1. Introduction
Mung bean (Vigna radiata L.), otherwise known as Phaseolus aureus or green gram, is a popular edible legume consumed worldwide [1]. It is widely grown in Asia, Africa, Europe, and Australia, but India, China, Pakistan, and Thailand are among the most important producers [2]. Therefore, it is a major commercial crop, and in the first two quarters of 2021, US imports of mung beans increased by 62% over 2020, mainly from China. Although India and Myanmar accounted for 30% of global output each, in 2020, Australia’s mung bean production, with 125,000 hectares planted, was fully mechanized and exported to Asia, North America, Europe, and the Middle East [3].
In brief, mung bean is consumed in various ways depending on the culinary culture or region. It can be consumed directly as cereal or processed to produce fresh bean sprouts, flour, soup, porridge, or transparent noodles made of starch or used as mung bean paste in sweets and cakes. Mung bean is estimated to provide a good amount of carbohydrates, proteins, minerals (potassium, iron, phosphorus, and calcium), isoflavonoid, folate, and dietary fiber [4]. Moreover, it effectively improves metabolic disorders, has an unsaturated nature of lipid composition and has a significant insoluble fiber fraction. Its phenolic content is related to its antioxidant activity [5]. Studies have shown that mung bean polysaccharides provide immunomodulatory activity and anti-tumor and anti-radiation health-promoting effects [6].
Mung bean consists of the seed coat, cotyledon, and hilum. The source of color is primarily from three parts: the surface texture layer and two parts from the seed coat. Pigments in the textural layer primarily consist of brown pigments; the seed coat has an opaque appearance when pigments are absent. In the seed coat, anthocyanins are distributed in the columnar layer, generating black spots, whereas green pigments in the parenchymal layer primarily consist of chlorophyll, without which the seed coat has a yellow color [7]. Chlorophyll is known to absorb light in the violet to red spectrum (approximately 400 to 700 nm wavelength range) and reflects green light (500 to 570 nm), which causes the characteristic green color of land plants. This is crucial for oxygen-containing photosynthesis. Nearly all plants contain chlorophyll a and b, whereas algae contain chlorophyll c, d, and e. Moreover, photosynthetic bacterial species have bacteriochlorophyll [8].
The basic unit of chlorophyll is the porphyrin ring, which is composed of four pyrrole rings linked into a large ring with a magnesium (Mg) atom in the center [9]. The pyrrole ring has special side chains, one of which is called phytol, a long chain. Under the effect of hydrochloric acid in the chlorophyll center, the Mg will be replaced by hydrogen to form pheophytin. Moreover, heating will accelerate the reaction. If heated continuously, the pheophytin will lose its methyl methacrylate to form pyropheophytin, thus causing the solution to turn brown. According to the chemical nature of chlorophyll, under the appropriate conditions (controlling the acidity and temperature of the solution, etc.), the Mg atom in the chlorophyll molecule can be replaced by copper, zinc, iron, and other metal elements [10,11]. Other derivatives such as chlorophyll and pheophorbide show opposite properties because of the loss of the phytol long chain. Chlorophyll degradation occurs primarily due to intrinsic factors such as leaf aging and fruit ripening or extrinsic factors such as pathogen invasion, temperature, lighting, oxygen, and pH. It occurs in the chloroplast matrix. First, chlorophyll is catalyzed by chlorophyllase to produce chlorophyllide; then, Mg is eliminated to form pheophorbide, following which it enters the cytosol to continue metabolism [12]. Therefore, the preservation of chlorophyll is intimately related to the quality of green plants. When the foods are processed thermally, chlorophyllase will be activated at 60 to 82 °C, and chlorophyll will turn into chlorophyllide, which is more stable to thermal treatment than chlorophyll [9]. Therefore, a plant can obtain better green color if treated with high-temperature sterilization.
Studies indicate that green fruits and vegetables show better and more stable colors when treated with alkaline conditions [8,9]. For enzymatic and non-enzymatic browning during processing or storage, hot water, and steam blanching (requiring prevention of overheating) before processing (e.g., canning or freezing) of fruits and vegetables to improve product quality is well established. A temperature spread of 60 to 100 °C for 30 to 150 sec effectively reduces the residual peroxidase activity in corn samples [13]. For this procedure, reducing the microbial load during mid-processing and cold storage of corn also preserved the product quality (color and vitamin retention) in further processing and storage. However, dehydration by drying decreases the enzyme levels without deactivating them; moreover, these drying methods cause severe damage to the nutritional value of the product (loss of vitamin C), as well as loss of color, appearance, and taste of the product, which may lead to hardening of the tissue, thus losing taste with the long drying time and high temperature [14,15].
Encouragingly, the global food industry focuses on innovative technologies to deactivate enzymes (by 90%, possibly more) to minimize adverse effects on texture, flavor, and nutrition. No study has examined the color change of mung beans during thermal processing. Hence, this study aimed to investigate the critical causes of color variation in mung beans during thermal processing, along with the differences within the pigments of mung bean cotyledons and seed coats. Finally, this study analyzed the chlorophyll changes in the seed coat of mung beans, revealing the critical causes of color changes after thermal processing.
2. Materials and Methods
2.1. Materials
Mung beans (Vigna radiata L. (Wilczek) cv. Tainan No. 5) were randomly harvested from around 10 mung bean plants from a local farm in Agricultural Research and Extension Station (Tainan District, Taiwan) and dried at 40 °C in an air-conveyor oven. The dried mung beans were vacuum-packed and stored at 4 °C until use. Methanol and ethyl acetate (BDH®), were purchased from Avantor® (Radnor, PA, USA). The standards (Sigma-Aldrich®, Kanagawa Prefecture, Japan) of chlorophyll a and b were purchased from Merck KGaA (Darmstadt, Germany). The other chemicals used were analytical-grade reagents.
2.2. Thermal Treatment of Mung Beans
Whole mung beans, seed coats, and cotyledons were evaluated. We took 5 g of whole beans and manually separated the seed coat and cotyledons. Reverse osmosis water was added 20-fold to each group according to dry weight. The samples were prepared by heating at 120 °C for 10–50 min. After cooling and centrifuging the sample at 4 °C for 10 min (Hettich Mikro 22R, Hitachi High-Tech Science, Tokyo, Japan), and the pH value of the supernatants, which were stored at 4 °C, was measured. Moreover, the bottom solids were freeze-dried (Eyela VOS301SD, Tokyo Rikakikai, Tokyo, Japan).
2.3. Characterization of Physicochemical Properties
Measurements of the content of moisture (Method 44-15A), protein (Method 46-11A), ash (Method 08-01), and lipid (Method 30-10) of mung bean seed coat and cotyledons were performed according to the American Association for Clinical Chemistry method [16]. The amylose content was determined by a rapid colorimetric method [17,18].
2.4. Color Characteristics
Color changes of samples during thermal processing were monitored for Hunter’s L (brightness), a (redness-greenness), and b (yellowness-blueness) values determined by using a colorimeter (Color Meter ZE-2000, Nippon Denshoku Industries, Tokyo, Japan).
