Article pubs.acs.org/JAFC

Quantitation of Chlorophylls and 22 of Their Colored Degradation Products in Culinary Aromatic Herbs by HPLC-DAD-MS and Correlation with Color Changes During the Dehydration Process Jean-Louis Lafeuille,* Stéphane Lefèvre, and Julie Lebuhotel EMEA Centre of Analytical Sciences, McCormick France, 999 Avenue des Marchés, 84200 Carpentras, France ABSTRACT: Chlorophylls and their green and olive-brown derivatives were successfully separated from culinary herb extracts by HPLC with photodiode-array and mass spectrometry detection. The method involved a ternary gradient elution and reversephase separation conditions capable of resolving 24 different pigments (2 chlorophylls and 22 of their derivatives) of different polarities within 28 min. The method was applied to monitor color changes in 50 samples of culinary aromatic herbs subjected to five different drying treatments. Of the 24 pigments, 14 were key to understanding the differences between the primary degradation pathways of chlorophyll a and chlorophyll b in culinary herbs during drying processes. A color degradation ladder based on the total molar percentage of all the remaining green pigments was also proposed as a tool to measure the impact of drying treatments on aromatic herb visual aspects. KEYWORDS: chlorophyll, pheophytin, pyrochlorophyll, pyropheophytin, culinary herb, drying



INTRODUCTION Chlorophylls are fat-soluble pigments and are responsible for the photosynthetic process and green hues in plants. They are present in the leaf chloroplasts,1 which are the entities implementing the energy-producing photosynthetic process using sunlight, carbon dioxide, and water. Chlorophylls (Figure 1) are

Most of the plants only have chlorophylls as photosynthetic green pigments when alive. Once they are harvested, protein denaturation occurs. Chlorophylls are then susceptible to chemical changes following multiple breakdown pathways according to the different process steps the plant is subjected to.2 There are some breakdown pathways leading to colored derivatives meaning their highly conjugated porphyrin ring remains unchanged. These colored derivatives are supposed to represent intermediary products of chlorophyll degradation before turning to discolored products by bleaching reactions.3 These reactions cleave the porphyrin ring by oxidation (photooxydation, dioxigenase, oxidase, peroxidase, lipoxygenase) producing fluorescent catabolites.4−6 Our study focuses only chlorophylls and their green and olive-brown colored degradation products.2,7 With regard to chlorophyll breakdown into colored derivatives, the first type of degradation is due to the asymmetry at carbon C132; chlorophylls a and b can turn into their respective epimers a′ and b′ even under mild conditions of processing. Other degradation pathways result from the presence of chemical constituents on the chlorophyll molecule periphery but not directly attached to the porphyrin ring. These groups can change or be removed, leading to green derivatives with the same chlorophyll visible spectrum. One of the groups is phytol at C17 (Figure 1) which can be removed by the action of an enzyme namely chlorophyllase naturally present in plant leaves.8 This hydrolytic reaction can also occur with other colored derivatives such as pheophytin (chlorophyll devoid of chelated Mg) to produce the corresponding dephytylated pigment namely pheophorbide,9 pyrochlorophyll to produce pyrochlorphillide, and pyropheophytin to produce pyropheophorbide. The phytol

Figure 1. Structure of chlorophyll a (R = CH3) and chlorophyll b (R = CHO).

composed of 4 pyrroles linked by 4 methyne bridges to form a porphyrin ring which is chelated with a magnesium ion placed in the center. As displayed in Figure 1, there are a phytol group esterified with propionate at C17, a keto group at C131, and a carboxymethoxy group at C132 which is an asymmetric carbon in the chlorophyll molecule. The most widely distributed forms in terrestrial plants are chlorophyll a and b having, respectively, a methyl group and a formaldehyde group at C7. © 2014 American Chemical Society

Received: Revised: Accepted: Published: 1926

August 22, 2013 January 28, 2014 February 1, 2014 February 1, 2014 dx.doi.org/10.1021/jf4054947 | J. Agric. Food Chem. 2014, 62, 1926−1935

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designed an interesting color degradation ladder enabling us to classify drying processes according to their impact on culinary herb surface color.

group is a long nonpolar alkyl group, and all the derivatives that lose it form a polar, colored chlorophyll derivative family and a suffix (-ide) is added to their names. Some processes call for a high-temperature blanching pretreatment (dipped in boiling water for a few seconds), which deactivates the chlorophyllase enzyme;10 thus, chlorophyllides can no longer be formed from chlorophylls. The loss of the carboxymethoxy group at C132 is due to a decarboxymethylation reaction that happens during severe heat treatment such as canning.11 This reaction eliminates the asymmetry at C132, and thus, no a′ and b′ derivative epimers can be formed. All the derivatives losing their carboxymethoxy group get a prefix (pyro-) added to their names. Another colorrelated transformation is the removal of chelated magnesium from the center of the porphyrin ring leading to derivatives with an olive-brown pigmentation. When plants are harvested leaf cell membranes break allowing sap acidic compounds to interact with chlorophylls. Acid conditions promote the loss of Mg2+ from the porphyrin ring, converting chlorophylls into olive-brown pheophytins. This conversion is the most frequent chlorophyll degradation, giving the product a brownish-greying surface color. Pheophytins can be formed through both dry-heating and moistheating processes.7 All the derivatives losing their chelating Mg2+ get a prefix (pheo-) added to their names. Finally, the formation of hydroxylchlorophylls and hydroxypheophytins caused by oxidation at C132 has been described in analytical studies on traditional Chinese herbs as well as other colored chlorophylls allomers like lactone derivatives.12−14 Chlorophylls and their derivatives have been extensively studied for both their implication in the color preservation in plants and for their reported biological activities.15 Surface color is an important quality attribute of herbal products; that is why numerous studies have been conducted to investigate color changes or chlorophyll degradation during various processes such as blanching, drying, or heat-treatment. The bright green color of chlorophylls is more pleasing to the consumer than the olive-brown color brought on by certain derivatives. Browning and greying of culinary herbs during drying processes is the most obvious visible symptom. A challenge to culinary herb processors is to preserve the natural color associated with quality attributes in a particular herb. Many publications in food process engineering and related fields deal with processes such as blanching, heat-treatment, or cooking that keep the plant wet at the end. However, culinary aromatic herbs such as oregano, basil, or dill are generally sold in a dried form to inhibit the growth of microorganism at ambient temperature. Freeze-drying, oven-drying, and sun-drying are the processes commonly used in the aromatic herb industry, but they must be used carefully so as to preserve as much of both the aroma and the color of the (fresh) raw material as possible. International standards for the quality specifications, including moisture content, of most culinary aromatic herbs are available, but none mention color specifications. Therefore, the drying treatment step is crucial to preserve the color quality of culinary herbs. In this Article we focused our attention on chlorophylls and their colored derivatives (both polar and nonpolar) present in dried culinary herbs. The first objectives were to detect and quantitate the largest range of chlorophyll colored derivatives possible. From this work we developed a knowledge and understanding of the various degradation pathways responsible for color changes in the dried herbs. With these findings our next goal was to find a novel way of determining the global level of dried culinary herb quality in terms of color. We thus



