Antioxidative phenolic compounds from the aerial parts of Cyperus exaltatus var. iwasakii and their HPLC analysis
Applied Biological Chemistry volume 66, Article number: 61 (2023)
The constituents and antioxidant activities of Cyperus exaltatus var. iwasakii (CE) have not been studied to date. In this study, the 2,2-diphenyl-1-picrylhydrazyl (DPPH) and 2,2′-azino-bis(3-ethylbenzothiazoline-6 sulfonic acid) (ABTS) assays were used to evaluate the radical-scavenging activities of the ethanol extract, four fractions, and isolated compounds of CE. In addition, phenolic acids and flavonoids were isolated from the ethanol extract of CE using column chromatography. The compounds identified by spectroscopy were gallic acid, protocatechuic acid, vanillic acid, p-coumaric acid, rutin, ferulic acid, isoquercitrin, astragalin, quercetin, luteolin, apigenin, tricin, and kaempferol. Quantitative analysis using high-performance liquid chromatography (HPLC) revealed that the major flavonoids of CE were astragalin and tricin and that the major phenolic acid was p-coumaric acid. In addition, comparative analysis of CE from Ganghwa and Hampyeong habitats using HPLC showed that the Hampyeong CE had a higher phytochemical content. Comparative analyses of the isolated compounds were also conducted among five Cyperus species. The highest antioxidant activities were found in the ethyl acetate (EtOAc) fraction, and among the compounds isolated from CE, vanillic acid and quercetin showed remarkable antioxidant activity even when compared with ascorbic acid. The results demonstrate the usefulness of CE, which has not been sufficiently studied previously, and will facilitate the evaluation of its potential effectiveness as antioxidant functional plant material.
Natural products are an essential source of potential drug intermediates for developing new drugs or health functional supplements. One of the many approaches employed in research of natural products is to select plants that have been commonly used since ancient times but have not been pharmacologically evaluated for possible antioxidant properties. These investigations should aim to identify the various biological functions and components of plant materials. Almost all Cyperus species are classified as weeds in agriculture, and herbicide development to remove them has been studied. Since Cyperus species are considered weeds, little research has been conducted on them, particularly in terms of their phytochemical composition and bioactivity [1,2,3].
Among Cyperus species, Cyperus exaltatus var. iwasakii (CE) is a sedge of the family of Cyperaceae and is commonly found in East Asia, Australia, Africa, and North America. CE culms have a triangular cross-section, a smooth surface, and a growing height of 100–180 cm. The leaves are 5–15 mm long, the spikelets are flat, and their color is yellow to dark yellowish-brown . In Africa, the rhizome of CE has been powdered and used to treat pus or anemia caused by malaria . In many countries, including the Republic of Korea, tough culms of CE have been used to make a variety of household products as well as cushions, slats, and huts. Thus, CE is a representative industrial crop among sedges that is used to make daily necessities. In Korea, the main CE production regions are Hampyeong and Ganghwa, and since much of the focus on CE since ancient times has been on its usefulness for making local products, its pharmacological aspects have rarely been explored.
Reactive oxygen species (ROS), such as singlet oxygen (1O2), superoxide anion (O2−), and hydroxyl (•OH) radical, and non-free radical species, such as hydrogen peroxide (H2O2), are frequently produced as byproducts of biological reactions or external stimuli [6, 7]. ROS have been shown to exert both positive and harmful effects. In some signal transduction pathways, ROS may function as second messengers at extremely low concentrations . However, excess generation of ROS is regarded as the primary cause of oxidative stress as a result of imbalanced antioxidant defense systems, and the creation of ROS may damage important biomolecules such as DNA, lipids, and proteins [9, 10]. Finally, oxidative stress causes the onset of age-related diseases such as cancer, hypertension, atherogenesis, Parkinson’s disease, and Alzheimer’s disease . Recent studies have shown that antioxidants derived from plants with free radical-scavenging characteristics may play a key role as therapeutic agents in the aging process and free radical-mediated illnesses such as neurodegeneration .