2.5. Measurement of Browning Index
The browning index was determined in the supernatant as described previously [19,20], with modifications. Each group of the cooking solution was filtered, and 15 mL was taken for centrifugation (4 °C, 1000× g, 15 min). Then, 3 mL of supernatant was mixed with the same amount of 95% ethanol in an ice bath for 15 min and again centrifuged, with the absorbance value determined from the supernatant. Pigments from the Maillard reaction have extreme absorbance values at 420 nm; thus, the extent of the browning index can be determined from the 420 nm value by UV-visible absorption spectrometry (Spectrophotometer, U-2000, Hitachi High-Tech Science, Tokyo, Japan).
2.6. Determination of Total Flavonoids and Total Polyphenols
The methods for the determination of total flavonoids and total polyphenols were modified from those of other studies [21,22,23]. The assessed mung bean samples were whole mung bean, seed coat, and cotyledon. The sample was mixed with 2% of AlCl3·6H2O by vibration and was allowed to stand for 10 min at room temperature. The absorbance value at 430 nm was measured by a spectrophotometer. The absorbance values of the test samples were substituted into the regression equation of the standard curve (0–100 μg/mg of quercetin solution was used as the standard), and the total flavonoid content was obtained by conversion. The sample (400 μL) was mixed with 400 μL of Folin and Ciocalteu’s phenol reagent, and placed at room temperature for 3 min. Then added 40 μL of 10% Na2CO3 solution and shook it every 10 min. The absorbance value at 735 nm was measured by spectrophotometer after 1 h. The above-measured value was calculated to obtain the total polyphenol content using the regression equation (standard curve) and the results were expressed as gallic acid equivalent (mg GAE/g extract).
2.7. Extraction and Determination of Pigments of Mung Bean Seed Coat and Cotyledons
The enrichment of chlorophyll from the seed coat and cotyledons was as described with modification [24,25,26].
2.7.1. Acetone Extraction Method
A total of 5 g of the above-dried sample and 0.1 g of CaCO3 was added with 95% acetone 50 mL, homogenized, and packed into brown bottles. The extractions were carried out at 4 °C under a light-proof environment according to the experiment time. After centrifugation (4 °C, 1600× g, 10 min), the supernatant was quantified to 50 mL with 95% acetone, and the absorbance value at 650–670 nm was scanned by spectrophotometer, and the highest wavelength (649 nm) was measured by the following formula for chlorophyll content.
2.7.2. Hot Ethanol Extraction Method
A total of 5 g of dried sample and 0.1 g of CaCO3 were added to 50 mL of 80% ethanol (heated for 2 min in the 80 °C water bath) and then treated with ultrasonication for 10 min. The above extract was transferred to a brown flask and placed under 4 °C sheltered from light for the duration of the experiment. The extract was centrifuged (1600× g) for 10 min at 4 °C, and the supernatant was repeated 3 times. The supernatant was fixed to 50 mL with 95% ethanol and scanned by spectrophotometer for absorbance values at 650–670 nm, which was the highest wavelength (649 nm), with the measured absorbance values substituted into the following equation for chlorophyll content.
2.7.3. Ethanol Extraction Method
The treatment was consistent with the method above (Section 2.7.2), except that ultrasound-assisted extraction did not use.
The absorbance values of Chlorophyll a and b measured by the above method were then used to calculate the content according to the following equation:
Chlorophyll a = 13.7 × OD665 − 5.76 × OD649(1)
Chlorophyll b = 25.8 × OD649 − 7.60 × OD665(2)
Chlorophyll = Chlorophyll a + Chlorophyll b(3)
Unit: (mg/100 g dry sample weight).
2.8. Total Carotenoid Content
The following measurements were made according to Alam et al. [27] with minor modifications. First, 2 g of dried sample was homogenized with 40 mL acetone, 60 mL n-hexane, and 0.1 g MgCO3 for 5 min and then subjected to vacuum filtration. Then, the filtrate was washed twice with 25 mL acetone and once with 25 mL n-hexane. The filtrate was poured into a separatory funnel and then washed 5 times with 100 mL of distilled water. The absorbance value of 436 nm was measured by a spectrophotometer. Then, the absorbance values were quantified by comparing them with the standard curve of β-carotene.
2.9. Anthocyanin Measurement
Total anthocyanin content was determined by modifying the pH differential method because it has been adopted as a standard method by the Association of Official Analytical Chemists [28,29]. Finally, the results were presented as milligrams of anthocyanin equivalent per 100 g mung bean. The anthocyanin content concentration per 100 g mung bean was calculated with the following formula:
{(A × DF × MW × V) × 100}/ε × W(4)
where A is the difference of absorbance under pH 1 and pH 4.5 at ABS 520 nm, DF is the dilution factor (10 × following the procedure given), V is total extract volume, MW is the molecular weight (cyanidin 340 g mol−1), ε is the molar extinction coefficient (24,600 M−1 cm−1), and W is the total weight of mung bean.2.10. Purification with Mung Bean Seed Coat Pigments
The above chlorophyll extract of mung bean seed coat was concentrated by rotary evaporation at 30 °C under a vacuum. Then, the concentrates were processed by a Perista pump (SJ-1211L, ATTO Co., Tokyo, Japan) for light-protected gel filtration chromatography. The gel filtration chromatography purification method was modified from earlier reports [30,31]. A total of 1.0 mL concentrates were applied on a Sephadex G-15 column (1.0 cm × 30.0 cm; flow rate, 40 mL/h) (GiMiTEC, Milton, DE, USA) for separation. The column was equilibrated with 80% ethanol buffer and then eluted with separation. The 5 mL/tube fractions were collected and assayed for chlorophyll. Each tube of chlorophyll purified fraction was detected at OD436 nm, collecting those with higher absorbance values, and repeated 20 times for collection. Finally, the collected fractions were concentrated and freeze-dried. Then, the powder was stored at −20 °C.
2.11. High-Performance Liquid Chromatography (HPLC)
The purified chlorophyll drier powder was dissolved in methanol (50:50, v/v) diluted to a concentration of 3 mg L−1, filtrated by a 0.45 μm membrane, and analyzed by HPLC as described by Zhang et al. [32] and Zeb et al. [33] with modification. The HPLC system (Hitachi High-Tech Science, Tokyo) with a Mightysil RP-18 GP column (250 mm × 4.6 mm, 5.0 μm) (Kanto Chemical Holdings, Tokyo, Japan) was equilibrated with a mobile phase consisting of ethyl acetate/methanol/water (50/37.5/12.5, v/v/v) at a flow rate of 1 mL/min. The volume of standard or extract in each analysis was 5 μL, and procymidone was monitored at 400–700 nm using a photodiode array detector.
2.12. Statistical Analysis
All experiments were performed in triplicate and repeated at least twice. All data are expressed as mean ± SD and were analyzed by one-way ANOVA and Duncan’s test with SPSS v22.0 (International Business Machines Corp., Armonk, New York, NY, USA). p < 0.05 was considered statistically significant.