MATERIALS AND METHODS

Materials. Fresh spinach was obtained from a local store in Avignon. No commercial frozen spinach was used, because it is generally subjected to a blanching step leading to important epimerization of chlorophylls.7 Culinary aromatic herbs were cultivated and dried in Turkey and Egypt. Fresh culinary herbs were obtained from fields in east-southern France and kept between 2 and 8 °C prior to analysis. Chemicals. Analytical grade (acetone) and HPLC grade solvents (methanol, acetone, water) as well as all analytical grade chemicals (ammonium acetate, sodium citrate, hydrochloric acid, sodium hydroxide) were obtained from Sigma-Aldrich (St. Louis, MO). Nylon filters were obtained from Macherey-Nagel (Hoerdt, France). Instrumentation. The HPLC system consisted of an Alltech Elite degassing system (Nicholasville, KY), a Thermo SpectraSystem P1000 pump (San Jose, CA), a Thermo Spectra Series AS100 autosampler equipped with a 20-μL loop, and a Kinetex stainless-steel column (100 × 4.6 mm i.d.) packed with XB-C18 (2.6-μm) from Phenomenex (Torrance, CA). A Thermo SpectraSystem UV6000LP diode array detector (DAD) was used to both monitor the separation and determine the absorption spectra of chlorophylls and their derivatives. Spectral data from the DAD system were recorded between 350 and 700 nm. For mass spectra, the HPLC described above was equipped with an AB-Sciex 4000 Q-trap mass spectrometer (MS) (Foster City, CA) with an electrospray ionization source. A UV-1800 spectrophotometer from Shimadzu (Kyoto, Japan) was used. An IKA T25 Ultra-Turrax homogenizer (Staufen, Germany) equipped with a stainless steel disperser (18 mm in diameter) and a Waring Lab blender (Winsted, CT) equipped with 1-L stainless-steel container were used to grind plant samples in solvents. A Retsch MM400 benchtop mixer mill (Haan, Germany) was used to grind dry plants. A Minolta CR310 colorimeter (Osaka, Japan) was used to measure in the LCh° colorimetric space. Data reported are the average of 3 determinations. Xlstat version 2012.4.03 from Addinsoft (Paris, France) was used for processing principal components analyses (PCA) and principal component regression (PCR). Dehydration Processes. Several dehydration processes were used: sun-drying as the commonly used approach, freeze-drying as the reference in term of preserving green color, and last oven-drying. The latter two methods were used as control treatments. Two other proprietary types of dehydration processes were used and were named DP1 and DP2. DP1 is an innovation brought to the traditional sundrying process. DP2 is a recently developed process designed to preserve the green appearance of aromatic herbs. Preparation of Standard Solutions. Chlorophyll a and b standard solutions were extracted and prepared from fresh spinach leaves. The chlorophyll derivatives comprising the pheophorbides and pyropheophorbides, although available commercially,16 were synthesized as deemed necessary. Approximately 54 g of fresh spinach leaves were weighed into a 1-L container to which 200 mL of acetone was added. The mixture was homogenized for 1 min. Part of the extract was passed through a 0.45-μm nylon filter. The ratio of 54 g of spinach−200 mL acetone was prepared to obtain a final chlorophyll solution composed of 80:20 acetone−water. This solution was subjected to a 10-fold dilution with acetone−water 80:20 and used as the standard chlorophyll solution for realizing regression curves for external calibration. A pheophytin standard solution was prepared by acidification of the chlorophyll standard solution according to Schwartz et al.17 Hydrochloric acid (1 N) was added dropwise to 100 mL of the chlorophyll standard solution, and the solution was shaken after each addition. The complete conversion was reached once the color of the solution turned 1927

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from green to olive brown. The absence of remaining chlorophylls could be confirmed by HPLC. Chlorophyllides were prepared in situ by the action of chlorophyllase enzyme on chlorophylls as substrate.10 Fresh spinach leaves were gently dehydrated for 48 h in an oven at 40 °C and finely ground with a mixer mill. This resulting powder contained both enzyme and substrate (the substrate was the chlorophyll already present in the spinach leaves). A 500 mg portion of the dried spinach leaf powder and 10 mL of a 50:50 acetone−aqueous citrate solution (0.04 M) were added. The mixture stood for 2 h at ambient temperature and was then passed through a 0.45-μm nylon filter. Pheophorbides were obtained by using a chlorophyllase enzyme solution described below on the pheophytins substrate. Around 5 g of dry spinach leaf powder, dried according to the procedure above for chlorophyllides, was weighed on a glass Buchner filtration system with a round paper filter. All the glassware as well as the sample and 500 mL of acetone were kept in a freezer at −20 °C. Immediately after taking all the materials out of the freezer, the cold acetone was passed through the spinach leaf powder in the cold Büchner filtration system in order to remove the chlorophylls. Once the acetonic eluent became transparent, the sample was put in an oven at 37 °C for solvent evaporation. To 500 mg of the resulting powder was added 5 mL of neutralized standard pheophytin solution along with 5 mL of deionized water previously equilibrated to pH 8 with aqueous sodium hydroxide solution (0.1 N). Pyrochlorophylls were obtained from the chlorophyll standard solution. Briefly, 10 mL of the chlorophyll standard solution was extracted with diethyl ether and the ether extract evaporated to dryness. The residue was dissolved in 20 mL of pyridine and heated under reflux in an oil bath at 100 °C for 2 h.18 Pyridine was removed in a rotary evaporator, and 10 mL of an acetone−water 80:20 mixture was added. Pyropheophytins were prepared from the standard pheophytin solution using the same procedure with additional steps consisting of neutralization of the pheophytin solution with 0.1 M NaOH solution prior to evaporation. Pyropheophorbides were prepared from the pheophorbide solution using the protocol described above and time allotted to turn the chlorophylls into pyrochlorophylls. Finally, pyrochlorphillides were prepared from the chlorophillide solution using the same procedure described, but in this chemical preparation, no heat was necessary for the reaction to occur. To summarize, a solution with chlorophylls and all the synthesized derivatives was prepared. The more acidic solutions (i.e., pheophytins and pheophorbides) were neutralized by a 0.1 N NaOH solution so that the chlorophylls and other derivatives with chelated magnesium remained stable. All solutions prepared were kept at −20 °C. Extraction of Chlorophylls and Their Derivatives from Culinary Herb Samples. Approximately 1 g of fresh or dry herb was weighed into a 150 mL beaker to which 100 mL of an 80:20 acetone−sodium citrate solution (0.1M) was added. The mixture was homogenized for 8 min at medium speed. The homogenate was filtered on a nylon filter prior to chromatographic analysis. The mixture volume lost upon extraction was accurately measured for adjustment in the final calculations. A 10-fold dilution can be necessary for samples of fresh herbs or herbs subjected to efficient color-protective dehydration processes. Preparation of the Stock and Working Solutions for External Calibration. Chlorophyll and pheophytin concentrations in the stock solutions were determined according to Vernon.19 The pheophytin stock solution was neutralized with 0.1 M NaOH. Five working solutions containing chlorophylls and pheophytins (between 1 and 20 mg/L for chlorophyll a and pheophytin a) were prepared by mixing the stock solutions and dilution with acetone−water (80:20). The calibration curves were generated by plotting the five concentrations of each compound against peak area measured at 654 nm. A working solution was injected every five sample injections to check for drift effect on the calibration. Final results of pigment levels in herbs were expressed in mol/g on dry wt basis; therefore, the moisture content was measured for every sample. Analysis of Chlorophylls and Their Derivatives by HPLCDAD-MS. A solvent system of acetone−methanol−0.5 M NH4OAc