Plants have an important function in sustaining human health and improving human quality of life. Plant extracts and phytochemicals such as flavonoids and other polyphenolic compounds have been demonstrated to exhibit biological activity in vitro and in vivo, justifying traditional medicine research focused on the characterization of the biological activity of these plants . Due to their diverse pharmacological properties, including antimicrobial, antioxidant, anticancer, analgesic, anti-inflammatory, and apoptosis-inducing properties, plant-derived natural products, such as polyphenols, tannins, terpenes, alkaloids, and flavonoids, have garnered considerable attention in recent years [14,15,16]. Many synthetic antioxidant molecules have been demonstrated to be harmful and/or to have mutagenic properties, which has piqued the curiosity of numerous researchers in natural antioxidants [17, 18]. Thus, research into natural antioxidants has become a crucial topic. In light of its widespread application and chemical composition, the in vitro antioxidative activity of the extract, fractions, and isolated compounds of CE is worth evaluation.
In this study, various experiments were performed with CE, which is mainly used as a fiber and industrial crop, to broaden knowledge of its antioxidant properties and phytochemical components.
Materials and methods
The aerial parts of CE were collected from Hampyeong, Korea in 2019. The methanol (MeOH) extracts of CE, C. difformis (CD), C. microiria (CM), C. sanguinolentus (CS), and C. cyperoides (CC) were obtained from KRIBB in Daejeon, Korea. We also acquired CE grown in Ganghwa in 2019 for comparative analysis.
Apparatus and chemicals
Open-column chromatography was performed on silica gel (60–200 Mesh ASTM, Germany) with Sephadex LH-20 (Sigma-Aldrich, St. Louis, MO, USA) and LiChroprep RP-18 (Sigma-Aldrich, St. Louis, MO, USA). Chromatographic analysis was performed using a high-performance liquid chromatography (HPLC) system (Agilent Technology 1260 Infinity II; Santa Clara, CA, USA) and INNO C18 column (250 × 4.6 mm, 5 µm) equipped with a pump, an auto-sampler, and a diode array detector (DAD). The solvents ethanol (EtOH), methanol (MeOH), n-hexane, chloroform (CHCl3), acetic acid, ethyl acetate (EtOAc), and n-butanol (n-BuOH) used for extraction and fractionation were purchased from Samchun Pure Chemicals Co. (Pyeongtaek, Korea). Extraction and evaporation were performed in an EYELA apparatus (Japan) and with the EYELA rotary evaporator system (Japan). In addition, 2,2'-Azino-bis(3-ethylbenzothiazoline-6-sulfonic acid) (ABTS) was purchased from Roche Diagnostics GmbH (Mannheim, Germany), and 2,2-diphenyl-1-picrylhydrazyl (DPPH, 95%) was purchased from Alfa Aesar (Haverhill, USA).
DPPH radical-scavenging activity
The DPPH method is widely used to measure the antioxidant activity of water-soluble or organic solvent extracts from natural products. In this experiment, the antioxidant activity of extracts and fractions of CE was measured using the method described by Choi et al. . First, 200 μL of 2 mM DPPH dissolved in 95% ethanol and 10 μL of each experimental group were added to an E.P. tube, vortexed, and reacted in the dark for 30 min. The concentrations of the remaining radicals were measured using a microplate reader (514 nm). Ascorbic acid (Sigma, USA) was used as a positive control. The scavenging ability (IC50) of the extract against DPPH was expressed as the concentration required to reduce the absorbance of the control group using only the solvent by 50%.
ABTS radical-scavenging activity
The ABTS radical-scavenging activity of extracts and fractions of CE was measured by modifying a previously reported method . For the experimental method, 7.4 mM ABTS and potassium persulfate (2.6 mM) dissolved in distilled water were added in a 1:1 ratio and diluted with distilled water (pH 7.4) to an absorbance value of 1.00 ± 0.02. After leaving the mixture in the dark for 24 h, 10 μL of each sample prepared by concentration in 200 μL of the radical stock solution was added. After leaving the mixture for 30 min, the concentrations of the remaining radicals were measured using a microplate reader (734 nm). Ascorbic acid was used as a positive control. The concentration range was measured to be 0.5–0.025 mg/mL, and the scavenging ability (IC50) of the extract against ABTS was expressed as the concentration required to reduce the absorbance of the control group by 50% using only the solvent.
