Formation and characterization of polyphenol-derived red chromophores

Apr 9, 2019 - Very recently, we described the formation of (+)-catechin and (-)-epicatechin derived polar chromophores by means of a cocoa alkalizatio...
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Formation and Characterization of Polyphenol-Derived Red Chromophores. Enhancing the Color of Processed Cocoa Powders: Part 2 Daniel Germann, Timo D. Stark, and Thomas Hofmann*

J. Agric. Food Chem. Downloaded from pubs.acs.org by UNIV OF LOUISIANA AT LAFAYETTE on 04/16/19. For personal use only.

Food Chemistry and Molecular Sensory Science, Technische Universität München, Lise-Meitner-Strasse 34, 85354, Freising, Germany S Supporting Information *

ABSTRACT: Very recently, we described the formation of (+)-catechin- and (−)-epicatechin-derived polar chromophores by means of a cocoa alkalization process. In this study we focus on the generation of unpolar chromophores using model reactions via Dutching with softer alkaline conditions. UPLC-HR-ESI-TOF-MSe analysis and one-dimensional and two-dimensional NMR spectroscopy led to the unequivocal identification of dehydrodicatechin- and hydroxyxanthene-derived chromophores. The previously unknown 6-C-linked constitutional isomers of C-6′B/C-6D-(2R,3S)-dehydrodicatechin (3, 5) were first described, and furthermore, the dimers dehydrocatechin-C-6′B/C-8D-(2S,3S)-epicatechin (2) and dehydrocatechin-C-6′B/C6D-(2S,3S)-epicatechin (4, 6) as well as the reddish-colored chromophores 8-C-xanthenocatechin (7), 8-C-xanthenoepicatechin (8), 6-C-xanthenocatechin (9), and 6-C-xanthenoepicatechin (10) were identified as new compounds. A LC-MS/MS method was developed to screen and quantify both classes of chromophores as well as their precursors in alkalized cocoa powders. The dehydrocatechin dimers showed degradation via the alkalization process; in contrast, the xanthenocatechins highlighted an increase in concentration with stronger alkalization, and, therefore, contribute to cocoa reddening. These results, together with those previously published, give a clear insight into the chemistry of polyphenol-derived chromophores generated by cocoa powder alkalization and enable a better understanding of chromophore formation mechanisms toward a more comprehensive color design of cocoa powders. KEYWORDS: cocoa powder, alkalization, model reaction, polyphenol, catechin, epicatechin, catechinic acid, dehydrodicatechin, dehydrocatechin dimer, chromophore

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grade (Merck). Deuterated solvents for NMR experiments were obtained from Sigma-Aldrich (Steinheim, Germany). Food samples were obtained from a local supermarket. General Experimental Procedure. One-dimensional and twodimensional (1D and 2D) NMR spectroscopy 1H, 1H−1H-gCOSY, gHSQC, gHMBC, and 13C were performed on an Avance III 500 MHz spectrometer with a CTCI probe or an Avance III 400 MHz spectrometer with a BBO probe (Bruker, Rheinstetten, Germany). Purity of all standards for external calibration was determined by means of qNMR after the method of Frank et al.2 Mass spectra of the compounds were measured on a Waters Synapt G2-S HDMS mass spectrometer (Waters, Manchester, UK) coupled to an Acquity UPLC core system (Waters, Milford, MA, USA). Chromophore screening and quantitation in cocoa powders were performed on a Xevo TQ-S mass spectrometer (Waters) coupled to an Acquity UPLC core system (Waters). HPLC analysis was performed using an analytical HPLC system (PU-2080 Plus; Jasco, Groß-Umstadt, Germany). HPLC separations were performed using a preparative HPLC system (PU-2087 Plus; Jasco). Solid phase extraction (SPE) for preseparation of model setups was performed using Chromabond C18 ec cartridges (70 mL, 10 g, Macherey Nagel, Düren, Germany). Debuffering of isolated compounds was performed by means of SPE using Strata C18-E cartridges (55 μm, 70 A, 1000 mg/6 mL, Phenomenex, Aschaffenburg, Germany) previously conditioned with

INTRODUCTION Recently, we investigated the formation of (+)-catechin- and (−)-epicatechin-derived polar chromophores in an alkaline milieu (model I) (accompanying article in this issue).1 The target compounds were isolated by means of preparative HPLC from model reactions imitating the cocoa alkalization process, and subsequently their structures were elucidated by means of UPLC-HR-ESI-TOF-MSe analysis, one- and twodimensional (1D and 2D) NMR spectroscopy, and electron paramagnetic resonance (EPR). With this strategy we found hydroxycatechinic acid radicals as contributors to cocoa reddening and identified the dehydrocatechinic acid−catechin dimer chromophores as new compounds.1 Therefore, the purpose of the present study was to characterize the unpolar chromophores derived from (+)-catechin or (−)-epicatechin alkaline model reactions using softer conditions as well as to quantify these compounds in order to estimate their contribution to cocoa color change during Dutching.

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MATERIALS AND METHODS

Chemicals. The following reagents were obtained commercially in p.A. quality: (+)-catechin (C), (−)-epicatechin (EC), ammonium formate (Sigma-Aldrich, Steinheim, Germany); potassium hydroxide, potassium carbonate, sodium hydroxide, iron(III) chloride, n-pentane (Merck, Darmstadt, Germany). Water for chromatographic separations was purified by means of a Milli-Q gradient Integral5 system (Millipore, Schwalbach, Germany), and solvents used were of HPLC© XXXX American Chemical Society

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February 18, 2019 April 8, 2019 April 8, 2019 April 9, 2019 DOI: 10.1021/acs.jafc.9b01050 J. Agric. Food Chem. XXXX, XXX, XXX−XXX