Table 1. Solvent Composition Evolution of the Elution Phase During the HPLC-DAD Analysis time

MeOH

ammonium acetate 0.5 M

acetone

flow (mL/min)

0 2 15 18 24 26 28 30

80 80 65 65 30 30 0 0

20 20 5 5 5 5 0 0

0 0 30 30 65 65 100 100

0.5 0.5 1 1.2 1.2 1.2 1.2 1.2

was used to separate chlorophylls and their breakdown compounds (see Table 1). The injection volume was 20 μL. Chlorophylls and their various derivatives were identified by retention time and by comparison of the absorption spectra between 350 and 700 nm of unknown peak with those in the literature, as well as with reference standards obtained by synthesis. In addition, the pigments were confirmed for their molecular weight ([M + H]+) by a quadrupole MS in positive mode with a 500−1000 Da scanning range. The source temperature was at 500 °C, ion spray voltage at 5500 V, curtain gas at 30 psi, declustering potential at 125 V, and the entrance potential at 10 V. Moisture Content Determination. For culinary aromatic herbs the recommended method to measure the moisture content is by distillation with toluene according to ISO 939:1980. Chlorophylls and Derivatives Breakdown: PCA and PCR Analyses. Using the experimental protocol described in this paper, chlorophylls and their derivatives were detected and quantified in 50 samples of diverse culinary herbs prepared fresh and after five different drying treatments described below. Herb samples included tarragon, coriander, parsley, dill, chive, oregano, mint, basil, and marjoram. All the samples were independent of each other. Drying treatments were the following: no process (fresh), sun-drying, freezedrying, oven-drying, DP1 and DP2. The data (average of duplicate analyses) was evaluated using principal component analysis (PCA) and principal component regression (PCR) techniques.



RESULTS AND DISCUSSION Extraction of Chlorophylls and Their Derivatives from Culinary Aromatic Herb Samples. An experimental design was carried out to optimize the parameters ruling the pigment extraction yields from fresh and dried herbs by means of an Ultra-Turrax homogenizer. The extraction mixture contained a 0.1 M sodium citrate solution to maintain a neutral pH while protecting the pigments against oxidation. The extraction time and the homogenizer rate of revolution were two key parameters. Other important factors were the extraction solution temperature and total molar contents of chlorophylls and their derivatives. The experimental design showed a strong interaction between the key parameters. For example, regardless the type of culinary herb or whether it was fresh or dry, the final optimized parameters were found to be an extraction time of 8 min at medium rotation speed. Preparation of the Stock and Working Solutions for External Calibration. Internal standards like Fast green FCF or zinc-phtalocyanine were used in some publications with varying levels of success.12,20 Chlorophyll and derivate quantitation was also realized using external calibration.7 The UV−vis absorption properties of chlorophylls and their derivatives can be attributed to three parameters: the primary porphyrin ring structure, the presence of chelated magnesium at its center, and the type of functional group or constituent at C7. With that in mind we could approximate to first order that both epimerization and decarboxymethylation at C132, and also 1928

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phytol loss at C17, do not affect the global UV−vis spectra of chlorophyll derivatives because C132 and C17 are not part of the porphyrin ring. Consequently, there are 4 types of UV−vis spectra: two groups for the green pigments (chlorophyll, chlorophyllide, pyrochlorophyllide, and pyrochlorophyll) according to the substituent at C7 (a or b) and likewise two groups for the olive-brown pigments (pheophytin, pheophorbide, pyropheophytin, and pyropheophorbide, a or b). This first approximation also takes into consideration the slight effect on the absorption spectra of the elution phase composition change throughout the chromatographic analysis. This effect is very low for the visible part of the spectrum, that is, why 654 nm was chosen as the quantitation wavelength for all green and olive-brown pigments. Furthermore, at this wavelength the carotenoid interferences are also eliminated. An external calibration method was used to quantitate each pigment. The two groups of green pigments were quantified using the calibration curve of chlorophyll a and b. Likewise, the 2 groups of olive-brown pigments were quantified using the calibration curves of pheophytin a and b. With working solutions containing known concentrations of chlorophylls a and b and pheophytins a and b, 5 injections were sufficient to cover the quantitation of all 24 chlorophylls and

Table 2. Analytical Parameters of Calibration Regression Curves chlorophyll a chlorophyll b pheophytin a pheophytin b conc range (mg/L) slope ± SD (n = 6) (1 × 103 AU)a r2 (av, n = 6) a

1−19

05−10

1−22

0.5−13

315 ± 10

139 ± 6

140 ± 7

69 ± 4

0.9993

0.9995

0.9997

0.9998

AU = area units.

derivatives. The calibration curves showed a high linearity, with a correlation coefficient ≥0.9993, as long as the concentration was below 25 mg/L (see Table 2). The final results of pigment contents in fresh or dried culinary herbs were expressed in mol/g on dry weight basis. In order to assess the global level of green pigment breakdown and to further estimate the visual color appearance of the herb product, the molar percentage of the green pigments was calculated (see equation below), with the remainder of the components representing the percentage of the olive-brown pigments.