3. Results and Discussion
3.1. Component Analysis of Mung Bean
Mung bean mainly consists of the seed coat and cotyledon, the elemental compositions of which are presented in Table 1. Mung bean seed coat contained about 2.35% ash, 4.05% crude protein, and 0.99% crude lipid, and the seed leaves contained approximately 47.73% carbohydrate, 3.39% ash, 24.60% crude protein, and 0.69% crude fat. In addition, the amylose content of the cotyledons was about 15.13%. There was a significant difference in the component analysis of cotyledon and seed coat (p < 0.05). The seed coat did not contain amylose. Starch in a 3D gel network is extensively applied to produce desirable textures and viscosities and to improve the water holding capacity, structural properties, and shelf stability of foods [34]. It imparts desirable sensory properties to foods. Conversely, mung bean has a higher amylose content than grain or legumes, as evidenced by relatively high amylose content (30–45%, w/w) and lower gelatinization temperature in wheat starch [35].
3.2. Determination of pH Value
In this study, samples were heated in an enclosed manner to avoid moisture loss from the cotyledons and seed coats, which would cause concentration loss during the thermal process and errors in pH value determination. Changes in supernatant pH were observed after the thermal processing of cotyledons and seed coats. The pH of the supernatant of cotyledons heated at 120 °C for 10–50 min ranged from 6.06 to 6.35. However, we found no significant association in pH by different thermal times (Figure 1a). In the same conditions, the pH of treated seed coats ranged from 5.46 to 6.25, which was somewhat lower and significantly different compared to the cotyledon treated group (p < 0.05).
3.3. Color Characteristics
The product’s color is usually the first impression that influences consumer judgment. Therefore, good color and appearance are essential aspects of commercialization [36]. However, the description of the color is highly subjective, so a standard is needed to describe the various colors and strengths. In general, chromatic changes observed with agricultural products are due to enzymatic/non-enzymatic reactions [37].
Moreover, plants such as fruits and vegetables experienced significant color changes visually (i.e., green to yellow), possibly related to the metabolism of the chlorophyll content and carotenoids and flavonoids [38]. Three primary sources of color in mung beans are the surface texture layer and two from the seed coat. The texture layer is mainly brown pigment, and without this pigment, the seed coat will appear transparent; anthocyanin resides in the columnar layer of the seed coat, producing black spots, and the green color appears in the parenchyma layer, which is mainly composed of chloroplasts and will be yellow without it. The seed coat contains chromophores [7], which were easily leached by thermal processes during heating, causing the cotyledon cells and the liquid to change color. While heating at 120 °C for 10–50 min, the L value of the seed coat decreased from 66.57 to 55.17; however, the brightness tended to decrease with increasing thermal processing time, following the same trend observed with the steam-boiling sap of the cotyledons (Figure 1b) (p < 0.05). This shows the changes caused by the dissolution of the chromophore from the seed coat and the Maillard reaction of the cotyledon cells containing proteins and carbohydrates, which are prone to carbohydrates undergoing caramelization with either of these two reactions and will change the color of cotyledons [39,40]. Preoptimization is the most common chlorophyll change in food processing.
The chlorophyll-bound Mg is released when the green plant undergoes thermal processing caused by the denaturation of lipoproteins and loses their protective effect, H+ replaces Mg2+ in tetrapyrrole to pheophytin, and the green color vanishes to form a brownish-yellow [41]; the reactions will be promoted by thermal and acidity conditions, oxidizing compounds, or active species [42,43]. The pheophytin formation rate will increase following the temperature increase in acidic conditions [44,45]. According to the variation in chlorophyll a and b values, in the thermal treatment process at 120 °C for 10–50 min, the color trend was significantly greater in seed coats (red and yellow) than in cotyledons (Figure 1c,d). Chromatic a and b variations were obtained at 120 °C for 10–50 min; the cotyledon chromatic changes increased from 1.81 to 5.45 and 11.97 to 13.52, respectively. In contrast, seed coat chromatic changes rose from 5.88 to 14.37 and 21.55 to 34.85. The above results were statistically significantly different (p < 0.05). This phenomenon is due to the damage of surface cell membranes in the seed coat caused by thermal treatment, which resulted in the leaching of organic acids and the deterioration of chlorophyll cysts (thylakoid members), which accelerated the release of chlorophyll from the seed coat. Therefore, presumably, the chlorophyll formed in the seed coat by the thermal and acidic effects during the thermal process transformed into pheophytin with a yellowish-brown color, thus enhancing the color.
3.4. Browning Index
The reactants of the Maillard reaction include mainly sugars, amino acids, and ascorbic acid, whereas the middle products included 5-hydroxymethylfurfural and furfural [26]. In addition, the final products include browning pigments such as melanoidin and melanin, with other secondary products of various odors [46]. The above intermediate product, 5-hydroxymethylfurfural produced by the Maillard reaction, is a pigment with strong absorbance at 420 nm; the browning index can respond in degrees determined by spectrophotometry detection at 420 nm. The rate of the Maillard reaction follows the relation between thermal (temperature) and processing time [47]. The browning index of both seed coat and cotyledons increased with thermal time (Figure 1e). At 120 °C for 10–50 min, the browning index of cotyledons rose from 0.06 to 0.18 and that of seed coats from 0.11 to 0.58. These results showed statistically significant differences (p < 0.05). Thus, elevated temperatures accelerated the browning degree and reduced the required lag time for the initial Maillard reaction, which also increased the rate of the Maillard browning reaction. With thermal treatment, hexose will interact with amino acids producing the Amadori rearrangement product, followed by 1,2-embolization to form hydroxymethylfurfural, the most important intermediate product of the Maillard reaction [40].
Nonetheless, the maximum absorbance wavelength of hydroxymethylfurfural was 290 nm (Figure 1f). In the present study, the hydroxymethylfurfural content did not increase or decrease with thermal time in seed coats and cotyledons. A study of mung bean noodle products also found that if related to anthocyanin degradation, the final product’s color tended to be black (decreasing L value) [48].
3.5. Change of Chlorophyll Content
Chlorophyll is a pigment that participates in photosynthesis. The chlorophyll found in the plant cell chloroplast has a small and well-arranged structure, forming ions combined with neighboring proteins and lipids, thus enabling the plant to appear green by reflecting green light and absorbing red and blue light [42]. The color of the food on its own or during processing will directly affect consumer acceptability and appetite; however, the color is also an easy way to evaluate the food quality. In this study, our analysis of the three main pigment contents in mung beans showed the highest chlorophyll content in the seed coat (7.37 ± 0.11), followed by anthocyanins (4.85 ± 1.14) and carotenoids (1.98 ± 0.29) (Table 2). Hence, chlorophyll in the seed coat is estimated to be one of the preferred pigments responsible for the emerald green color of mung bean products. Notably, the chlorophyll content in the seed coat decreased from 6.57 to 1.28 (mg/100 g) during heating at 120 °C for 10–50 min (Figure 2a), with statistically significant differences (p < 0.05), which meanwhile shows a negative relation with thermal processing time (Figure 2b). The results were consistent with the previous chromatic analysis. The thermal treatment caused the seed coat to break down. The chlorophyll release occurred during processing, which showed reduced chlorophyll content in the seed coat, thus enabling the cotyledons’ release and green coloration (Figure 2c).