Article

Journal of Agricultural and Food Chemistry methanol and water. The sample was applied on the cartridge and flushed with water (10 × 5 mL); afterward, the buffer-free sample was eluted with methanol (2 × 5 mL). Absorbance maxima and corresponding extinction coefficients of identified chromophores were determined using a UV−vis 2401PC spectrophotometer (Shimazdu, Kyoto, Japan). Analytes were dissolved in ultrapure water or methanol, transferred to a quartz glass Suprasil cuvette (QS, 10 mm, 200−2500 nm, Carl Zeiss, Jena, Germany), and measured at the UV−vis range (200−650 nm) in reference to a blank of corresponding solvent. Model Reactions. The low alkalization model II was used for the generation of dehydrocatechin dimers: (+)-C (1 g) was suspended in an aqueous solution of potassium hydroxide (40 mL, 1 g/100 mL, pH 11), and oxygen was bubbled for 5 min via a semipermeable membrane into solution, and, afterward, heated for 2 h at 60 °C. After cooling with ice to ambient temperature, the pH was adjusted to 7 with formic acid (10%, aq). Then, the reaction mixture was preseparated by means of SPE. After activation and conditioning, elution was started with water (2 × 60 mL, aq fraction), followed by acetonitrile (2 × 60 mL, ACN fraction) and methanol (60 mL, MeOH fraction). The ACN fraction was further concentrated under reduced pressure to a total volume of 5 mL. An aliquot of this solution (1 mL) was diluted with methanol (1 mL) and acetonitrile (2 mL) and fractionated using preparative hydrophilic interaction liquid chromatography (HILIC) HPLC. The hydrogen peroxide alkalization model III was used for the generation of the hydroxyxanthene derivatives: (+)-catechin (1 g) was suspended in aqueous potassium hydroxide (40 mL, 1 g/100 mL, pH 11), hydrogen peroxide (500 μL, 30%) was added, and oxygen was bubbled for 10 min via a semipermeable membrane into solution. Afterward, the solution was heated for 2 h at 60 °C. Then, the reaction solution was immediately cooled with ice to ambient temperature, and the pH was adjusted to 7 with formic acid (10%). Afterward, the solution was concentrated under reduced pressure to a total volume of 5 mL. An aliquot of this solution (1.5 mL) was directly used for preparative RP-HPLC chromatography. Isolation and Structural Characterization of Target Chromophores. Further separation of the model approach II was performed using preparative HPLC using a Luna HILIC column (250 × 21.2 mm, 5 μm; Phenomenex) as the stationary phase. The effluent (21 mL/min) was monitored at 280 and 392 nm. The separation started with a mixture (A/B, 0/100, v/v) of aqueous ammonium formate (5 mmol/L, pH 5.8) as solvent A and aqueous ammonium formate in acetonitrile (10/90, v/v, 5 mmol/L, pH 5.8) as solvent B, and the B content was increased up to 90% within 10 min, afterward in 10 min to 70%, and in 2 min to 50% B. The compounds 1−6 were collected as a yellow-orange-colored dehydrocatechin dimer fraction (P2), C and EC (P1), orange dehydrocatechin trimers (P3), and oligomers (P4). The last eluting peak was collected as reddish-brown high molecular weight (HMW) fraction (P5). The dehydrocatechin dimer fraction was further purified by means of semipreparative HPLC. Chromatography was performed using a RP-column (250 × 10 mm, phenylhexyl, 5 μm; Phenomenex) as the stationary phase. The effluent (4.2 mL/min) was monitored at 280 and 392 nm. Using the solvents (A, B) mentioned above, the separation was started with a mixture (88/12, v/v) of A and B, and the B content was increased up to 30% within 30 min, and finally up to 35% within 8 min. Collected fractions were concentrated under reduced pressure, and buffer was removed by means of SPE using the method described above. The buffer free samples were concentrated under reduced pressure and freeze-dried, affording the dehydrodicatechin (dC) A isomers 1, 3, 5 and dehydrocatechin-epicatechin (dEC) isomers 2, 4, 6 (Figure 1). Separation of the model approach III was performed using preparative HPLC using a Luna phenyl-hexyl column (250 × 21.2 mm, 5 μm; Phenomenex) as the stationary phase. The effluent (21 mL/min) was monitored at 280 and 512 nm. The separation started with a mixture (A/B, 95/05, v/v) of aqueous ammonium formate (5 mmol/L, pH 5.8) as solvent A and aqueous ammonium formate in acetonitrile (10/90, v/v, 5 mmol/L, pH 5.8) as solvent B, and the B

Figure 1. Chemical structures of compounds 1−10. content was increased to 30% within 41 min and finally in 5 min to 100% B. Collected fractions were concentrated under reduced pressure, and buffer was removed by means of SPE as described above. The buffer free samples were concentrated under reduced pressure and freeze-dried, affording 7,8-xanthenocatechin (7,8-XC, 7), 7,8-xanthenoepicatechin (7,8-XEC, 8), 5,6-xanthenocatechin (5,6-XC, 9), and 5,6-xanthenoepicatechin (5,6-XEC, 10). B1′,D7,B3′,C3,B6′,D8-Dehydrodicatechin A (1, Figure 1). Yellow powder; UV (MeOH) λmax = 280, 380, 428 nm; ελmax (MeOH, 298 K, pH 7.6) = 7147 (380 nm), 2399 (428 nm) L·mol−1·cm−1; (−) HRESIMS: m/z = 575.1189 [M − H]− (calcd for C30H23O12, 575.1190). 1H NMR (500 MHz, acetonitrile-d3), COSY): δ 2.50 [m, 4H, J = 10.1, 8.9, 5.1 Hz, H-C (B2′α, C4α, F4 α)], 2.69 [m, 1H, J = 13.0 Hz, H-C(B2′β)], 2.88 [m, 1H, J = 11.0 Hz, H-C(C4β)], 4.04 [m, 2H, J = 9.2, 5.6, 3.9 Hz, H-C(C2, C3)], 4.09 [m, 1H, J = 8.1, 7.0 Hz, H-C(F3)], 4.90 [d, 1H, J = 6.7 Hz, H-C(F2)], 5.93 [m, 2H, J = 2.3, 2.2 Hz, H-C(A6, A8)], 6.20 [s, 1H, H-C(D6)], 6.34 [s, 1H, HC(B5′)], 6.76 [dd, 1H, J = 7.7, 2.1 Hz, H-C(E6′)], 6.82 [d, 1H, J = 6.0 Hz, H-C(E5′)], 6.85 [d, 1H, J = 1.8 Hz, H-C(E2′)]. 13C NMR (125 MHz, acetonitrile-d3, HSQC, HMBC): δ 27.5 [C-C4], 27.7 [CF4], 44.9 [C-B2′], 66.0 [C-C3], 67.2 [C-F3], 78.7 [C-C2], 83.0 [CF2], 89.5 [C-B1′], 91.0 [C-D6], 94.6 [C-B3′], 95.4 [C-A8], 96.4 [CA6], 98.5 [C-A4a], 103.7 [C-D4a], 105.5 [C-D8], 112.9 [C-B5′], 114.6/114.7 [C-E2′], 116.2 [C-E5′], 119.9/120.0 [C-E6′], 131.3 [CE1′], 145.6 [C-E3′/4′], 145.7 [C-E3′/4′], 154.5 [C-D8a], 155.4 [CA8a], 156.8 [C-A5], 157.5 [C-A7], 162.8 [C-B6′], 164.4 [C-D5], 166.9 [C-D7], 192.7 [C-B4′]. B1′,D7,B3′,C3,B6′,D8-Dehydrocatechin-(2S,3S)-epicatechin dimer (2, Figure 1). Yellow powder; UV (MeOH) λmax = 280, 393, 429 nm; ελmax (MeOH, 298 K, pH 7.6) = 1363 (393 nm), 821 (429 nm) L·mol−1·cm−1; (−) HRESIMS: m/z = 575.1189 [M − H]− (calcd for C30H23O12, 575.1190). 1H NMR (500 MHz, acetonitriled3), COSY): δ 2.51 [m, 2H, J = 12.5, 11.3 Hz, H−C(B2′α, C4α)], 2.69 [d, 1H, J = 12.0 Hz, H-C(B2′β)], 2.74 [m, 1H, H-C(F4α)], 2.87 B