b,b′

%green pigments = 100 ×

∑i = a , a ′ (chlorophylli + pyrochlorophylli + chlorophyllidei + pyrochlorophillidei) b,b′

∑i = a , a ′ (chlorophylli and derivatesi)

Analysis of Chlorophylls and Their Derivatives by HPLC-DAD-MS. In order to separate chlorophylls and their polar and nonpolar colored derivatives, a combination of two gradient mobile phases reported in previous studies was utilized.9,12 Polar derivatives were separated using a solvent system of methanol/aqueous ammonium acetate (0.5 M).9 These polar pigments are acidic pigments with pKa ranging from 2 to 4.21 Ammonium acetate was used in HPLC analysis because it provides excellent results in the separation of highly polar acidic chlorophyll derivatives. In addition, it buffers the elution phase at a pH around 7 and thus reduces the likelihood of porphyrin rings losing their chelating magnesium during analysis. Nonpolar derivatives were separated using a solvent system of methanol/acetone.12 Figure 2 shows the HPLC chromatogram of chlorophylls and their derivatives from a standard solution by employing a ternary solvent system of acetone/methanol/ammonium acetate (0.5 M). Following the described gradient program, good resolution of the chlorophyll derivative peaks was achieved. All the peaks showing chlorophyll-like or pheophytin-like spectra were checked for their molecular weight by LC-MS. With a single injection and a 28 min run, a total of 24 pigments could be detected. The retention time increased in the order presented in Figure 2 in agreement with results reported previously for chlorophillide/pheophorbide/chlorophyll/ pheophytin.22 We noticed different clusters of peaks on the pigment chromatogram. Each cluster contains the peak of the main pigment (chlorophyll or chlorophillide or pheophytin or pheophorbide, either a or b), the peak of its epimer form, and the peak of the -pyro form. The -pyro form elutes last in each cluster likely due to the elimination of the carbomethoxy group. This slightly reduces the polarity of the pigments resulting in greater interactions with the C18 stationary phase, as well as a lower solubility in the eluent.23 Therefore, the entire

chromatogram in Figure 2 shows eight clusters of 3 peaks that make the final identification easier. Chlorophylls and Derivatives Breakdown: PCA Analyses. In our study we decided to quantitate chlorophylls and their colored derivatives in fresh and dried culinary aromatic herbs subjected to different dehydration processes. There were 50 different herb samples. Of the 24 chlorophylls and their derivatives detected and identified by our method, only 14 of them were found in the different herb samples, namely the following: chlorophyll a and a′, chlorophyll b and b′, chlorophyllide a, pheophytin a and a′, pheophytin b and b′, pheophorbide a, pyrochlorophyll a and b, pyropheophytine a and b. No compounds were identified by MS as hydroxychlorophyll or hydroxypheophytin with a hydroxyl group at C132. The raw data are presented in Table 3, and they were used to build Table 4. Chlorophyllide a and pheophorbide a were rarely found in the 50 samples (only in samples 3 and 1, respectively). Chlorophillide a appeared after a low-temperature blanching step on fresh herbs prior to dehydration which allowed the chlorophyllase to be activated. Chlorophyll epimers were present in highest concentrations in the samples with ovenand sun-drying processes. Table 4 shows that chlorophyll a′/chlorophyll a ratio is generally lower than chlorophyll b′/chlorophyll b ratio. Similar results were reported previously in cooked wet spinach.20 In our study, these epimer ratios varied noticeably according to the type of treatment. Surprisingly, the pheophytin a′/pheophytin a ratio values were less dependent on the type of process used for drying the herbs. Pheophytin b′/pheophytin b ratio values (when detected) were high, showing that pheophytin b is subjected to a strong epimerization in every type of process. In the plants with less degraded chlorophylls (freeze-drying, DP1, and DP2 treatments), chlorophyll a was in higher amounts than chlorophyll b, which was also mentioned elsewhere.13 1929

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Figure 2. HPLC chromatogram of a mixture of chlorophylls and their colored derivatives. Chromatographic conditions described in the text. Peaks: 1a,b = chlorophyllide b,b′; 1c = pyrochlorophyllide b; 2a,b = chlorophyllide a,a′; 2c = pyrochlorophyllide a; 3a,b = pheophorbide b,b′; 3c = pyropheophorbide b; 4a,b = pheophorbide a,a′; 4c = pyropheophorbide a; 5a,b = chlorophyll b,b′; 5c = pyrochlorophyll b; 6a,b = chlorophyll a,a′; 6c = pyrochlorophyll a; 7a,b = pheophytin b,b′; 7c = pyropheophytin b; 8a,b = pheophytin a,a′; 8c = pyropheophytin a.

process leading to further pigment degradation toward the appearance of pyropheophytins, with pyrochlorophylls and pheophytins as potential precursors. In that case, pyrochlorophyll b turns into pyropheophytin b more rapidly whereas pyrochlorophyll a tends to accumulate. This means that pheophytin b tends to accumulate as well, allowing an important conversion into pheophytin b′ that is well-observed for herbs subjected to sun-drying processes (Table 3). However, the presence of pyropheophytin a in measurable amounts, especially in sun-dried herb samples, suggests that it might have been formed from pheophytin a. These findings demonstrate that pyropheophytins cannot only be formed under relative severe heat treatment as previously described,11 but also during long mild drying processes such as sun-drying. In conclusion, the total of pyroderivatives in the b series (pyrochlorophyll b + pyropheophytin b) is always higher than the total of pyroderivatives in the a series and contrary to what happens to chlorophylls a and b. This kinetic tendency in the pyroderivative formation has not been described elsewhere to date. Figure 3 attempts to describe the degradation pathways and the novel kinetic qualitative tendencies described above. Normally, the loss of Mg2+ ion is supposed to be the primary and direct way of chlorophyll breakdown, but in the case of dried aromatic herbs both Mg2+ removal and decarboxymethylation can be dominant reactions, depending on whether one considers the a or b chlorophyll form. The primary focus of literature references on chlorophyll degradation pathways includes wet plants (leaf senescence, cooking processes, fruit ripening) or vegetable oils.2−11,16,17,19,20 For dried culinary herbs, the dehydration process is done rapidly after harvesting, and thus,