3.6. Flavonoid Condensation Reaction
The basic structure of flavonoids is 2-phenyl-benzo-α-pyrones, composed of two A and C rings of benzo-α-pyrone bonded to a phenolic (phenyl ring). Because the form can connect various functional groups to the structure, the variety of species is enormous. Currently, there are more than 4000 species [49]. According to the above observation, thermal processing conferred no significant color change in mung bean cotyledons (p < 0.05). At the same time, the seed coats showed a pronounced color change when heated alone. This study investigated the difference in the composition of the seed coats further. The total flavonoid content (quercetin) of mung bean seed coat heated at 120 °C for 20–50 min increased from 2.93 ± 0.51 to 18.74 ± 0.41 (μg/mg); thus, the content was statistically significantly different from the heat processing time (Figure 3a) (p < 0.05). However, flavonoids are chemically reactive and easily oxidized to quinones; quinones can be condensed with other polyphenolic compounds to form dark-brown polymers [50]. The condensation of flavonoids to form a dark-brown mass may be closely related to the reduced L-value obtained in this study by thermal treatment of seed coats.
3.7. Oxidative Condensation of Polyphenols
Polyphenols are secondary metabolites widely found in plants with a C6-C3-C6 aromatic ring structure with six major groups: (1) phenolic acids, (2) flavonoids, (3) catechins, and theaflavins, (4) chalcones, (5) anthocyanins, and (6) anthraquinones [51,52]. The chemical properties of polyphenols are active with easy oxidation to quinone, which is an electron-phallic solid group that easily binds with hydrogen atoms to carry out many reactions, especially (1) as an oxidizing agent to promote oxidation with other molecules, (2) as fast polymerization to obtain dark-brown polymers, and (3) forming large complexes with proteins as well as thearubigins and theaflavins [53,54]. Thearubigins and theaflavins are a group of compounds with a brownish-red color, with strong absorbance values at 380 nm [55]. During the thermal treatment of the seed coats, the 380 nm absorbance value increased with thermal treatment, from 0.51 ± 0.02 to 1.53 ± 0.08 at 120 °C for 10 to 50 min (Figure 3b). However, the 380 nm absorbance value of the thermal treatment for 50 min showed the maximum value, which was statistically significant compared to other groups (p < 0.05). Turkmen et al. [56] studied seven vegetables processed by boiling, steaming, and microwaving and showed increases in total phenolic content for bell peppers, beans, leeks, and broccoli regardless of the cooking method whereas peas and pumpkin showed a decrease in content. The phenolic contents were influenced by vegetable species more than the processing method. Our study also found that the total polyphenols content (in gallic acid equivalents) increased with thermal processing time. The total polyphenols content increased from 2.10 ± 0.03 to 2.44 ± 0.01 (mg/g) at 120 °C for 10 to 50 min (Figure 3c). In particular, the total phenolic content showed a significant difference in 40 min at 120 °C (p < 0.05). The thearubigins and theaflavins are the most important products produced by the oxidation of polyphenols in black tea leaves and thus are responsible for the red color of black tea broth [57]. These two studies and our study revealed that the solution of the seed coats reddened gradually with increasing thermal treatment time. The reason for such phenomenon could be that the substrates of the polyphenols yielded the thearubin product caused by thermal catalysis and hence the color change.
3.8. Pigment Content of Different Parts in Mung Beans
In this study, the contents of chlorophyll, carotenoids, and anthocyanins in mung bean were analyzed by spectroscopy to find the natural pigments that had the best effect on color. The content of chlorophyll, anthocyanins, and carotenoids was in the order of seed coat > the whole bean > cotyledons (Table 2). The pigment was concentrated in the seed coat. The content of carotenoids in the whole bean was 0.12 ± 0.02 (mg/100 g), which was significantly lower than that for the other pigments and was not the primary pigment affecting mung bean color. Furthermore, the chlorophyll content of the seed coat, whole bean, and cotyledon was 7.37 ± 0.11, 0.70 ± 0.04, and 0.26 ± 0.15 (mg/100 g), respectively, significantly higher than the other two pigments (p < 0.05). Thus, chlorophyll was the primary source of green color in mung beans among the three pigments.
3.9. Extraction and Purification of Chlorophyll
Most studies have used acetone to extract chlorophyll; otherwise, methanol, ethanol, petroleum ether, and ethyl acetate are used alone or in combination [58]. The optimal conditions and solvents for extracting chlorophyll from mung bean seed coats were investigated with acetone, ethanol, and hot ethanol. The effectiveness of solvents used to extract chlorophyll (a + b) for 24 h was in the order of acetone > ethanol > hot ethanol; the extraction rates were 7.64, 7.39, and 5.00 (mg/100 g) (Table 3), respectively. The acetone and ethanol extraction rates were nearly identical, with no significant differences (p < 0.05). The hot ethanol method obtained the highest chlorophyll (a + b) content, 4.9 (mg/100 g), at 2 h of extraction, so hot ethanol could accelerate the dissolution of chlorophyll at the beginning of extraction.
In contrast, the content with this method increased from only 4.9 to 5.0 (mg/100 g) at the subsequent extraction times of 24 h, with no trend of increase and poor effect. However, ethanol (vapor treatment) was used to help delay harvested cauliflower’s senescence (retarded chlorophyll degradation) [59]. Ethanol has an inhibitory effect on chlorophyllase activity and gene expression, reducing the degradation of chlorophyll, which is related to the photosystem [36]. This situation also explains the accelerated leaching of chlorophyll at the beginning of extraction. Hence, the content was higher than in other treatments, and the chlorophyll degradation was higher with treatment time and thermal effects.
3.10. Chlorophyll Purification and Assay Analysis
In heated vegetables, the cleavage rate is 7- to 9-fold faster for chlorophyll a than b [60]. Chlorophyll b was more stable than chlorophyll a during heating, with the crude extract of chlorophyll b measured by complete wavelength scanning (UV-visible light) to show high absorption peaks at 440, 460, and 640 nm, as well as the highest absorption peak at 460 nm, and is used as an indicator for chlorophyll collection. However, those methods are time-consuming and cannot separate complex components. Nowadays, column chromatography with HPLC commonly uses separate target compounds [60]. Following the identification of the target substances by HPLC, column chromatography can separate the different molecular weights of chlorophyll and derivatives by gel filtration chromatography. In this study, first, the chlorophyll was purified by gel filtration chromatography and then the chlorophyll and its components were analyzed by reverse-phase HPLC (RP-HPLC). The identification involved using a photodiode array detector, referring to Canjura, F. L. and S. J. Schwartz [61]. Chlorophyll belongs to the small-molecule polar substances, so it is not suitable for normal-phase chromatography, because the polar filler of the normal phase is active and quickly causes the pigment to form isomers [62].
However, the reverse phase is chiefly the effect of the hydrophobic group, and it causes more minor damage to chlorophyll. Then, the separation by Sephadex G-15 colloid depends on the molecular weight for purification purposes. With the fraction collected by the fraction collector, the analysis involved spectrophotometry to detect the highest absorption peak of chlorophyll at OD 460 nm to obtain the chromatogram of gel filtration (Figure 4a). Hence, the chlorophyll extract of mung-bean seed-coat fractions appeared between fraction tubes numbers 11 to 24 when the column was eluted, and the chlorophyll or its components showed the maximum peak absorption in tube 13. Therefore, after each sample loading, the sample was repeatedly collected from tubes 11 to 24 with a splitting collector. Each tube was composed after the above gel filtration, then compared with the standard by HPLC in terms of retention time. Tube 12 had the same retention time of 52.14 min as chlorophyll b (Figure 4b #A); meanwhile, tube 15 showed a retention time of 61.93 min like chlorophyll a (Figure 4b #B).