DOI: 10.1021/acs.jafc.9b01050 J. Agric. Food Chem. XXXX, XXX, XXX−XXX

Article

Journal of Agricultural and Food Chemistry

H-C(2′)], 8.30 [s, 1H, H-C(9)]. 13C NMR (125 MHz, MeOD-d3, HSQC, HMBC): δ 27.8 [C-4], 67.6 [C-3], 83.7 [C-2], 95.9 [C-6], 103.1 [C-5′′], 107.3 [C-4a], 107.6 [C-2′′], 115.0 [C-2′], 116.3 [C5′], 118.6 [C-1′′], 119.8 [C-6′], 131.0 [C-1′], 134.8 [C-9], 146.5 [C3′/4′], 146.6 [C-3′/4′], 151.4 [C-3′′], 153.4 [C-8a], 154.0 [C-5], 154.9 [C-7], 158.1 [C-6′′], 176.1 [C-4′′]. (2S,3S)-7,8-Xanthenoepicatechin, (2S,3S)-2-(3,4-dihydroxyphenyl)-3,5,10-trihydroxy-3,4-dihydro-2H,9H-pyrano[2,3-a]xanthen-9-one (8, Figure 1). Red powder; UV (MeOH) λmax = 278, 449, 478, 514 nm, ελmax (MeOH, 298 K, pH 7.6) = 19439 (449 nm), 22070 (478 nm), 14267 (514 nm) L·mol−1·cm−1; (−) HRESIMS: m/z = 407.0764 [M − H]− (calcd for C22H15O8, 407.0767). 1H NMR (500 MHz, MeOD-d3, COSY): δ 2.90 [dd, 1H, J = 17.6 Hz, HC(4α)], 2.99 [dd, 1H, J = 17.4 Hz, H-C(4β)], 4.33 [m, 1H, J = 1.0 Hz, H-C(3)], 5.09 [d, 1H, J = 1.0 Hz, H-C(2)], 6.52 [s, 1H, HC(5′′)], 6.54 [s, 1H, H-C(6)], 6.87 [d, 1H, J = 8.2 Hz, H-C(5′)], 6.89 [s, 1H, H-C(2′′)], 6.90 [d, 1H, J = 7.0, 1.3 Hz, H-C(6′)], 7.09 [d, 1H, J = 1.3 Hz, H-C(2′)], 8.38 [s, 1H, H-C(9)]. 13C NMR (125 MHz, MeOD-d3, HSQC, HMBC): δ 29.0 [C-4], 66.4 [C-3], 81.2 [C2], 95.8 [C-6], 103.1 [C-5′′], 107.7 [C-4a], 107.9 [C-2′′], 114.9 [C5′], 115.4 [C-2′], 118.1 [C-1′′], 119.5 [C-6′], 131.2 [C-1′], 134.7 [C9], 146.2 [C-3′,4′], 151.1 [C-3′′], 153.0 [C-8a], 154.5 [C-5], 155.5 [C-7], 157.8 [C-6′′], 179.1 [C-4′′]. (2R,3S)-5,6-Xanthenocatechin, (2R,3S)-3-(3,4-dihydroxyphenyl)2,6,9-trihydroxy-2,3-dihydro-1H,10H-pyrano[2,3-c]xanthen-10-one (9, Figure 1). Red powder; UV (MeOH) λmax = 278, 451, 480, 509 nm, ελmax (MeOH, 298 K, pH 7.6) = 8161 (451 m), 9011 (480 nm), 6366 (509 nm) L·mol−1·cm−1; (−) HRESIMS: m/z = 407.0764 [M − H]− (calcd for C22H15O8, 407.0767).). 1H NMR (500 MHz, MeODd3, COSY): δ 2.66 [dd, 1H, J = 16.6, 7.4 Hz, H-C(4α)], 2.89 [dd, 1H, J = 16.7, 3.5 Hz, H-C(4β)], 4.17 [m, 1H, J = 12.6, 6.3 H-C(3)], 4.92 [d, 1H, J = 6.9 Hz, H-C(2)], 6.50 [s, 1H, H-C(5′′)], 6.52 [s, 1H, HC(8)], 6.76 [d, 1H, J = 7.9, 1.4 Hz, H-C(6′)], 6.80 [d, 1H, J = 8.1 Hz, H-C(5′)], 6.86 [m, 2H, H-C(2′,2′′)], 8.31 [s, 1H, H-C(9)]. 13C NMR (125 MHz, MeOD-d3, HSQC, HMBC): δ 27.6 [C-4], 67.6 [C3], 83.7 [C-2], 95.4 [C-8], 103.1 [C-5′′], 107.4 [C-4a], 106.9 [C-2′′], 115.0 [C-2′], 116.1 [C-5′], 118.2 [C-1′′], 119.8 [C-6′], 130.0 [C-1′], 134.8 [C-9], 145.8 [C-3′/4′], 146.6 [C-3′/4′], 151.2 [C-3′′], 153.2 [C-8a], 153.8 [C-5], 154.8 [C-7], 158.3 [C-6′′], 176.8 [C-4′′]. (2S,3S)-5,6-Xanthenoepicatechin, (2S,3S)-3-(3,4-dihydroxyphenyl)-2,6,9-trihydroxy-2,3-dihydro-1H,10H-pyrano[2,3-c]xanthen-10one (10, Figure 1). Red powder; UV (MeOH) λmax = 278, 451, 480, 509 nm, ελmax (MeOH, 298 K, pH 7.6) = 8161 (451 m), 9011 (480 nm), 6366 (509 nm) L· mol−1·cm−1; (−) HRESIMS: m/z = 407.0764 [M − H]− (calcd for C22H15O8, 407.0767).). 1H NMR (500 MHz, MeOD-d3, COSY): δ 2.89 [dd, 1H, J = 17.7, 4.4 Hz, H-C(4α)], 2.98 [dd, 1H, J = 17.7, 4.2 Hz, H-C(4β)], 4.33 [m, 1H, H-C(3)], 5.08 [d, 1H, J = 1.0 Hz, H-C(2)], 6.50 [s, 1H, H-C(5′′)], 6.52 [s, 1H, HC(8)], 6.83 [d, 1H, J = 8.0 Hz, H-C(5′)], 6.89 [d, 1H, H-C(2′′)], 6.90 [d, 1H, J = 8.6, 2.1 Hz, H-C(6′)], 7.09 [d, 1H, J = 2.0 Hz, HC(2′)], 8.36 [s, 1H, H-C(9)]. 13C NMR (125 MHz, MeOD-d3, HSQC, HMBC): δ 28.8 [C-4], 66.4 [C-3], 81.2 [C-2], 95.7 [C-8], 103.1 [C-5′′], 106.1 [C-6], 106.8 [C-4a], 107.8 [C-2′′], 115.4 [C-2′], 116.1 [C-5′], 118.2 [C-1′′], 119.5 [C-6′], 131.2 [C-1′], 134.8 [C-9], 145.8 [C-3′/4′], 146.2 [C-3′/4′], 151.0 [C-3′′], 153.5 [C-8a], 153.9 [C-5], 154.8 [C-7], 158.1 [C-6′′], 176.6 [C-4′′]. Quantitation of Compounds 1−8 in Alkalized Cocoa Powders. The target compounds were analyzed in the same manner as previously described.1 The MS/MS parameters of each compound were tuned using IntelliStart of MassLynx v4.1 software, and quantitative analysis was performed by means of the multiple reaction monitoring (MRM) mode with fixed mass transitions as follows: m/z 289.1 → 245.0 for (+)-C, (−)-EC and CA, m/z 575.1 → 394.1 for 1− 6, and m/z 407.1 → 255.0 for 7−10. All compounds were quantified by means of external calibration. dC and dEC dimers were quantitated as sum of all isomers according to the calibration curve of dC A.1 A defined amount of dC A was dissolved in methanol (1 mg/mL) and diluted with water to concentrations ranging from 1 mg/mL to 1 μg/ mL. After linear regression of the peak area versus concentration, calibration curves showed linear responses with correlation