Total polyphenol content analysis
Total polyphenol content analysis was performed by modifying a previously reported Folin-Ciocalteu method  First, 60 μL of 2N Folin-Ciocalteu phenol reagent (St. Lewis, Sigma-Aldrich, USA) was added to the extract. Then, after adding 60 μL of 15% Na2CO3 to the solution for 30 min, absorbance was measured at 760 nm using a microplate reader (Epoch, BioTek, Winooski, Vietnam, USA). Finally, a calibration curve was created using gallic acid as the standard to quantify the total polyphenol content.
Extraction, fractionation, and isolation of CE
The dried aerial parts of CE (4.5 kg) from Hampyeong were cut into small pieces and extracted under reflux with 95% ethanol at 80–84℃ for three hours; this process was repeated three times with an equipped extractor. Using a rotary evaporator, the resultant extract solution was filtered and concentrated to obtain a crude ethanol extract (930 g). The ethanol extract of CE (920 g) was suspended in distilled water and subsequently partitioned with n-hexane, CHCl3, EtOAc, and n-BuOH to obtain the n-hexane (355 g), CHCl3 (25 g), EtOAc (14 g), and n-BuOH fractions (40 g). The EtOAc fraction (13.1 g) of CE was placed onto a silica gel column (6 × 80 cm) with a step gradient of CHCl3:MeOH (10:0 to 4:6) and pooled to produce a fraction. The fractions were analyzed and combined using thin layer chromatography to create six additional subfractions. These additional fractions were purified using Sephadex LH-20 column and then dissolved at a step gradient of H2O:MeOH (9:1–0:1 v/v) to produce compounds 1, 7, 8, 9, 10, 11, 12, and 13. The n-BuOH fraction (36 g) of CE was also eluted on an open column (8 × 100 cm) using a previously described method. The n-BuOH fraction revealed four fractions and recrystallized compounds 2–6.
Compound 1: White powder; EI-MS: m/z 170 [M]+ (100), 162 (40.0), 133 (50.0), 90 (24.0), 80 (12.0); 1H−NMR (500 MHz, DMSO-d6): δ 7.10 (2H, s, H−1, H−5); 13C-NMR (125 MHz, DMSO-d6): δ 110.8 (C-2, C-6), 122.4 (C-1), 139.3 (C-4), 146.4 (C-3, C-5), 168.7 (C-7) (Fig. S1 and S2).
Compound 2: White crystal; EI-MS: m/z 154 [M]+ (100), 162 (40.0), 135 (50.0), 90 (25.0), 76 (12.0); 1H-NMR (500 MHz, DMSO-d6): δ 7.43 (d, 1H, J = 2.0 Hz, H-2), 7.38 (dd, 1H, J = 8.0, 2.0 Hz, H-6), 6.77 (d, 1H, J = 8.0 Hz, H-5); 13C-NMR (125 MHz, DMSO-d6): δ 172.0 (C = O), 150.7 (C-4), 146.2 (C-3), 125.5 (C-6), 123.7 (C-1), 117.8 (C-2), 115.6 (C-5) (Fig. S3 and S4).
Compound 3: White powder; EI-MS: m/z 168 [M]+ (100), 284 (65.0), 258 (56.0), 168 (13.0), 137 (15.0), 119 (15.0), 83 (70.0), 65 (75.0); 1H-NMR (500 MHz, DMSO-d6): δ 7.54 (1H, d, J = 2.0 Hz, H-2), 7.55 (1H, dd, J = 8.0, 2.0 Hz, H-6), 6.82 (1H, d, J = 8.0 Hz, H-5), 4.00 (3H, s, OCH3); 13C-NMR (125 MHz, DMSO-d6): δ 166.5 (COOH), 152.3 (C-3), 148.4 (C-4), 123.0 (C-1), 122.0 (C-6), 113.4 (C-2), 111.9 (C-5), 54.1 (OCH3) (Fig. S5 and S6).
Compound 4: White powder; EI-MS: m/z 164 [M]+ (100), 257 (12.0), 228 (5.0), 181 (2.0), 154 (20.0), 128 (11.0), 106 (1.0), 88 (1.0), 70 (4.0); 1H-NMR (500 MHz, DMSO-d6): δ 7.51 (3H, t, J = 8.0, 15.0 Hz, H-3, H-5, H-9), 6.75 (2H, d, J = 8.0 Hz, H-6, H-8), 6.21 (1H, d, J = 16.5 Hz, H-2); 13C-NMR (125 MHz, DMSO-d6): δ 168.1 (C-9), 158.3 (C-4), 143.2 (C-7), 131.4 (C-2, C-6), 124.2 (C-1), 113.4 (C-3, C-5), 114.3 (C-8) (Fig. S7 and S8).