Furthermore, the scanning wavelength range in Figure 4b, #A, was 400 to 700 nm at the retention time of 52.14 min, with a broad wave-shape peak due to the absorption of the light source by the moieties, with absorbance λ max at 462, 650 nm (Figure 4c #A). However, for #B, the maximum absorbance values were 430, 656, and 668 (Figure 4c #B), with the same absorbance values as chlorophyll b and a. The HPLC analysis showed the same retention time as the standard chlorophyll and spectral properties when detected by a photodiode array. According to the results, mung bean extracts #A and #B were shown to be chlorophyll b and chlorophyll a. In brief, the change in chlorophyll pattern and the Maillard reaction of mung bean seed coat were the leading causes of chromatic changes with thermal treatment.
4. Conclusions
In this study, the color change of mung beans during thermal processing was found. It showed that in a weak acidic condition with heat, chlorophyll content in mung bean seed coat decreased (6.57–1.28 mg/100 g), which was shown to be statistically significant in each group (p < 0.05), and tawny pheophytin was formed; this phenomenon was related to increased chlorophyll b value (yellowness, 21.55–34.85). In contrast to the mung bean seed coat and cotyledon groups, all had significant differences (p < 0.05). Consequently, total flavonoid content (2.93 to 18.74 μg/mg) in the seed coat showed an increasing trend with thermal time. The condensation reaction of flavonoids with other polyphenols formed dark-brown polymers, closely related to the reduced brightness (L-value) in the supernatant of the seed coat with thermal processing time, which was statistically significant (p < 0.05). Therefore, the chlorophyll extraction and purification of the mung bean seed coat confirmed the morphological change of chlorophyll combined with the Maillard reaction, which caused the color change after thermal treatment. This study fills an existing knowledge gap and synergistically manipulates the interactions between different food components for potential application in mung bean processing, which might be valuable in the beverage or baking industries.
5. Highlights
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The color changes of mung beans during thermal processing were evaluated.
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Under weak acidic conditions with heat, the chlorophyll content in the seed coat decreased.
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Chromatic changes of chlorophyll were combined with the Maillard reaction after thermal treatment.
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The results of this study might be applied in the beverage or baking industries.
Conceptualization, Y.-T.C.; Data curation, P.-H.L.; Formal analysis, Y.-J.C. and W.-C.L.; Funding acquisition, Y.-T.C. and P.-H.L.; Investigation, Y.-J.C. and P.-H.L.; Methodology, Y.-J.C., W.-C.L. and P.-H.L.; Project administration, Y.-T.C.; Software, W.-C.L.; Validation, P.-H.H.; Writing—original draft, P.-H.H.; Writing—review & editing, W.-C.L. and P.-H.L. All authors have read and agreed to the published version of the manuscript.
Not applicable.
Not applicable.
Data sharing not applicable. No new data were created or analyzed in this study. Data sharing is not applicable to this article.
This study was also supported by grants provided by the Ministry of Science and Technology (MOST 111-2410-H-126-003) in Taiwan. The authors also thank Tainan Agricultural Research and Extension Station (Tainan District, Taiwan) for providing the mung bean samples.
The authors declare no conflict of interest. They have no known competing financial interests or personal relationships that could have influenced the work reported in this paper.
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Enlarge this image.Figure 1. Changes in physicochemical properties in heating aqueous solutions of mung bean seed coat and cotyledon obtained by thermal treatment at 120 °C for 10–50 min. Data are expressed as the mean ± SD; those with the same letter are not significantly different (p > 0.05). (a) pH value; (b) L value (brightness); (c) a value (Redness); (d) b value (yellowness); (e) Browning index (OD 420 nm); (f) HMF (hydroxymethylfurfural) (OD 290 nm).
Enlarge this image.Figure 2. Chlorophyll content and color of heating aqueous solutions of mung bean seed coats and cotyledons with the appearance changes of cotyledons, obtained by thermal treatment at 120 °C for 10–50 min. Data are expressed as the mean ± SD; those with the same letter are not significantly different (p > 0.05). (a) Chlorophylls content (mg/100 g); (b) The changes of boiled aqueous solutions color; (c) The appearance change of cotyledon color.
Enlarge this image.Figure 3. Evolution of total flavonoid content, oxidative condensation of polyphenols, and total polyphenol content of mung bean seed coats by thermal treatment at 120 °C for 10–50 min. Data are expressed as the mean ± SD; those with the same letter are not significantly different (p > 0.05). (a) Total flavonoid content (µg/mg); (b) Oxidative condensation of polyphenols (OD 380 nm); (c) Total polyphenolic content (mg/g).
Enlarge this image.Figure 4. Purification, HPLC chromatography, and photodiode array identification of the pigment found in mung bean seed coat. (a) Chromatography of mung bean seed coat pigment extracted by Sephdax G-15 gel filtration; (b) Determination of retention time of purified compounds (#A and #B) by HPLC; (c) Chromatography of purified compounds (#A and #B) by Photodiode array.
Enlarge this image.Figure 4. Purification, HPLC chromatography, and photodiode array identification of the pigment found in mung bean seed coat. (a) Chromatography of mung bean seed coat pigment extracted by Sephdax G-15 gel filtration; (b) Determination of retention time of purified compounds (#A and #B) by HPLC; (c) Chromatography of purified compounds (#A and #B) by Photodiode array.
Component analysis of mung bean seed coat and cotyledons.
| Content (%) | Ash | Crude Protein | Crude Lipid | Carbohydrate | Amylose | |
|---|---|---|---|---|---|---|
| Sample | ||||||
| Cotyledon | 3.39 a ± 0.23 | 24.60 a ± 2.17 | 0.69 b ± 0.18 | 47.73 a ± 2.25 | 15.13 a ± 0.11 | |
| Seed coat | 2.35 b ± 0.11 | 4.05 b ± 0.73 | 0.94 a ± 0.12 | 3.51 b ± 0.58 | 0.00 b ± 0.03 | |
Date is mean ± SD of three replications on a dry weight basis. a,b: Means within a row with different letters in the superscript are significantly different (p < 0.05).
Pigment content of mung bean seed coat and cotyledons.
| Chlorophylls | Carotenoids | Anthocyanins | |
|---|---|---|---|
| Whole bean | 0.70 b ± 0.04 | 0.12 b ± 0.02 | 0.39 b ± 0.15 |
| Seed coat | 7.37 a ± 0.11 | 1.98 a ± 0.29 | 4.85 a ± 1.14 |
| Cotyledon | 0.26 c ± 0.15 | 0.02 c ± 0.01 | 0.29 b ± 0.08 |
Date is mean ± SD of three replications on a dry weight basis. a–c: Means within a row with different letters in the superscript are significantly different (p < 0.05).