[m, 1H, H-C(F4β)], 2.93 [m, 1H, H-C(C4β)], 4.06 [m, 2H, J = 9.9, 6.6, 1.8 Hz, H-C(C2,C3)], 4.27 [m, 1H, J = 0.9 Hz H-C(F3)], 5.05 [d, 1H, J = 0.9 Hz, H-C(F2)], 5.25 [s, 1H, HO-C(B3′)], 5.58 [d, 1H, J = 2.0 Hz, H-C(A8)], 5.94 [d, 1H, J = 2.0 Hz, H-C(A6)], 6.21 [s, 1H, H-C(D6)], 6.48 [s, 1H, H-C(B5′)], 6.84 [m, 1H, H-C(E5′)], 6.86 [m, 1H, H-C(E6′)], 7.00 [d, 1H, J = 1.5 Hz, H-C(E2′)]. 13C NMR (125 MHz, acetonitrile-d3, HSQC, HMBC): δ 27.7 [C-C4], 28.9 [C-F4], 45.0 [C-B2′], 65.9 [C-F3], 66.0 [C-C3], 78.7 [C-C2], 80.1 [C-F2], 89.4 [C-B1′], 90.9 [C-D8], 94.6 [C-B3′], 95.7 [C-A8], 97.0 [C-A6], 100.7 [C-A4a], 103.1 [C-D4a], 105.7 [C-D8], 112.5 [CB5′], 114.6 [C-E2′], 116.0 [C-E5′], 119.3 [C-E6′], 131.3 [C-E1′], 145.2 [C-E3′/4′], 145.5 [C-E3′/4′], 155.2 [C-D8a], 156.1 [C-A8a], 156.8 [C-A5], 157.7 [C-A7], 162.9 [C-B6′], 165.0 [C-D5], 166.8 [CD7], 192.6 [C-B4′]. B1′,D5,B3′,C3,B6′,D6-Dehydrodicatechin A (3, Figure 1). B1′,D7,B3′,C3,B6′,D6-Dehydrodicatechin A (5, Figure 1). Yellow powder; UV (MeOH) λmax = 280, 385, 428 nm; ελmax (MeOH, 298 K, pH 7.6) = 7147 (380 nm), 2399 (428 nm) L·mol−1·cm−1; (−) HRESIMS: m/z = 575.1189 [M − H]− (calcd for C30H23O12, 575.1190). 1H NMR (500 MHz, acetonitrile-d3), COSY): δ 2.23 [m, 1H, H−C(B2′α)], 2.50 [m, 1H, J = 10.1 Hz, H−C(F4α)], 2.58 [d, 1H, J = 4.6 Hz, H-C(C4α)], 2.69 [m, 1H, H-C(C4β)], 2.78 [m, 2H, J = 5.9, 3.8, H-C(F4α,β)], 2.88 [d, 1H, J = 11.0, H-C(B2′β)], 3.91 [d, 1H, J = 0.9 Hz, H-C(C2)], 4.09 [m, 1H, J = 8.1, 6.2 Hz, H-C(F3)], 4.40 [d, 1H, J = 0.9 Hz, H-C(C3)], 4.90 [d, 1H, J = 6.4, H-C(F2)], 5.54 [d, 1H, J = 2.3, H-C(A8)], 5.92 [m, 1H, J = 2.3 Hz, H-C(A6)], 6.23 [s, 1H, H-C(D8)], 6.37 [s, 1H, H-C(B5′)], 6.76 [dd, 1H, J = 7.7, 2.1 Hz, H-C(E6′)], 6.82 [d, 1H, J = 6.0 Hz, H-C(E5′)], 6.85 [d, 1H, J = 1.8 Hz, H-C(E2′)]. 13C NMR (125 MHz, acetonitrile-d3, HSQC, HMBC): δ 24.5 [C-C4], 27.4 [C-F4], 40.0 [C-B2′], 64.2 [C-C3], 67.2 [C-F3], 72.3 [C-C2], 83.0 [C-F2], 91.5 [C-B1′], 91.9 [C-D8], 94.8 [C-B3′], 95.7 [C-A8], 97.0 [C-A6], 100.62 [C-A4a], 104.2 [CD4a], 104.6 [C-D6], 113.9 [C-B5′], 114.6/114.7 [C-E2′], 116.2 [CE5′], 119.9/120.0 [C-E6′], 131.3 [C-E1′], 145.6 [C-E3′/4′], 145.7 [C-E3′/4′], 155.0 [C-D8a], 155.8 [C-A8a], 157.0 [C-A5], 157.4 [CA7], 163.2 [C-B6′], 164.5 [C-D5], 165.5 [C-D7], 192.5 [C-B4′]. B1′,D5,B3′,C3,B6′,D6-Dehydrocatechin-(2S,3S)-epicatechin Dimer (4, Figure 1). B1′,D7,B3′,C3,B6′,D6-Dehydrocatechin(2S,3S)-epicatechin dimer (6, Figure 1). Yellow powder; UV (MeOH) λmax = 280, 393, 429 nm; ελmax (MeOH, 298 K, pH 7.6) = 1363 (393 nm), 821 (429 nm) L·mol−1·cm−1; (−) HRESIMS: m/z = 575.1189 [M − H]− (calcd for C30H23O12, 575.1190). 1H NMR (500 MHz, MeOD-d3), COSY): δ 2.20 [m, 1H, J = 11.0 Hz, HC(B2′α)], 2.67 [m, 1H, J = 17.4, 5.2 Hz, H-C(C4α)], 2.83 [d, 1H, J = 16.7, 11.3, 5.5, 4.6 Hz, H-C(F4α, C4β)], 2.91 [m, 1H, J = 17.1, 9.1, 4.6, H-C(F4β)], 2.96 [d, 1H, J = 11.2, H-C(B2′β)], 3.86 [d, 1H, J = 1.5 Hz, H-C(C2)], 4.31 [m, 1H, J = 1.2 Hz, H-C(F3)], 4.49 [d, 1H, J = 5.0, 1.5 Hz, H-C(C3)], 5.07 [d, 1H, J = 1.2, H-C(F2)], 5.93 [d, 1H, J = 2.2, H-C(A6)], 5.93 [d, 1H, J = 2.0 Hz, H-C(A8)], 6.16 [s, 1H, HC(D8)], 6.56 [s, 1H, H-C(B5′)], 6.83 [d, 1H, J = 8.0 Hz, H-C(E5′)], 6.88 [dd, 1H, J = 7.9, 1.5 Hz, H-C(E6′)], 7.05 [d, 1H, J = 1.9 Hz, HC(E2′)]. 13C NMR (125 MHz, MeOD-d3, HSQC, HMBC): δ 24.7 [C-C4], 29.3 [C-F4], 40.8 [C-B2′], 64.9 [C-C3], 66.3 [C-F3], 72.8 [C-C2], 80.7 [C-F2], 91.6 [C-B1′], 91.8 [C-D8], 95.2 [C-A8], 95.4 [C-B3′], 96.4 [C-A6], 98.7 [C-A4a], 103.7 [C-D4a], 104.3 [C-D6], 112.8 [C-B5′], 114.9 [C-E2′], 115.9 [C-E5′], 119.0 [C-E6′], 131.1 [C-E1′], 146.0 [C-E3′/4′], 155.5 [C-A8a], 156.0 [C-D8a], 157.4 [CA5], 157.7 [C-A7], 164.6 [C-B6′], 166.5 [C-D7], 193.4 [C-B4′], not detectable [C-D5]. (2R,3S)-7,8-Xanthenocatechin, (2R,3S)-2-(3,4-dihydroxyphenyl)3,5,10-trihydroxy-3,4-dihydro-2H,9H-pyrano[2,3-a]xanthen-9-one (7, Figure 1). Red powder; UV (MeOH) λmax = 278, 449, 478, 514 nm, ελmax (MeOH, 298 K, pH 7.6) = 19439 (449 nm), 22070 (478 nm), 14267 (514 nm) L·mol−1·cm−1; (−) HRESIMS: m/z = 407.0764 [M − H]− (calcd for C22H15O8, 407.0767). 1H NMR (500 MHz, MeODd3, COSY): δ 2.66 [dd, 1H, J = 16.6, 6.8 Hz, H-C(4α)], 2.90 [dd, 1H, J = 16.3, 4.2 Hz, H-C(4β)], 4.16 [m, 1H, J = 7.1, 5.1, 2.0 Hz, HC(3)], 4.92 [d, 1H, J = 6.2 Hz, H-C(2)], 6.50 [s, 1H, H-C(5′′)], 6.51 [s, 1H, H-C(6)], 6.76 [d, 1H, J = 8.3, 1.8 Hz, H-C(6′)], 6.80 [d, 1H, J = 8.1 Hz, H-C(5′)], 6.85 [s, 1H, H-C(2′′)], 6.87 [d, 1H, J = 1.8 Hz, C