Compound 5: Yellow amorphous powder; FAB-MS: m/z 625. 1H-NMR (DMSO-d6, 500 MHz): δ = 6.18(1H, d, J = 2.0 Hz, H-6), 6.34(1H, d, J = 2.0, H-8), 7.49(1H, d, J = 2.0, H-2′), 6.79(1H, d J = 9.0, H-5′), 7.53(1H, dd, J = 9.0, 2.0, H-6′), 12.53(1H, s, 5-OH), 5.30(1H, d, J = 7.0, H-1′′), 4.34(1H, d, J = 2.0, H-1′′′), 1.00(3H, d, J = 6.0, H-6′′′). 13C-NMR (125 MHz, DMSO-d6): δ 156.4 (C-2), 131.2 (C-3), 176.3 (C-4), 156.5 (C-5), 100.6 (C-6), 164.4 (C-7), 93.6 (C-8), 161.2 (C-9), 104.1 (C-10), 121.6 (C-1′), 115.1 (C-2′), 144.7 (C-3′), 148.5 (C-4′), 116.1 (C-5′), 120.9 (C-6′), 101.2 (C-1′′), 71.8 (C-2′′), 74.1 (C-3′′), 71.6 (C-4′′), 72.8 (C-5′′), 67.0 (C-6′′) 103.8 (C-1′′′), 70.0 (C-2′′′), 70.2 (C-3′′′), 70.5 (C-4′′′), 68.2 (C-5′′′), 17.7 (C-6′′′) (Fig. S9 and S10).
Compound 6: Yellow amorphous powder; EI-MS: m/z 194 [M]+ (100), 180 (25), 160 (7), 134 (32), 101 (14), 90 (15), 71(27), 52(15); 1H-NMR (500 MHz, DMSO-d6) δ: 3.94 (3H, s, H-4'), 6.32 (1H, d, J = 14.0 Hz, H-2′), 6.85 (1H, d, J = 9.0 Hz, H-6), 7.14 (1H, dd, J = 8.0, 2.0 Hz, H-5), 7.01 (1H, d, J = 2.0 Hz, H-3), 7.74 (1H, d, J = 14.0 Hz, H-1′); 13C-NMR (125 MHz, DMSO-d6): δ 55.0 (C-4′), 108.3 (C-5), 114.5 (C-2), 114.8 (C-2'), 124.6 (C-3), 124.7 (C-4), 142.8 (C-1′), 147.0 (C-6), 148.3 (C-1), 171.3 (C-3′) (Fig. S11 and S12).
Compound 7: Yellow powder; FAB-MS: m/z 464. 1H-NMR (DMSO-d6, 500 MHz): δ 12.65 (5-OH), 7.58 (1H, d, J = 1.5 Hz, H-2′), 7.57 (1H, d, J = 2.0 Hz, H-6′), 6.85 (1H, dd, J = 1.5, 7.5 Hz, H-5′), 6.41 (1H, d, J = 2.0 Hz, H-8), 6.21 (1H, d, J = 2.0 Hz, H-6), 5.47 (1H, d, J = 7.0 Hz, H-1″), 3.60–3.10 (6H, m, H-2″-6′′). 13C-NMR (125 MHz, DMSO-d6): δ 155.1 (C-2), 133.4 (C-3), 177.3 (C-4), 162.3 (C-5), 98.6 (C-6), 165.1 (C-7), 93.4 (C-8), 156.2 (C-9), 104.9 (C-10), 121.6 (C-1′), 115.1 (C-2'), 143.7 (C-3'), 148.5 (C-4′), 116.1 (C-5′), 120.9 (C-6′), 100.7 (C-1″), 74.2 (C-2′), 76.4 (C-3′), 70.0 (C-4′), 77.7 (C-5′), 61.0 (C-6′) (Fig. S13 and S14).