Evaluation of chlorophyll extraction with acetone, hot ethanol, and ethanol.
| Solvent | Acetone | Hot-Ethanol | Ethanol | |||
|---|---|---|---|---|---|---|
| Extraction Time (h) | 2 | 24 | 2 | 24 | 2 | 24 |
| chlorophyll a | 0.95 D ± 0.12 | 2.69 A ± 0.15 | 1.48 C ± 0.12 | 1.55 C ± 0.15 | 1.20 C ± 0.16 | 2.35 B ± 0.15 |
| chlorophyll b | 2.15 D ± 0.04 | 4.95 A ± 0.06 | 3.42 B ± 0.06 | 3.46 B ± 0.02 | 2.91 C ± 0.01 | 5.04 A ± 0.03 |
| chlorophyll a + b | 3.10 D ± 0.10 | 7.64 A ± 0.18 | 4.9 B ± 0.18 | 5.00 B ± 0.16 | 4.11 C ± 0.14 | 7.39 A ± 0.17 |
Data are the pigment (mg)/dry weight of mung bean (100 g). A–D: Means within a column with different letters in the superscript are significantly different (p < 0.05).
References
1. Pataczek, L.; Zahir, Z.A.; Ahmad, M.; Rani, S.; Nair, R. Beans with benefits—The role of mungbean (Vigna radiata) in a changing environment. Am. J. Bot.; 2018; 9, 24.
2. Australian Centre for International Agricultural Research (ACIAR). International Mungbean Improvement Network (IMIN): Mung Central. 2016; Available online: https://avrdc.org/download/project_newsletters/mung_central_newsletter/Mungbean-Central-ED1-2016-final-version.pdf (accessed on 16 May 2022).
3. Batzer, J.C.; Singh, A.; Rairdin, A.; Chiteri, K.; Mueller, D.S. Mungbean: A preview of disease management challenges for an alternative U.S. cash crop. J. Integr. Pest. Manag.; 2022; 13, 4. [DOI: https://dx.doi.org/10.1093/jipm/pmab044]
4. Lambrides, C.J.; Godwin, I.D. Mungbean. Pulses, Sugar and Tuber Crops; Chittaranjan, K. Springer: Berlin/Heidelberg, Germany, 2007; pp. 69-90.
5. Kumar, A.Y.; Seema, P.; Kumar, R.R.; Kushwaha, A.; Rashid, M.H.; Tarannum, N.; Chakraborty, S. Mungbean (Vigna radiata (L.) R. Wilczek): Progress in breeding and future challenges. Int. J. Plant Soil Sci.; 2022; 34, pp. 50-59. [DOI: https://dx.doi.org/10.9734/ijpss/2022/v34i330846]
6. Wang, F.; Huang, L.; Yuan, X.; Zhang, X.; Guo, L.; Xue, C.; Chen, X. Nutritional, phytochemical and antioxidant properties of 24 mung bean (Vigna radiate L.) genotypes. Food Prod. Process. Nutr.; 2021; 3, 28. [DOI: https://dx.doi.org/10.1186/s43014-021-00073-x]
7. Watt, E.E.; Poehlman, J.M.; Cumbie, B.G. Origin and composition of a texture layer on seeds of mungbean. Crop Sci.; 1977; 17, pp. 121-125. [DOI: https://dx.doi.org/10.2135/cropsci1977.0011183X001700010033x]
8. Villaño, D.; García-Viguera, C.; Mena, P. Colors: Health effects. Encyclopedia of Food and Health; Caballero, B.; Finglas, P.M.; Toldrá, F. Academic Press: Oxford, UK, 2016; pp. 265-272.
9. Pither, R.J. Canning|Quality changes during canning. Encyclopedia of Food Sciences and Nutrition; 2nd ed. Caballero, B. Academic Press: Oxford, UK, 2003; pp. 845-851.
10. Scheer, H. Chlorophylls and carotenoids. Encyclopedia of Biological Chemistry; 2nd ed. Lennarz, W.J.; Lane, M.D. Academic Press: Waltham, MA, USA, 2013; pp. 498-505.
11. Yilmaz, C.; Gökmen, V. Chlorophyll. Encyclopedia of Food and Health; Caballero, B.; Finglas, P.M.; Toldrá, F. Academic Press: Oxford, UK, 2016; pp. 37-41.
12. Giuliani, A.; Cerretani, L.; Cichelli, A. Colors: Properties and determination of natural pigments. Encyclopedia of Food and Health; Caballero, B.; Finglas, P.M.; Toldrá, F. Academic Press: Oxford, UK, 2016; pp. 273-283.
13. Popalia, C.; Kumar, N. Effect of temperature and processing time on physico-chemical characteristics in hot water blanching of sweet corn kernels. J. Inst. Eng. Ser. A; 2021; 102, pp. 163-173. [DOI: https://dx.doi.org/10.1007/s40030-021-00508-1]
14. Popaliya, C.; Kumar, N. Physicochemical characteristics of sweet corn kernels during microwave blanching. J. Food Process. Preserv.; 2022; 46, e16413. [DOI: https://dx.doi.org/10.1111/jfpp.16413]
15. Patel, K.K.; Kar, A. Heat pump assisted drying of agricultural produce—An overview. J. Food Sci. Technol.; 2012; 49, pp. 142-160. [DOI: https://dx.doi.org/10.1007/s13197-011-0334-z]
16. AACC. Approved Method of the AACC; 10th ed. American Association of Cereal Chemists: Heathrow, UK, 2000.
17. Williams, P.C.; Kuzina, F.D.; Hlynka, I. A rapid colorimetric procedure for estimating the amylose content of starches and flours. Cereal Chem.; 1970; 47, pp. 411-420.