DOI: 10.1021/acs.jafc.9b01050 J. Agric. Food Chem. XXXX, XXX, XXX−XXX

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Journal of Agricultural and Food Chemistry

Figure 2. UPLC-ESIneg-TOF-MSe spectra of isolated dehydrodicatechin fractions (P2e-g). coefficients of R2 > 0.99. All calibration curves were generated in duplicates.

■

S2) containing dC dimers (1−6) as confirmed by means of HR-UPLC-TOF-ESI-MSe (Figure S3). 1−6 were obtained as yellow amorphous powders with typical absorption maxima expected for dehydrodicatechins. Results from negative electrospray ionization indicated for 1−6 an [M − H]− ion with m/z 575, as well as a fragment ion with m/z 394, typical for dC A.3 High resolution LC-MS analysis confirmed the target compounds to have a molecular formula of C30H23O12 and the fingerprint fragment C21H14O8 (Figure S3). The UPLC-TOF-MSe chromatogram showed two main peaks for P2e as well as P2g indicating the same molecular mass as well as fragmentation pattern, and therefore, revealed two isomers of dC A (Figure 2). 1H and 13C NMR chemical shifts of P2f only revealed one compound which was proposed as a further isomer of P2g. However, it was not possible to further separate the isomers of P2e and P2g by means of HPLC. First, the 1D and 2D NMR spectra of 1−6 were recorded in MeOD-d3 according to Guyot et al.3 However, the vanishing of acid proton signals could be observed over time, e.g., the indispensable proton signals H-C(6,8) for the linkage confirmation of the two monomers. Consequently, the spectra of P2e and g were also recorded in acetonitrile-d3. The 13C NMR signals of P2e could be assigned to a total of 60 carbon