Pyrochlorophyll b was found in most of the samples, even in fresh herb samples, and the only degradation pathway available is from chlorophyll b. Fresh plants were kept several days in a refrigerator prior to analysis, and the degradation process may have already started, which could explain the presence of pyrochlorophyll b. This phenomenon has not been described elsewhere. As for pyrochlorophyll a, it has not been detected in fresh herb samples, nor after the DP2 process, the quickest process. The presence of pheophytin b was not observed in fresh herbs or in DP2 processed samples. In the other processes, this pigment was not present all the time, except for samples subjected to sun-drying. Unlike pheophytin b, pheophytin a was detected in all 50 samples. It is well-known from numerous reports11 that chlorophyll a is more susceptible to forming pheophytin than chlorophyll b. This slower conversion of chlorophyll b to pheophytin presents two aspects: on one hand, after the long sun-drying process that produces the highest level of chlorophyll breakdown, the concentration of chlorophyll a became lower than that of chlorophyll b. On the other hand, it allows an easier conversion of chlorophyll b into pyrochlorophyll b through decarboxymethylation, which is thus the dominant breakdown reaction for chlorophyll b. This degradation pathway is confirmed by looking at the pyrochlorophyll b/chlorophyll b + b′ ratio (Table 4) which is much higher than the pyrochlorophyll a/chlorophyll a + a′ ratio, except in the sun-drying process. Pyrochlorophyll b content was generally found to be higher than that of pyrochlorophyll a, except in the sun-dried herb samples. This unique sun-drying exception can be explained: it is a long 1930

dx.doi.org/10.1021/jf4054947 | J. Agric. Food Chem. 2014, 62, 1926−1935

olive-brown pigments (1 × 10−9 mol/g dry weight)a

1931

4.1 4.2 4.3 4.4 4.5 4.6 4.7

5.1 5.2

mint 2 mint 3

3.0 0.6

772.7 204.0 808.0 826.9 761.3 813.6 516.0

433.2 24.1

3.4 3.5

dill 3 oregano 3 parsley 6 parsley 7 parsley 8 parsley 9 coriander leaves 4

77.4 11.3 124.5

3.1 3.2 3.3

oregano 1 oregano 2 marjoram 1 basil 2 marjoram 2

206.4 269.1 253.2

1.9 1.10 1.11

927.4 245.2 566.3 423.4 603.5 504.6

347.3 335.8

1.7 1.8

2.1 2.2 2.3 2.4 2.5 2.6

470.3 32.1 140.6 203.9 46.6 279.9

1.1 1.2 1.3 1.4 1.5 1.6

dill 2 tarragon 3 mint 1 basil 1 parsley 5 chive 2

parsley 1 tarragon 1 chive 1 dill 1 tarragon 2 coriander leaves 1 parsley 2 coriander leaves 2 parsley 3 parsley 4 coriander leaves 3

0.0 0.6

81.3 21.9 19.1 19.8 27.6 31.3 8.7

8.6 7.2

ND 0.3 1.9

86.7 15.6 33.8 19.9 27.6 16.8

51.7 64.7 66.2

90.9 66.3

88.4 9.1 33.0 52.1 12.3 68.2

52.1 19.1

263.3 103.6 334.4 362.7 329.3 355.4 272.0

219.0 32.4

47.1 10.3 67.7

374.9 101.4 243.6 204.8 265.2 242.3

85.6 108.4 96.2

179.0 145.8

165.2 15.1 108.2 124.7 26.9 148.0

14.1 5.1

47.0 17.5 19.9 19.4 22.6 23.1 1.7

10.7 10.3

2.0 0.4 2.2

67.8 18.8 25.7 23.8 42.1 16.0

31.6 38.7 28.8

54.6 39.0

44.7 18.7 31.8 39.3 9.7 47.8

ND ND

ND ND ND ND ND ND 0.4

ND ND

ND ND ND

ND ND ND ND ND 9.8

ND ND ND

ND ND

ND ND ND ND ND ND

0.7 6.6

ND ND ND ND ND ND 0.6

ND 0.1

1.4 ND ND

ND ND 1.2 1.3 ND ND

2.7 2.4 4.9

2.8 1.0

4.0 1.0 1.5 ND 1.4 2.3

61.7 97.8 71.4

203.5 90.3

1.4 1.9

9.7 ND 7.5 11.6 7.6 7.7 6.2

11.9 2.1

10.4 12.6 3.8 ND 4.3 5.3 2.7

36.1 14.9

Sun-Drying 199.6 88.5

6.8 13.0

6.5 1.6 0.7

7.6 1.2 10.3 8.2 19.4 21.5

11.5 22.9 14.8

41.5 19.3

11.0 1.3 44.5 32.8 2.4 35.7

37.5 65.1 18.2 16.0 24.3 36.1 18.6

DP2

42.1 65.6

Freeze-Drying 35.0 46.4 13.9 6.7 24.2 50.0 27.0 49.0 25.9 109.8 8.8 119.4 DP1 2.1 34.6 0.4 7.6 3.8 5.5

23.2 31.7 19.7

18.2 7.3

Oven-Drying 4.2 46.1 13.8 6.6 9.6 232.6 21.6 187.8 20.8 11.0 18.0 162.8

32.9 70.1

ND ND ND ND ND ND ND

1.9 8.7

2.2 2.6 ND

8.5 ND ND 20.1 19.2 10.5

ND 6.1 ND

ND ND

ND ND 17.8 6.8 6.3 10.8

20.1 59.2

ND ND ND ND ND ND ND

6.5 6.9

9.7 2.2 4.0

7.8 ND ND 17.1 17.6 11.0

ND 14.1 ND

ND ND

ND 3.9 18.4 21.6 5.7 25.5

12.3 16.0

13.3 ND ND ND ND 1.6 ND

0.8 2.0

0.4 ND ND

ND ND ND ND 3.4 2.0

3.0 9.6 8.0

19.3 ND

ND ND 7.5 5.8 4.6 14.5

4.8 9.8

ND ND ND ND ND ND ND

5.8 3.6

8.1 ND 3.3

ND ND ND ND ND ND

ND 13.3 ND

ND ND

ND 3.2 21.7 17.7 ND 17.7

ND ND

ND ND ND ND ND ND ND

ND ND

ND ND ND

ND ND ND ND ND 27.7

ND ND ND

ND ND

ND ND ND ND ND ND

0.06 0.03

2.94 1.97 2.42 2.28 2.31 2.29 1.90

1.98 0.74

1.64 1.10 1.84

2.47 2.42 2.32 2.07 2.28 2.08

2.41 2.48 2.63

1.94 2.30

2.85 2.12 1.30 1.63 1.74 1.89

18.9 11.6

95.0 81.7 98.2 98.7 97.6 96.6 97.4

91.4 43.3

67.9 61.8 93.6

95.5 98.0 93.7 88.1 85.1 80.6

84.0 75.9 83.3

72.4 84.4

93.2 85.7 48.7 61.8 79.7 67.9

type of herb and no. sample on chlorophyll chlorophyll chlorophyll chlorophyll chlorophyllide pyrochlorophyll pyrochlorophyll pheophytin pheophytin pheophytin pheophytin pyropheophytin pyropheophytin pheophorbide chloro % number PCA a a′ b b′ a a b a a′ b b′ a b a a/bb greenc