18. Yadav, B.S.; Sharma, A.; Yadav, R.B. Resistant starch content of conventionally boiled and pressure-cooked cereals, legumes and tubers. J. Food Sci. Technol.; 2010; 47, pp. 84-88. [DOI: https://dx.doi.org/10.1007/s13197-010-0020-6]
19. Pham, H.T.T.; Bista, A.; Kebede, B.; Buvé, C.; Hendrickx, M.; van Loey, A. Insight into non-enzymatic browning of shelf-stable orange juice during storage: A fractionation and kinetic approach. J. Sci. Food Agric.; 2020; 100, pp. 3765-3775. [DOI: https://dx.doi.org/10.1002/jsfa.10418] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/32270878]
20. Pandey, S.; Sundararajan, S.; Ramalingam, S.; Pant, B. Elicitation and plant growth hormone-mediated adventitious root cultures for enhanced valepotriates accumulation in commercially important medicinal plant Valeriana jatamansi Jones. Acta Physiol. Plant.; 2021; 44, 4. [DOI: https://dx.doi.org/10.1007/s11738-021-03319-w]
21. Anwar, F.; Latif, S.; Przybylski, R.; Sultana, B.; Ashraf, M. Chemical composition and antioxidant activity of seeds of different cultivars of mungbean. J. Food Sci.; 2007; 72, pp. S503-S510. [DOI: https://dx.doi.org/10.1111/j.1750-3841.2007.00462.x] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/17995664]
22. Liu, Y.; Chen, W.; Fan, L. Effects of different drying methods on the storage stability of barley grass powder. J. Sci. Food Agric.; 2022; 102, pp. 1076-1084. [DOI: https://dx.doi.org/10.1002/jsfa.11443]
23. Chen, J.Y.; Yen, G.C.; Tsai, N.T.; Lin, J.A. Risk and benefit of natural and commercial dark brown sugars as evidenced by phenolic and Maillard reaction product contents. J. Agric. Food Chem.; 2021; 69, pp. 767-775. [DOI: https://dx.doi.org/10.1021/acs.jafc.0c04795]
24. Arnon, D.I. Copper enzymes in isolated chloroplasts. Polyphenol oxidase in Beta vulgaris. Plant Physiol.; 1949; 24, pp. 1-15. [DOI: https://dx.doi.org/10.1104/pp.24.1.1]
25. Zhang, S.; Sheng, Y.N.; Feng, Y.C.; Diao, J.J.; Wang, C.Y.; Zhang, D.J. Changes in structural and functional properties of globulin-polyphenol complexes in mung beans: Exploration under different interaction ratios and heat treatment conditions. Int. J. Food Sci. Technol.; 2022; 57, pp. 1920-1935. [DOI: https://dx.doi.org/10.1111/ijfs.15180]
26. Zhang, S.; Feng, Y.C.; Fu, T.X.; Sheng, Y.N.; Diao, J.J.; Wang, C.Y. Effect of processing on the phenolics content and antioxidant properties of mung bean. Cereal Chem.; 2021; 98, pp. 355-366. [DOI: https://dx.doi.org/10.1002/cche.10375]
27. Alam, M.K.; Rana, Z.H.; Islam, S.N. Comparison of the proximate composition, total carotenoids and total polyphenol content of nine orange-fleshed sweet potato varieties grown in Bangladesh. Foods; 2016; 5, 64. [DOI: https://dx.doi.org/10.3390/foods5030064]
28. Johnson, J.; Collins, T.; Walsh, K.; Naiker, M. Solvent extractions and spectrophotometric protocols for measuring the total anthocyanin, phenols and antioxidant content in plums. Chem. Pap.; 2020; 74, pp. 4481-4492. [DOI: https://dx.doi.org/10.1007/s11696-020-01261-8]
29. Lee, J.; Durst, R.; Wrolstad, R. AOAC 2005.02: Total Monomeric Anthocyanin Pigment Content of Fruit Juices, Beverages, Natural Colorants, and Wines—pH Differential Method. Official Methods of Analysis of AOAC International; AOAC International: Rockville, MD, USA, 2005.
30. Wu, M.C.; Jiang, C.M.; Huang, P.H.; Wu, M.Y.; Wang, Y.T. Separation and utilization of pectin lyase from commercial pectic enzyme via highly methoxylated cross-linked alcohol-insoluble solid chromatography for wine methanol reduction. J. Agric. Food Chem.; 2007; 55, pp. 1557-1562. [DOI: https://dx.doi.org/10.1021/jf062880s] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/17263548]
31. Masoodi, K.Z.; Lone, S.M.; Rasool, R.S. Gel-filtration or size-exclusion chromatography. Advanced Methods in Molecular Biology and Biotechnology; Masoodi, K.Z.; Lone, S.M.; Rasool, R.S. Academic Press: Amsterdam, The Netherlands, 2021; Chapter 26 pp. 147-149.
32. Zhang, Z.; Lin, H.; Sui, J.; Han, X.; Wang, L.; Sun, X.; Cao, L. The effect of chlorophyll on the enzyme-linked immunosorbent assay (ELISA) of procymidone in vegetables and the way to overcome the matrix interference. J. Sci. Food Agric.; 2021; 102, pp. 3393-3399. [DOI: https://dx.doi.org/10.1002/jsfa.11686] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/34825360]
33. Zeb, A.; Khan, S.; Ercişli, S. Characterization of carotenoids, chlorophylls, total phenolic compounds, and antioxidant activity of Brassica oleracea L. var. botrytis leaves from Pakistan. Biologia; 2022; 77, pp. 315-324. [DOI: https://dx.doi.org/10.1007/s11756-021-00905-8]
34. Min, C.; Ma, W.; Kuang, J.; Huang, J.; Xiong, Y.L. Textural properties, microstructure and digestibility of mungbean starch-flaxseed protein composite gels. Food Hydrocoll.; 2022; 126, 107482. [DOI: https://dx.doi.org/10.1016/j.foodhyd.2022.107482]
35. Li, K.; Zhang, M.; Bhandari, B.; Xu, J.; Yang, C. Improving storage quality of refrigerated steamed buns by mung bean starch composite coating enriched with nano-emulsified essential oils. J. Food Process. Eng.; 2020; 43, e13475. [DOI: https://dx.doi.org/10.1111/jfpe.13475]
36. Awad, A.H.R.; Parmar, A.; Ali, M.R.; El-Mogy, M.M.; Abdelgawad, K.F. Extending the shelf-life of fresh-cut green bean pods by ethanol, ascorbic acid, and essential oils. Foods; 2021; 10, 1103. [DOI: https://dx.doi.org/10.3390/foods10051103] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/34067518]
37. Kannan, V.S.; Arjunan, T.V.; Vijayan, S. Drying characteristics of mint leaves (Mentha arvensis) dried in a solid desiccant dehumidifier system. J. Food Sci. Technol.; 2021; 58, pp. 777-786. [DOI: https://dx.doi.org/10.1007/s13197-020-04595-z]
38. Monribot-Villanueva, J.L.; Altúzar-Molina, A.; Aluja, M.; Zamora-Briseño, J.A.; Elizalde-Contreras, J.M.; Bautista-Valle, M.V.; de los Santos, J.A.; Sánchez-Martínez, D.E.; Rivera-Reséndiz, F.J.; Vázquez-Rosas-Landa, M. et al. Integrating proteomics and metabolomics approaches to elucidate the ripening process in white Psidium guajava. Food Chem.; 2022; 367, 130656. [DOI: https://dx.doi.org/10.1016/j.foodchem.2021.130656]
39. Lotfy, S.N.; Saad, R.; El-Massrey, K.F.; Fadel, H.H.M. Effects of pH on headspace volatiles and properties of Maillard reaction products derived from enzymatically hydrolyzed quinoa protein-xylose model system. LWT; 2021; 145, 111328. [DOI: https://dx.doi.org/10.1016/j.lwt.2021.111328]
40. Moldoveanu, S.C. 8—Analytical pyrolysis of caramel colours and of maillard browning polymers. Analytical Pyrolysis of Natural Organic Polymers; 2nd ed. Moldoveanu, S.C. Elsevier: Amsterdam, The Netherlands, 2021; Volume 20, pp. 315-333.