RESULTS AND DISCUSSION

Besides the previously described polar chromophores from (+)-C and (−)-EC, a range of unpolar chromophores could be observed in alkalized cocoa powders as well (Supporting Information, Figure S1). Reaction parameter optimization for the formation of these unpolar chromophores yielded a softer alkalization process, and, therefore in models II (O2, compounds 1−6) and III (O2 + H2O2, compounds 7−10) with reduced concentrations of alkalizing agent as well as a decreased reaction temperature, but longer reaction times up to 2 h. The dehydrodicatechins (1−6) could be detected in the acetonitrile fraction of the SPE and were further separated via HILIC HPLC into five different fractions (P1−P5). LC-TOFMS screenings indicated the presence of dC oligomers in HPLC fractions P3 and P4 and confirmed dC dimers in fraction P2.3−5 HPLC fraction P5 was determined as a high molecular weight (HMW) fraction above 30 kDa by means of ultrafiltration. The target fraction P2 was further separated by means of RP-HPLC yielding three main peaks (P2e-g, Figure D

DOI: 10.1021/acs.jafc.9b01050 J. Agric. Food Chem. XXXX, XXX, XXX−XXX

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Journal of Agricultural and Food Chemistry

Figure 3. Excerpt of the HMBC spectrum (500/125 MHz, 298 K, MeOD-d4) of P2e. Crucial correlations for confirmation of the C-linkages are labeled black for 1 and dashed for 3/5.

target peak and P2g two more target peaks differing to P2f. These observations strengthened the theory of the existence of three constitutional isomers in total as well as it showed that the compounds found in P2f and g were further reaction/ degradation products of 1, 3, and 5. In summary, 3 and 5 could be assigned to B1′,D5,B3′,C3,B6′,D6-dehydrodicatechin A (3, Figure 1) and B1′,D7,B3′,C3,B6′,D6-dehydrodicatechin A (5, Figure 1). According to the UPLC-TOF-MSe chromatogram, P2f only consisted of one dC dimer, namely, B1′,D5,B3′,C3,B6′,D6- or B1′,D7,B3′,C3,B6′,D6-dehydrocatechin-(2S,3S)-epicatechin dimer (4/6, Figure 1). Furthermore, the 1H NMR spectrum of P2f revealed one set of signals with almost identical chemical shifts as the identified 6-C-linked dC A (3/5) isomers. In comparison to 3/5 the 1H NMR spectrum showed a downfield shift for H-C(F2) to 5.07 ppm with a coupling constant of J < 1 Hz. Also the aromatic proton H-C(E2′) appeared in a more downfield range at 7.05 ppm, which is the characteristic chemical shift of the corresponding EC proton H-C(2′). These signals clearly confirmed an EC moiety instead of a C moiety in the chemical structure. The HMBC experiment revealed only one correlation between H-C(B2′) and a carbonyl group at the typical chemical shift of 193.4 ppm, confirming the hemiacetal structure of P2f, alongside with the assignment of C(B1′) at 91.6 ppm and C(B3′) at 95.4 ppm by correlations with H-C(B2′) and H-C(B5′), respectively. The intramolecular C-linkage was determined at position 6 by comparison of the chemical shift of proton H-C(D8) of 3 and 5 with the corresponding aromatic proton of P2f showing the exact same chemical shift of 6.16 ppm. The corresponding HMBC correlation between H-C(D8) and C(D7) at 166.5 ppm also fitted to the 6-C-linked dC A (1). Furthermore, the chemical shifts of methylene protons H-C(B2′) resonating at 2.20 and 2.96 ppm exactly matched the corresponding proton signals of the 6-C-linked dC A isomer. As described above, the LC-MS chromatogram of P2g as well as the 1H NMR spectrum showed two more dC dimers with a double set of signals with similar chemical shifts of the identified dCs (1, 3, 5). Therefore, these two sets of signals were assigned to the B1′,D7,B3′,C3,B6′,D8-dehydrocatechin-

atoms which confirmed that the spectrum showed signals of two different isomers. The 1H and 13C NMR signal shifts of one isomer were completely in line with 8-C-dC A, and consequently 1 could be identified as B1′,D7,B3′,C3,B6′,D8dehydrodicatechin A.3 The set of signals of the second isomer was very similar to dC A (1) but showing slightly different chemical shifts. Therefore, it was suspected that both isomers only differed in the C-linkage. Confirmation of the C-linkages for both isomers was achieved by means of an HMBC experiment showing correlations between proton H-C(D6/8) and carbon atoms C(D5) and C(D7) resonating at 164.4 and 164.5 ppm and the corresponding correlation between protons H-C(F4) and C(D5) (Figure 3). The proton signal of the first isomer (1) resonating at 6.11 ppm showed two correlation signals at 165.0 and 166.7 ppm as well as the correlation signal of protons H-C(F4) to C(D5) at 165.0 ppm. The proton signal of the second isomer (3/5) resonating at 6.17 ppm showed only one corresponding correlation at 165.3 ppm and the correlation between protons H-C(F4) and C(D5) at 165.5 ppm confirming only a correlation to C(D7) for the aromatic proton H-C(D6/8) of 3/5. Vice versa for 3/5 a correlation of H-C(D6/8) to C(D5) was not detectable. In summary, dC A (1) showed the intramolecular C-linkage at position D8 and (3/5) at position D6 (Figure 3). Previously Guyot et al. confirmed the formation of the 8-Clinked dC A (1) by means of enzymatic catalysis and postulated the reaction pathway of dC dimers.3,7 In the present study we demonstrated the chemically formation of 6-C-linked isomers of dC A (3, 5) under alkaline conditions. Further, the position of the oxygen linkage to C(1′B) of 6-C-linked isomers was still unclear. Theoretically the formation of the new furan cycle was either possible from the hydroxy group D5 (3) or D7 (5). However, according to the recorded data, it was not possible to give a final conclusion if one of both constitutions was preferred or even that both constitution isomers coexist. A deeper insight gave the UPLCTOF-MSe chromatogram of the isolated peaks P2e, f, and g recorded after 1 week storage in methanol (Figure 2). The chromatogram showed a transformation of P2e to P2f and g structures as well as vice versa. Furthermore, P2f showed one E

DOI: 10.1021/acs.jafc.9b01050 J. Agric. Food Chem. XXXX, XXX, XXX−XXX

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Journal of Agricultural and Food Chemistry

Figure 4. UPLC-ESIneg-TOF-MSe spectrum of 7. (A) High and (B) low collision energy spectrum.