green pigments (1 × 10−9 mol/g dry weight)a

Table 3. Pigment Contents in the Culinary Aromatic Herb Samples

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1932

1.3 2.6 3.0 7.5 154.3 63.6

5.1

4.7 136.2

5.6 5.7 5.8 5.9 5.10 5.11

5.15

5.16 5.17

441.2 714.8 609.1 557.9 643.9 720.7 428.0

2.6 2.8

5.4 5.5

6.1 6.2 6.3 6.4 6.5 6.6 6.7

1.2

5.3

2.6 3.8 3.7 3.7 3.9 4.2 ND

0.5 5.4

0.5

0.5 4.1 1.0 1.6 1.9 9.9

0.6 0.6

0.3

175.0 322.5 266.2 252.6 272.6 326.4 181.0

48.1 108.8

50.1

10.0 30.7 15.7 21.0 128.3 32.1

25.4 24.5

22.5

3.1 3.1 3.8 1.6 3.1 2.9 ND

13.8 12.2

14.6

3.1 8.8 4.8 5.5 5.8 6.2

6.5 7.3

6.5

ND ND ND ND ND ND 94.1

ND ND

ND

ND ND ND ND ND ND

ND ND

ND

ND ND ND ND ND ND 2.8

5.6 1.2

3.4

0.2 1.5 0.3 0.4 1.1 0.4

8.6 0.3

3.5

199.4

70.8 196.9 132.0 155.3 94.9 23.5

190.2 103.7

Sun-Drying 108.9

220.4 164.5 Fresh Herb 3.6 7.3 2.8 13.5 10.0 11.8 11.6 9.7 3.8 10.3 18.4 14.7 1.8 7.0

2.8 4.7

2.7

0.7 1.4 1.0 1.6 5.9 1.9

4.0 1.0

0.8

ND ND ND 2.0 0.6 1.7 0.7

37.0 27.1

36.2

13.4 37.4 24.2 22.5 18.0 4.9

27.5 16.7

18.3

Mean of duplicate analyses. bChloro a/b = chlorphyll a/chlorophyll b. c% green, percentage of total green pigments.

oregano 8 parsley 10 parsley 11 parsley 12 parsley 13 parsley 14 coriander leaves 6

marjoram 3 basil 3 marjoram 4 oregano 4 oregano 5 oregano 6 basil 4 oregano 7 coriander leaves 5 marjoram 5 basil 5 chive 3

a

green pigments (1 × 10−9 mol/g dry weight)a

olive-brown pigments (1 × 10−9 mol/g dry weight)a

ND ND ND ND ND ND ND

52.1 10.2

32.2

33.7 94.9 75.6 83.1 5.7 6.3

84.3 43.3

49.6

ND ND ND ND ND ND ND

58.5 32.7

34.1

13.3 67.8 28.6 62.6 20.2 3.1

70.0 31.0

35.1

ND ND ND ND ND ND ND

17.4 5.8

6.8

42.1 7.0 128.0 13.3 0.9 1.7

10.9 5.7

10.6

ND ND ND ND ND ND ND

38.7 27.2

23.7

2.8 42.6 9.5 6.0 17.4 2.4

3.1 2.0

5.2

ND ND ND ND ND ND ND

ND ND

ND

ND ND ND ND ND ND

ND ND

ND

2.52 2.22 2.29 2.21 2.36 2.21 2.36

0.10 1.25

0.10

0.13 0.09 0.19 0.36 1.20 1.98

0.10 0.11

0.05

98.8 98.7 98.7 98.6 98.8 98.5 98.9

15.1 50.1

18.7

8.2 9.9 6.1 9.9 65.4 73.1

11.0 15.3

13.2

type of herb and no. sample on chlorophyll chlorophyll chlorophyll chlorophyll chlorophyllide pyrochlorophyll pyrochlorophyll pheophytin pheophytin pheophytin pheophytin pyropheophytin pyropheophytin pheophorbide chloro % number PCA a a′ b b′ a a b a a′ b b′ a b a a/bb greenc

Table 3. continued

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Table 4. Pigment Ratio Results (Means and Standard Deviations) process

n

oven-drying freeze-drying DP1 DP2 sun-drying fresh herb

11 6 5 7 14 7

a

% green pigmentsa 75.6 90.7 71.9 95.3 22.7 98.7

± ± ± ± ± ±

12.6 7.1 21.1 5.5 22.4 0.1

chlorophyll a′/ chlorophyll a 0.24 0.06 0.07 0.05 0.32 0.01

± ± ± ± ± ±

0.03 0.02 0.13 0.04 0.43 0.002

chlorophyll b′/ chlorophyll b 0.40 0.14 0.10 0.09 0.25 0.01

± ± ± ± ± ±

pheophytin a′/ pheophytin a

pheophytin b′/ pheophytin b

± ± ± ± ± ±

1.96 ± 1.0 0.93 ± 0.08 2.37 ± 1.8

0.28 0.05 0.12 0.06 0.08 0.01

0.21 0.18 0.18 0.16 0.17 0.07

0.02 0.01 0.03 0.08 0.02 0.08

1.10 ± 1.0

pyrochlorophyll a/ chlorophyll (a + a′) 0,01 0.0008 0.004 0.0002 0.95 0.0009

± ± ± ± ± ±

0.008 0.001 0.008 0.0004 1,6 0.003

pyrochlorophyll b/ chlorophyll (b + b′) 0.18 0.09 0.05 0.02 0.05 0.03

± ± ± ± ± ±

0.17 0.03 0.007 0.01 0.03 0.02

% green pigments = percentage of total green pigments.