Compound 8: Yellow powder; ESI–MS: m/z 447.2 [MH]-. 1H-NMR (DMSO-d6, 500 MHz) δ: 8.06 (2H, dd, J = 12.0, 2.5 Hz, H-2′, H-6'), 6.87 (2H, dd, J = 9.5, 2.5 Hz, H-3', H-5'), 6.40 (1H, d, J = 2.0 Hz, H-8), 6.18 (1H, d, J = 2.0 Hz, H-6), 5.15 (1H, d, J = 7.8 Hz, H-1″), 3.76 (1H, dd, J = 14.0, 4.2 Hz, H-6″b), 3.57 (1H, dd, J = 12.0, 4.5 Hz, H-6″a), 3.49 (1H, d, J = 11.4 Hz, H-3″), 3.40 (1H, d, J = 11.4 Hz, H-2″), 3.27 (1H, d, J = 7.8 Hz H-5″). 13C-NMR (125 MHz, DMSO-d6) δ: 178.3 (C-4), 163.6 (C-7), 161.8 (C-2), 160.2 (C-5), 157.6 (C-9), 157.3 (C-4'), 134.1 (C-3), 130.9 (C-2', C-6'), 121.2 (C-1'), 114.7 (C-3', C-5'), 104.8 (C-10), 103.5 (C-1″), 98.5 (C-6), 93.3 (C-8), 75.7 (C-5″). 73.8 (C-3″), 71.6 (C-2″), 68.6 (C-4″), 60.5 (C-6″) (Fig. S15 and S16).
Compound 9: Yellow powder; ESI–MS: m/z 301.1 [MH]- (C15H10O7). 1H-NMR (DMSO-d6, 500 MHz) δ: 7.69 (1H, d, J = 2.1 Hz, H-2'), 7.60 (1H, dd, J = 8.4, 2.1 Hz, H-6′), 6.88(1H, d, J = 8.4 Hz, H-5'), 6.46 (1H, d, J = 2.0 Hz, H-8), 6.14 (1H, d, J = 2.0 Hz, H-6). 13C-NMR (125 MHz, DMSO-d6) δ: 175.9 (C-4), 163.3 (C-7), 161.3 (C-9), 157.8 (C-5), 148.3 (C-4'), 146.5 (C-1), 143.0 (C-1'), 135.8 (C-3) 122.7 (C-1'), 120.2 (C-6'), 115.0 (C-5'), 113.5 (C-2'), 103.3 (C-10), 98.0 (C-6), 93.1 (C-8) (Fig. S17 and S18).
Compound 10: Yellow powder; ESI–MS: m/z 270 [MH]−. 1H-NMR (DMSO-d6, 500 MHz) δ: 7.41(1H, dd, J = 8.2, 2.2 Hz, H-6'), 7.50(1H, d, J = 2.2 Hz, H-2′), 6.90(1H, d, J = 8.3 Hz, H-5'), 6.68(1H, s, H-3), 6.45(1H, d, J = 2.0 Hz, H-8), 6.21(1H, d, Hz, H-6); 13C-NMR(125 MHz, DMSO-d6) δ: 181.7(C-4), 164.1(C-2), 163.9(C-7), 162.4(C-5), 156.2(C-9), 149.7(C-4′), 145.7(C-3′), 121.6(C-1′), 118.9(C-6′), 116.0(C-5′), 113.3(C-2′), 103.7 (C-10), 98.8(C-6), 93.8(C-8) (Fig. S19 and S20).
Compound 11: Yellow powder; ESI–MS: m/z 270 [MH]. 1H-NMR (DMSO-d6, 500 MHz): 7.83 (2H, d, J = 8.8 Hz, H-2', H-6'), 6.93 (2H, d, J = 8.8 Hz, H-3′, H-5′), 6.83 (1H, d, J = 2.0 Hz, H-6), 6.71 (1H, d, J = 2.0 Hz, H-8), 6.57 (1H, s, H-3); 13C-NMR(125 MHz, DMSO-d6): 180.4 (C-4), 165.9 (C-5), 164.4 (C-2), 162.5 (C-4′), 160.7 (C-9), 160.1 (C-7), 129.3 (C-2′, C-6′), 123.2 (C-10), 117.1 (C-3′, C-5′), 109.3 (C-10), 106.6 (C-3), 104.7 (C-6), 99.3 (C-8) (Fig. S21 and S22).