C14H7O5 (Figure 4). The 1H NMR spectrum of 7 confirmed the existence of a catechin moiety in the structure. Of the two expected catechin A-ring proton signals, only one could be assigned at 6.51 ppm. The missing of one A-ring proton signal gave a hint for a newly developed intramolecular C−C-linkage at either position C(6) or C(8). The 1H NMR spectrum further revealed two additional, noncatechin correlated, proton signals at 6.50 and 8.30 ppm. Furthermore, the HSQC spectrum revealed a second proton-carbon correlation at 6.85 ppm. The three additional proton signals highlighted an olefinic or aromatic π-electron system. A comparison of the 13 C NMR spectrum, in which 22 signals appeared, with the HSQC experiment showing 10 signals, revealed 12 signals corresponding to quaternary carbon atoms. Unequivocal assignment of these quaternary carbon atoms and the hydrogen-substituted carbon atoms, respectively, was successfully achieved by means of HMBC and HSQC experiments. The HMBC experiment revealed a correlation between the aromatic A-ring proton H-C(6) resonating at 6.51 ppm and C(4a) at 107.2 ppm as well as between C(4a) and protons HC(3) and H-C(4) indicating the position of the aromatic proton at C(6). In addition, the newly formed intramolecular C-linkage at the catechin A-ring was clearly confirmed between C(8) and C(9), by correlations of the protons H-C(4), HC(2), and H-C(9) to carbon atom C(8a) resonating at 153.4 ppm and no correlation with proton H-C(6), respectively. The two new carbon atoms C(2′′) and C(5′′) were assigned by the couplings to proton H−C(9). Further, the HMBC experiment revealed a correlation between proton H-C(5′′) resonating at 6.50 ppm and C(1′′) at 118.6 ppm. The carbon atom C(6′′) resonating at 158.1 ppm was assigned by correlations to the protons H-C(9), H-C(2′′) and H-C(5′′). The missing of a corresponding proton signal for carbon atom C(6′′) was explained by the fact that a new O-linkage between carbon atoms C(7) and C(6′′) has been established which could be confirmed by the downfield shift of this carbon atom to 158.1 ppm close to the chemical shift of equivalent carbon atoms C(3′) and C(4′). The carbon atom C(3′′) resonating at 151.4 ppm was assigned by correlations to the protons H-C(2′′) and H-C(5′′). The remaining signal of C(4′′) resonating at 176.1 ppm was identified as carbonyl group by its chemical shift and

(2S,3S)-epicatechin dimer (2) and the second 6-C-linked dC A isomer (4/6, Figure 1). Again, the 1H NMR proton signals of H-C(F2) and H-C(E2′) showed a downfield shift confirming EC moieties for both isomers. Furthermore, one set of signals showed identical chemical shifts as P2f for protons H-C(D8) resonating at 6.16 ppm and H-C(B5′) at 6.56 ppm indicating a 6-C-linkage. In comparison to 1 the chemical shift of H-C(D6) at 6.21 ppm of the second isomer matched to an 8-C-linkage. Additionally, the HMBC experiment revealed a correlation between the methylene proton H-C(B2′) resonating at 2.69 ppm and the carbonyl group C(B4′) at 192.6 ppm, similar to the chemical shift of proton H-C(B2′) and carbon atom C(B4′) of 1. The proton signals H-C(F4) resonating at 2.84 and 2.93 ppm showed a correlation with C(D5) at 169.0 ppm. As well as proton H-C(D6) resonating at 6.09 ppm showed correlations with C(D5) and C(D7) clearly confirming the 8C-linkage for the main isomer of P2g. For the second isomer, no correlations in methanol could be detected. However, the chemical shifts of proton signals H-C(B2′) perfectly fitted with the ones of P2f, clearly confirming the second isomer to be 6C-linked. The hemiacetal structure at position C(B3′) could be confirmed by the HMBC correlations in acetonitrile between the hydroxy-group proton HO-C(B3′) at 5.25 ppm and carbon atoms C(B2′), C(B3′) and C(B4′). In summary, the identification of the three dEC dimers detected in fractions P2f and g revealed the presence of one 8C-linked (2) and two 6-C-linked (4, 6) isomers. The dEC dimers have been previously described as natural occurring chromophores, e.g., in black tea.5,6 To the best of our knowledge, this is the first time NMR data are described for 6C-linked dC A and dEC dimers. These results give a deeper insight into the complexity, diversity, and formation of these compounds in processed food. Compounds 7−10 were isolated from model III and showed typical absorption maxima for red chromophores at 480−514 nm. Results from ESI MS indicated that these type of compounds form an [M − H]− ion with m/z 407, as well as a fragment ion with m/z 255, characteristic for a retro-Diels− Alder fragmentation of the catechin moiety. High resolution LC-MS analysis confirmed the target compounds to have a molecular formula of C22H16O8 and the fingerprint fragment F

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noticeable in a red coloring and a shift to higher wavelengths, which reflects the pH of cocoa powder suspensions after alkalization.11 After storage of compound 7 for at least of 3 days, a new set of signals was detectable which were assigned to the structure of 8 and confirmed an epimerization reaction of the carbon C(2) of the C moiety over time (Figure S8). The 1D/2D NMR data of compound 8 was very similar to 7 aside from differences in chemical shifts of corresponding EC signals. Furthermore, proton H-C(2) showed a smaller coupling constant (J = 1.0 Hz), comparable to the corresponding coupling constants of EC. Also the olefinic proton signal HC(9) appeared more down field at 8.38 ppm. Again, confirmation of the intramolecular linkage was proven by HMBC correlations, and, finally, compound 8 was identified as the epimer of 7, namely C-8-linked hydroxyxanthenon-EC. The 1 and 2D NMR data of 9 isolated from the second eluting peak 2 of model III was also very similar to 7. The HMBC correlations between proton H-C(9) and carbon atoms C(5,7) as well as between H-C(8) and C(8a), clearly confirmed the C-linkage at position 6. Also the aliphatic proton signals H-C(4) and H-C(2) (J = 6.9 Hz) assigned by HMBC correlations to C(8a) highlighted a C moiety. In conclusion, 9 was identified as the C-6-linked hydroxyxanthenon-C constitution isomer of 7. Analogous to 7 the NMR spectra of 9 revealed a second set of signals which was assigned to 10. The 1D/2D NMR data of 10 were very similar to 9 aside from differences in chemical shifts of protons H-C(2), H-C(9), and H-C(2′) underlying an EC moiety. The C-linkage of 10 was elucidated by the same strategy like for 9 and hence identified as the epimer of 9, namely, C-6-linked hydroxyxanthenon-EC. The O-linkage at position C(5) was highlighted by comparison of the chemical shifts of C(5) and C(7). C(7) resonated downfield shifted confirming a free hydroxy group according to NMR data of similar compounds.15 Therefore, compounds 9 and 10 were confirmed as O-5/O-6′′,C-6/C-9-XC and -EC, respectively.