maximum of information by converting a set of observations (herb samples) of possibly correlated variables (pigments) into a set of values of orthogonal variables called principal components. As explained above, the use of these variables did not aim to differentiate the types of dried herbs, but rather the types of processes they were subjected to. The first two components represent the plane of best fit through the data. As PCA is sensitive to the relative scaling of the original variables, observation values expressed in mol/g on dry wt basis were reduced and centered prior to calculations. In our case, there were 14 variables composed of the 14 detected chlorophylls and derivatives leading to a 14D space composed of the final orthogonal principal components. The 2D (F1, F2) principal plane with the orthogonal projections of the 14 initial variables as well as the scatterplot diagram with scores of the observations are displayed in Figure 4 showing an interesting cumulative variability: F1 and F2 axes account for 57% of the total variability, and the following principal components show Eigenvalues below 1.15. On principal plane with the variable projections (Figure 4A), most of the variables are away from the plane center meaning that they were not that distorted by the projection. Chlorophyllide a and pheophorbide a have a low contribution to F1 and F2, but they were rarely found in the 50 samples. Apart from pyrochlorophyll a, all the relevant variable projections related to green pigments are on the left half of the plane. All the relevant variable projections related to olive-brown pigments are on the right half. This suggests that globally green and olive-brown pigments are negatively correlated along the F1 axis. In practical terms, it reflects the fact that a drop in the green pigment concentrations leads to an increase of the brown-olive ones. As pyrochlorophyll a tends to accumulate, its behavior is like that of the olive-brown pigments. That is why its projection is atypically in the right half of the (F1, F2) plane. Chlorophyll epimer variables (a′ and b′) point upward on the principal plane suggesting that samples subjected to mild processes will be placed rather at the top of this plane. On the scatterplot (Figure 4B), observations coming from the same drying treatments were plotted so that they form clusters. As mentioned for the variable projection directions, fresh and freeze-dried plants, as well as the group of plants subjected to treatment DP2 (which is supposed to protect the green color effectively), are in the left half of the (F1, F2) plane considered as the region where green pigments are wellpreserved. Treatment DP1 and oven-drying are more central, and sun-drying on the right part is driven by the olive-brown pigment variables. The freeze-drying cluster extends toward the upper-left portion of the (F1, F2) plane in the direction of the pyrochlorophyll b projection. Consequently, this process produces the highest levels of pyrochlorophyll b.

Figure 3. Chlorophyll a and b degradation pathways in culinary aromatic herbs during drying processes. Arrow width is proportional to the qualitative importance of the degradation pathway kinetic.

the final water activity ranges from 0.3 to 0.55. This low range significantly reduces the effect of enzymatic reactions involved in Mg-dechelatase, chlorophyllase, or enzymatic reduction of chlorophyll b.24 Consequently, no chlorophyllide was detected in this study except in one product subjected to a blanching step prior to dehydration. Moreover, neither pyrochlorophyllides nor pyropheophorbides were detected, because both need the chlorophyllase action to occur somewhere in the chlorophyll degradation process. The chlorophyll catabolic pathway is stopped;24 a simpler chemical degradation occurs instead. The raw data in Tables 3 and 4 and illustrated pathways in Figure 3 show how different dry culinary herb species chosen independently of each other can show similar behavior with respect to noncatabolic chlorophyll degradation. In those conditions, the type of process became an important factor for differentiation. The raw data were evaluated by PCA, which is mainly a statistical treatment of the data. The objective was to present a 1933

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factors: cloudy or sunny conditions during drying, windy or calm, exposure to the sun light for long or short periods of time, high, low or moderate temperature fluctuations and, finally, whether or not a a pretedding step is included. The variability of these parameters can vary greatly and make the sun-drying cluster large. On the F1 axis representing the greenness scale, the DP2 process (high energy, short time) cluster is the closest to the fresh herb cluster and elongates toward the pyrochlorophyll b and chlorophyll epimer in direction. Very few olive-brown pigments are thus produced. Chlorophylls and Derivatives Breakdown: PCR Analyses. After the PCA treatment of the data we chose to extract quantitative information from the results. It was interesting to test if quantitative information could be forecasted for unknowns from the available measures. In that case, PCR (principal component regression) can help, since it calculates a single principal component regression model using the given number of principal components to predict unknown quantitative information (response) from available measurements (observations). The calculated orthogonal principal components from the PCA data were used to run the PCR model. Herb surface color was chosen as an interesting response for variables and especially the LCh° colorometric space according to three axes: lightness (L), chroma or saturation (C), and hue (h°). This colorometric space describes how well the human eye can perceive the different colors. Unfortunately, no significant correlation was found between the variables and the 3 colorimetric responses. A slight linearity was observed for the hue h° response. Additionally, the relationship between hue and the total green pigment content (on dry wt basis, mol/g) was plotted. Hue values ranged from 85° to 128°, which correspond to the yellow-green part of colorimetric space. The curve h° as a function of total green pigment content reached rapidly an asymptote at h° = 128° for contents above 4 × 10−7 mol/g. This interesting result shows that no surface color improvement can be realized by having greater concentrations of green pigments. It also tells us that below 4 × 10−7 mol/g the surface green color rapidly worsens. Color Degradation Ladder. In order to understand the tendency for chlorophyll to degrade toward olive-brown derivatives, a degradation ladder was designed. The idea was to find a sole and easy-to-use indicator using the complex set of chlorophyll and derivative results in order to assess the plant material color after processing. This ladder was thus based on the percentage of total green pigments present (presented earlier in this paper), while understanding that green chlorophyll derivatives do not bring about any change in color insofar as the light absorption properties remain globally the same. This ladder was divided into four categories. Category limits were fixed according to the visual color estimation of each sample. Figure 5 shows the ladder visually and allows one to check whether the processing and treatment of a herb has an impact on the visual quality of the final product when dried. The various herb process treatments from Table 4 were evaluated with the ladder definitions. Fresh herbs, freeze-dried herbs, and DP2 treatment (high energy short time) of herbs belong to the category “no significant impact” with percentage of total green pigments above 90% on average with a well-preserved green appearance of the dried herb samples. Oven-drying (high energy medium time) and DP1 treatment (low energy long time) belong to the “low impact” category, with a percentage ranging from 65% to 90%. Sun-drying (low energy long time) falls into the fourth

Figure 4. Projection of the pigment variables on (A) the principal (F1, F2) plane, and (B) scatterplot of the culinary herb samples on the first 2 principal component plane.