41. Sharma, S.; Katoch, V.; Kumar, S.; Chatterjee, S. Functional relationship of vegetable colors and bioactive compounds: Implications in human health. J. Nutr. Biochem.; 2021; 92, 108615. [DOI: https://dx.doi.org/10.1016/j.jnutbio.2021.108615]
42. Lin, Y.P.; Charng, Y.Y. Chlorophyll dephytylation in chlorophyll metabolism: A simple reaction catalyzed by various enzymes. Plant Sci.; 2021; 302, 110682. [DOI: https://dx.doi.org/10.1016/j.plantsci.2020.110682] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/33288004]
43. Shiekh, K.A.; Olatunde, O.O.; Zhang, B.; Huda, N.; Benjakul, S. Pulsed electric field assisted process for extraction of bioactive compounds from custard apple (Annona squamosa) leaves. Food Chem.; 2021; 359, 129976. [DOI: https://dx.doi.org/10.1016/j.foodchem.2021.129976] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/33957326]
44. Liaotrakoon, W.; Liaotrakoon, V.; Hongtongsuk, T. Effect of solid-based feed concentration and water temperature on physicochemical, chlorophyll and antioxidative properties of Pandanus amaryllifolius leaf extract. J. Food Process. Preserv.; 2021; 45, e15735. [DOI: https://dx.doi.org/10.1111/jfpp.15735]
45. Lichtenthaler, H.K. Chlorophylls and carotenoids: Pigments of photosynthetic biomembranes. Methods in Enzymology; Academic Press: London, UK, 1987; Volume 148, pp. 350-382.
46. Ribeiro, J.A.; Cantillano, R.F.F.; Nora, F.R.; Nora, L. Effect of 4-hexylresorcinol on post-cut browning and quality of minimally processed ‘fuji’ apple fruits. J. Food Meas. Charact.; 2020; 14, pp. 2461-2471. [DOI: https://dx.doi.org/10.1007/s11694-020-00494-1]
47. Rodier, L.C.; Hartel, R.W. Characterizing Maillard reaction kinetics and rheological changes in white chocolate over extended heating. J. Food Sci.; 2021; 86, pp. 2553-2568. [DOI: https://dx.doi.org/10.1111/1750-3841.15772] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/34056726]
48. Ma, Y.; Wang, A.; Yang, M.; Wang, S.; Wang, L.; Zhou, S.; Blecker, C. Influences of cooking and storage on γ-aminobutyric acid (GABA) content and distribution in mung bean and its noodle products. LWT; 2022; 154, 112783. [DOI: https://dx.doi.org/10.1016/j.lwt.2021.112783]
49. Qu, Z.; Tang, Z.; Liu, F.; Sablani, S.S.; Tang, J. Quality of green beans (Phaseolus vulgaris L.) influenced by microwave and hot water pasteurization. Food Control; 2021; 124, 107936. [DOI: https://dx.doi.org/10.1016/j.foodcont.2021.107936]
50. Palamutoğlu, R. Antibrowning effect of commercial and acid-heat coagulated whey on potatoes during refrigerated storage. J. Food Sci.; 2020; 85, pp. 3858-3865. [DOI: https://dx.doi.org/10.1111/1750-3841.15468]
51. Brglez Mojzer, E.; Hrnčič, M.K.; Škerget, M.; Knez, Ž.; Bren, U. Polyphenols: Extraction methods, antioxidative action, bioavailability and anticarcinogenic effects. Molecules; 2016; 21, 901. [DOI: https://dx.doi.org/10.3390/molecules21070901]
52. Alara, O.R.; Abdurahman, N.H.; Ukaegbu, C.I. Extraction of phenolic compounds: A review. Curr. Res. Food Sci.; 2021; 4, pp. 200-214. [DOI: https://dx.doi.org/10.1016/j.crfs.2021.03.011]
53. Imran, A.; Arshad, M.U.; Arshad, M.S.; Imran, M.; Saeed, F.; Sohaib, M. Lipid peroxidation diminishing perspective of isolated theaflavins and thearubigins from black tea in arginine induced renal malfunctional rats. Lipids Health Dis.; 2018; 17, 157. [DOI: https://dx.doi.org/10.1186/s12944-018-0808-3] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/30021615]
54. Takemoto, M.; Takemoto, H. Synthesis of theaflavins and their functions. Molecules; 2018; 23, 918. [DOI: https://dx.doi.org/10.3390/molecules23040918] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/29659496]
55. Tse, S.Y. Food and nutritional analysis|Coffee, cocoa, and tea*. Encyclopedia of Analytical Science; 2nd ed. Worsfold, P.; Townshend, A.; Poole, C. Elsevier: Amsterdam, The Netherlands, 2005; pp. 279-285.
56. Turkmen, N.; Poyrazoglu, E.; Sari, F.; Velioglu, Y. Effects of cooking methods on chlorophylls, pheophytins and colour of selected green vegetables. Int. J. Food Sci. Technol.; 2006; 41, pp. 281-288. [DOI: https://dx.doi.org/10.1111/j.1365-2621.2005.01061.x]
57. Koch, W. Theaflavins, thearubigins, and theasinensins. Handbook of Dietary Phytochemicals; Xiao, J.; Sarker, S.D.; Asakawa, Y. Springer: Singapore, 2020; pp. 1-29.
58. Xu, D.P.; Li, Y.; Meng, X.; Zhou, T.; Zhou, Y.; Zheng, J.; Zhang, J.J.; Li, H.B. Natural antioxidants in foods and medicinal plants: Extraction, assessment and resources. Int. J. Mol. Sci.; 2017; 18, 96. [DOI: https://dx.doi.org/10.3390/ijms18010096]
59. Suzuki, Y.; Uji, T.; Terai, H. Inhibition of senescence in broccoli florets with ethanol vapor from alcohol powder. Postharvest Biol. Technol.; 2004; 31, pp. 177-182. [DOI: https://dx.doi.org/10.1016/j.postharvbio.2003.08.002]
60. Almela, L.; Fernández-López, J.A.; Roca, M.A.J. High-performance liquid chromatographic screening of chlorophyll derivatives produced during fruit storage. J. Chromatogr. A; 2000; 870, pp. 483-489. [DOI: https://dx.doi.org/10.1016/S0021-9673(99)00999-1]
61. 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, pp. 1102-1105. [DOI: https://dx.doi.org/10.1021/jf00006a020]
62. Caesar, J.; Tamm, A.; Ruckteschler, N.; Leifke, A.; Weber, B. Revisiting chlorophyll extraction methods in biological soil crusts—Methodology for determination of chlorophyll a and chlorophyll a + b as compared to previous methods. Biogeosciences; 2018; 15, pp. 1415-1424. [DOI: https://dx.doi.org/10.5194/bg-15-1415-2018]
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; Yu-Tsung, Cheng 2 ; Yung-Jia, Chan 3 ; Wen-Chien, Lu 4 ; Li, Po-Hsien 5
1 College of Food, Jiangsu Food and Pharmaceutical Science College, No. 4, Meicheng Road, Higher Education Park, Huai’an 223003, China;
2 Cardiovascular Center, Taichung Veterans General Hospital, 1650 Sec. 4 Taiwan Boulevard, Xitun Dist., Taichung 40705, Taiwan;
3 College of Biotechnology and Bioresources, Da-Yeh University, 168, University Rd, Dacun, Changhua 51591, Taiwan;
4 Department of Food and Beverage Management, Chung-Jen Junior College of Nursing, Health Sciences and Management, 217, Hung-Mao-Pi, Chia-Yi City 60077, Taiwan
5 Department of Food and Nutrition, Providence University, 200, Sec. 7, Taiwan Boulevard, Shalu Dist., Taichung 43301, Taiwan