentration increased during bacterial fermentation. The
content of flazin increased drastically during storage at room temperature, but no change was seen at
+ 4 °C. The increased concentration of flazin in stored juices without any detected microbial growth
was surprising.
470
471
472
473
474
475
476
In summary, a comprehensive profile of phenolics was revealed in black currant juice as fresh,
as well as after oneꢀyear storage in various conditions. Altogether 43 phenolic compounds were
identified, eight of which were found for the first time in the juice. Novel findings related to stability
and hydrolysis patterns of various derivatives and also to their E/Zꢀisomerism were obtained. Both
structure of the phenolic compartment and type of the derivative had an influence on the stability.
The effects of light and temperature were analyteꢀdependent but all in all, storage at + 4 °C retained
the composition well.
477
478
479
480
481
482
ACKNOWLEDGMENTS
The authors thank the Natural Resources Institute Finland (Luke, Piikkiö, Finland) for providing the
black currant berries for the study. We are also grateful to Ville Ruusunen, Eerika Vuorinen and
Tuija Niva for their help in the anthocyanin analyses. Olli Martiskainen, Jari Sinkkonen and Maarit
Karonen are warmly acknowledged for the help in identification of unknown compounds.
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ASSOCIATED CONTENT
The Supporting Information is available free of charge via the Internet at http://pubs.acs.org.
484
485
486
487
488
489
490
491
492
493
494
495
496
497
498
499
500
501
502
503
504
Table S1: List of the reference compounds used for quantitation. Figure S1: Identification of
1
anthocyanidins. Figure S2: H NMR (CD3OD, 500.13 MHz) spectrum of peak 15. Figure S3: 13C
NMR (CD3OD, 125.77 MHz) spectrum of peak 15. Figure S4: Optimizing chromatographic
separation of the peaks 14 and 15. Figure S5: Decomposition of phenolic compounds and flazin.
Figure S6: Decomposition of individual HCA compounds. Figure S7: Decomposition of individual
flavonol compounds and anthocyanins. Table S2: Concentrations of the metabolites. Figure S8:
Increase of peak 17 during pasteurization (PDF).
AUTHOR INFORMATION
Corresponding Author
* Phone: + 358 2 333 6871. Fax: +358 2 333 6860. Eꢀmail: leenamaija.makila@utu.fi
Funding
The study was funded by the Tekes – Finnish Funding Agency for Technology and Innovation
together with Finnish food companies on the project “Black Currant as Unique Source of Functional
Ingredients of Food: Novel Processes and Innovations (26001642)”, and by the Finnish Cultural
Foundation and the Magnus Ehrnrooth Foundation.
Notes
The authors declare that there is no competing financial interest.
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