Oven-drying and sun-drying clusters are the most scattered groups of observations but elongate in different directions. For oven drying, higher energy is brought to the product in a medium period of time. High energy and medium time preserve relatively well the green color; that is why the cluster of oven-drying observations tends to extend toward the upper part of the (F1, F2) plane, the direction between the chlorophyll epimer and pheophytin projections. Thus, herbs subjected to this process showed that chlorophylls had been partly turned to both other green pigments and olive-brown pigments. The oven-drying cluster is scattered, presumably due to the impact of the oven temperature parameter on the color result. As for sun-drying, a process with low energy and long delay time impacting chlorophyll breakdown, samples are situated on the right part of the plane. That is the consequence of a massive chlorophyll conversion into olive-brown derivatives such as pheophytin a, pheophytin b and b′ and moderately pheophytin a′, pyropheophytin a and pyrochlorophylls. The efficiency of this process on color preservation depends on numerous 1934

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(14) Kao, T. H.; Chen, C. J.; Chen, B. H. An improved high performance liquid chromatography-photodiode array detectionatmospheric pressure chemical ionization-mass spectrometer method for determination of chlorophylls and their derivates in freeze-dried and hot-air-dried Rhinacanthus nasutus. Talanta 2011, 86, 349−355. (15) Lanfer-Marquez, U. M.; Barros, R. M. C.; Sinnecker, P. Antioxidant activity of chlorophylls and their derivatives. Food Res. Int. 2005, 38, 885−891. (16) Yamauchi, N.; Harada, K.; Watada, A. E. In vitro chlorophyll degradation in stored broccoli (Brassica oleracea L. var. italica Plen.) florets. Postharvest Biol. Technol. 1997, 12, 239−245. (17) Schwartz, S. J.; Woo, S. L.; von Elbe, J. H. High-performance liquid chromatography of chlorophylls and their derivatives in fresh and processed spinach. J. Agric. Food Chem. 1981, 29, 533−535. (18) Pennington, F. C.; Strain, H. H.; Svec, W. A.; Katz, J. J. Preparation and properties of pyrochlorophyll a, methyl pyrochlorophyllide a, pyropheophytin a, and methyl pyropheophorbide a derived from chlorophyll by decarbomethoxylation. J. Am. Chem. Soc. 1964, 86 (7), 1418−1426. (19) Vernon, L. P. Spectrophotometric determination of chlorophylls and pheophytins in plants extracts. Anal. Chem. 1960, 32 (9), 1144− 1150. (20) Bohn, T.; Walczyk, T. Determination of chlorophyll in plant samples by liquid chromatography using zinc−phthalocyanine as an internal standard. J. Chromatogr. A. 2004, 1024, 123−128. (21) Mantoura, R. F. C.; Llewellyn, C. A. The rapid determination of algal chlorophyll and carotenoid pigments and their breakdown products in natural waters by reverse-phase high-performance liquid chromatography. Anal. Chim. Acta 1983, 151, 297−314. (22) Canjura, F. L.; Schwartz, S. J. Separation of chlorophyll compounds and their polar derivatives by high-performance liquid chromatography. J. Agric. Food Chem. 1991, 39, 1102−1105. (23) Suzuki, N.; Saitoh, K.; Adachi, K. Reversed-phase highperformance thin-layer chromatography and column liquid chromatography of chlorophylls and their derivatives. J. Chromatogr. A. 1987, 408, 181−190. (24) Hörtensteiner, S.; Kräutler, B. Chlorophyll breakdown in higher plants. Biochim. Biophys. Acta 2011, 1807, 977−988.

Figure 5. Color degradation ladder of culinary aromatic herbs. The percentages are the level of total green pigments (figure colors are arbitrary).

category, named “important impact”, and characterized by an average percentage below 35%. Sun-drying is a commonly used process to dry herbs; however, the technique greatly impacts the global herb color.



AUTHOR INFORMATION

Corresponding Author

*Phone: +33 4 90 63 89 22. Fax: +33 4 90 63 89 82. E-mail: [email protected]. Notes

The authors declare no competing financial interest.



REFERENCES

(1) Matile, P.; Schellenberg, M.; Vicentini, F. Localization of chlorophyllase in the chloroplast envelope. Planta 1997, 201, 96−99. (2) Schwartz, S. J.; Lorenzo, T. V. Chlorophylls in foods. Food Sci. Nutr. 1990, 29 (1), 1−17. (3) Matile, P.; Hörtensteiner, S. Chlorophyll degradation. Annu. Rev. Plant Physiol. Plant Mol. Biol. 1999, 50, 67−95. (4) Ginsburg, S.; Matile, P. Identification of catabolites of chlorophyll-porphyrin in senescent rape cotyledons. Plant Physiol. 1993, 102, 521−527. (5) Marangoni, A. G. Kinetic model for chlorophyll degradation in green tissue. 2. Pheophorbide degradation to colorless compounds. J. Agric. Food Chem. 1996, 44, 3735−3740. (6) Yamauchi, Y.; Watada, A. E. Effectiveness of various phenolic compounds in degradation of chlorophyll by in vitro peroxydasehydrogen peroxide system. J. Jpn. Soc. Hortic. Sci. 1994, 63 (2), 439− 444. (7) Teng, S. S.; Chen, B. H. Formation of pyrochlorophylls and their derivatives in spinach leaves during heating. Food Chem. 1999, 65, 367−373. (8) Yamauchi, N.; Watada, A. E. Pigment changes in parsley leaves during storage in controlled or ethylene containing atmosphere. J. Food Sci. 1993, 58, 616−618. (9) Almela, L.; Fernandez-Lopez, J. A.; Roca, M. J. High-performance liquid chromatographic screening of chlorophyll derivatives produced during fruit storage. J. Chromatogr. A 2000, 870, 483−489. (10) Holden, M. The breakdown of chlorophyll by chlorophyllase. Biochem. J. 1961, 78, 359−364. (11) Schwartz, S. J.; Lorenzo, T. V. Chlorophyll stability during continuous aseptic processing and storage. J. Food Sci. 1991, 56 (4), 1059−1062. (12) Shioi, Y.; Tomita, N.; Tsuchiya, T.; Takamiya, K. Conversion of chlorophyllide to pheophorbide by Mg-dechelating substance in extracts of Chenopodium album. Plant Physiol. Biochem. 1996, 34, 41−47. (13) Huang, S. C.; Hung, C. F.; Wu, B. H.; Chen, B. H. Determination of chlorophylls and their derivatives in Gynostemma pentaphyllum Makino by liquid chromatography−mass spectrometry. J. Pharm. Biomed. Anal. 2008, 48 (1), 105−112. 1935

dx.doi.org/10.1021/jf4054947 | J. Agric. Food Chem. 2014, 62, 1926−1935

Quantitation of chlorophylls and 22 of their colored degradation products in culinary aromatic herbs by HPLC-DAD-MS and correlation with color changes during the dehydration process.

Chlorophylls and their green and olive-brown derivatives were successfully separated from culinary herb extracts by HPLC with photodiode-array and mas...
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