HMBC correlation to proton H-C(2′′) (Figure 5). In conclusion, compound 7 was identified as an 8-C-linked

Figure 5. Excerpt of the HMBC spectrum (500/125 MHz, 298 K, MeOD-d4) of 7 showing key correlations for the structure elucidation.

hydroxyxanthenon derivative of C. Previously, Yanase et al.8 described a very similar xanthylium-related red pigment with similar NMR data, confirming the same unexpected low chemical shift for proton H-C(9) and a C-8-linkage for this compound. Es-Safi et al.9−11 showed that the absorbance maxima, and therefore, coloring of C-derived xanthylium salts are strongly pH dependent. Similar to anthocyanidins these compounds show a shift in absorption maxima as expressed in color shift from yellow in acidic milieu, to red in neutral and slight alkaline milieu and to reddish-pink in strong alkaline milieu. Xanthylium salts derived of C have been previously described as yellowish pigments occurring in white wine.9,10,12−14 It was possible to confirm the presence of the neutral state of the xanthene-pyran body at pH 6.5−7.5,

Figure 6. Proposed reaction pathway of 7−10 starting from (+)-catechin. G

DOI: 10.1021/acs.jafc.9b01050 J. Agric. Food Chem. XXXX, XXX, XXX−XXX

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Journal of Agricultural and Food Chemistry In summary, the identified compounds 7−10 were determined as C or EC, derivatives containing a hydroxyxanthenone body with a strong dislocated π-electron system causing the reddish coloring of these compounds, and therefore, potentially contribute to cocoa powder reddening. The proposed reaction pathway of these compounds starts with a dimerization reaction of C attacking from position C(8) or C(6), respectively, to the diarylpropanoyl-intermediate of C (Figure 6). Afterward, the A-ring of the diarylpropanoyl moiety is oxidized and a new pyran ring is formed by a nucleophilic attack of C(7) or C(5) hydroxy group to generate the C-6linked isomer. After the alkaline catalyzed elimination of the corresponding aldehyde residue, the nonoxidized xanthene intermediate is formed and via oxidation, and cleavage of two protons 7 and 9 are generated and eventually epimerized to 8 and 10. Quantitation of Compounds 1−10 in Alkalized Cocoa Powder Samples. Very recently, we traced the degradation of the major precursors (−)-EC and (+)-C and the previously not identified dehydrocatechinic acid-catechin chromophores by means of cocoa processing followed by LC-MS/MS analysis.1 Similar to EC, the dEC dimers decreased via fermentation in cocoa beans as well as liquor of 230−238 μmol/kg to 76−161 μmol/kg (Figure 7) and further via roasting. However, lowest concentrations were detectable in the alkalized samples ranging from 25 in light to 11 μmol/kg in strong alkalized cocoa. Similar to C, the dC showed a degradation from 37 to 12 μmol/kg by means of a 5 day fermentation, and a further decrease was observable via roasting. However, via alkalization the dC showed constant concentrations due to the increasing epimerization of dEC to dC (Figure 7A). Calculated ratios of dEC/dC showed similar results as previously stated for EC/C ratios and therefore confirmed the increasing epimerization rate of dEC to dC.1 The raw cocoa samples showed dEC/dC ratios of 5.8−7.3, the 5 day fermented samples showed a slight decrease of 5.6−6.2, the roasted showed an overall decrease up to 3.3−4.0 in R and 1.7−2.1 in HR samples as well as the ratios from 1.4 (weak) to 1.1 (medium), and 0.9 (strong) in alkalized samples (Figure 7B). By means of UPLC-MS/MS, it was possible to distinguish between dEC and dC C-8- and C-6-linked isomers. Thus, a constant decrease of the 8-C-linked isomers, occurring in unfermented cocoa beans, by means of cocoa processing was observable. The 6-C-linked dehydrocatechin dimers could only be detected in traces in the roasted samples, and especially, in alkalized cocoa powders, whereas from light to heavy Dutching an increase in the range of 7−15 (6-C-dEC) and 6−8 μmol/kg (6-C-dC) was detectable (Figure 7A,C). The XCs (7−10) were quantified in total as the chromatography was hampered, most probably caused by the keto−enol tautomery. In Dutched cocoa powders, the XCs showed an increase from 38 in weak to 108 μmol/kg in strongly alkalized cocoa powders, strengthen the postulated reaction pathway under alkaline conditions and also potentially contribute to the red coloring of alkalized cocoa powders (Figure 7D). In summary, the constitutional isomers dCs 1−6 as well as four isomers of the so far unknown reddish-colored XCs 7−10 could be isolated from different alkaline model reactions. Their chemical structures were determined by means of LC-MS, 1D and 2D NMR spectroscopy and their concentrations were quantitated in various differently processed cocoa samples (fermentation, roasting, Dutching). With the discovery of the

Figure 7. Quantitation of (A) 6-C- and 8-C-linked dehydrocatechin A dimers and dehydrocatechin-epicatechin dimers, (B) ratio shifts of dEC to dC, (C) ratio shifts of 8-C-linked- to 6-C-linked-dEC/dC and (D) quantitation of xanthenocatechins.

xanthene-derived red chomophores in alkalized cocoa, a second class of red, polyphenol-derived chromophores was identified alongside the previously described hydroxycatechinic acid radicals.1 With increasing grade of alkalization, an increase of XC content was detectable, which strongly supports the idea of the contribution of these compounds to cocoa reddening. Furthermore, the naturally occurring 8-C-linked dC dimers showed a constant decrease via cocoa processing. However, the 6-C-linked dCs, which were not detectable in nonprocessed cocoa at all, showed an increase in concentration, and therefore, seem to be marker compounds for the Dutch processing.

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ASSOCIATED CONTENT

S Supporting Information *

The Supporting Information is available free of charge on the ACS Publications website at DOI: 10.1021/acs.jafc.9b01050. Analytical, preparative HPLC separations, mass and HMBC spectra of compounds 3−10 (PDF) H

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(15) Morimoto, S.; Nonaka, G.; Nishioka, I. Tannins and related compounds. LIX. Aesculitannins, novel proanthocyandins with doubly-bonded structures from Aesculus hippocastanum L. Chem. Pharm. Bull. 1987, 35, 4717−4729.

AUTHOR INFORMATION

Corresponding Author

*Phone: +49-8161-71-2902. Fax: +49-8161-71-2949. E-mail: [email protected]. ORCID

Timo D. Stark: 0000-0002-6502-173X Thomas Hofmann: 0000-0003-4057-7165 Notes

The authors declare no competing financial interest.

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ACKNOWLEDGMENTS We thank the NMR team of Lehrstuhl für Lebensmittelchemie und Molekulare Sensorik for performing the NMR measurements. We thank Cargill for providing the alkalized cocoa samples and especially Brian Guthrie, Ph.D. from Cargill for excellent assistance, discussions, and cooperation on this project.

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REFERENCES

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