Compound 12: Yellowish white powder; EI-MS m/z 330.0740 [M]+; 1H-NMR (500 MHz, DMSO-d6) δ 12.97 (1H, 5-OH), H-7.32 (2H, s, H-2′, H-6′) 6.98 (1H, s, H-3), 6.56 (1H, d, J = 3.0 Hz, H-8), 6.21 (1H d, J = 3.0 Hz, H-6), 3.89 (6H, s, 3′, 5′-OCH3); 13C-NMR (125 MHz, DMSO-d6) δ 182.1 (C-4), 164.6 (C-7), 164.1 (C-2), 161.8 (C-5), 157.7 (C-8a), 148.6 (C-3′, C-5′), 140.2 (C-4′), 120.8 (C-1′), 104.7 (C-2′, C-6′), 104.0 (C-4a, C-3), 99.3 (C-6), 94.7 (C-8), 56.8 (C-OCH3 × 2) (Fig. S23 and S24).
Compound 13: Yellow powder; EI-MS m/z [M + H]+ m/z 287.0546. 1H-NMR (500 MHz, DMSO-d6) δ: 12.48 (1H, s, 5-OH), 10.84 (1H, s, 7-OH), 10.13 (1H, s, 4′-OH), 8.04 (2H, d, J = 9.0 Hz, H-2′, H-6′), 6.95 (2H, d, J = 9.0 Hz, H′3′, H-5′), 6.45 (1H, d, J = 2.0 Hz, H-8), 6.22 (1H, d, J = 2.2 Hz, H-6). 13C-NMR (125 MHz, DMSO-d6) δ: 176.4 (C-4), 164.4 (C-7), 161.1 (C-5), 159.7 (C-4′), 156.6 (C-9), 147.2 (C-2), 136.1 (C-3), 130.0 (C-2′, C-6′), 122.2 (C-10), 116.0 (C-3′, C5), 103.5 (C-1′), 96.7 (C-6), 94.0 (C-8) (Fig. S25 and S26).
Preparation of samples for HPLC and calibration curves
Two types of CE are cultivated in Hampyeong and Ganghwa, and both were used to prepare MeOH extracts (50 mg/mL). The same sample-preparation method was used to prepare MeOH extracts of four Cyperus species. Standard flavonoid and phenolic acid solutions were also prepared by dissolving the respective compounds in MeOH (0.5 mg/mL). Before analysis, all samples were filtered through a syringe filter (0.45 µm, PVDF) using sonication for 20 min. The compounds were diluted in series to produce different concentrations (0.03125–0.5 mg/mL). The calibration curve for each standard was constructed by plotting the concentration (X, µg/mL) versus the peak area (Y). The correlation coefficient (r2) was used to determine linearity.
HPLC–DAD chromatographic condition
Chromatographic separation of individual compounds was performed using the INNO C18 column (250 × 4.6 mm, 5 µm). The mobile phase was 0.5% trifluoroacetic acid in 10% ACN (0.5:90:10, TFA/ACN/water) (sol A) and 0.5% TFA in 90% ACN (0.5:90:10, TFA/ACN/water) (sol B). The gradient elution system was as follows: 0 min, 90% A; 15 min, 90% A; 18 min, 83% A; 30 min, 80% A; 35 min, 70% A; 55 min, 20% A; 58 min, 100% B; 63 min, 100% B; 65 min, 90% A; and 70 min, 90% A. The injection volume was 10 µL, and the flow rate was 0.8 mL/min. The UV detection wavelength was 270 nm. All injections were performed three times.
Results and discussion
Identification of phytochemical constituents from CE
The results for total polyphenol content and the ABTS and DPPH radical-scavenging activities are shown in Table 1. The EtOAc and n-BuOH fractions showed the highest radical-scavenging activity.
For fractions with a high polyphenol content and antioxidant activity, phytochemical isolation was conducted using open-column chromatography. Recrystallized phytochemicals from the EtOAc and n-BuOH fractions of CE were identified by nuclear magnetic resonance (NMR) spectroscopy and mass spectrometry (MS). Spectroscopic data from the isolated compounds showed characteristic signals of flavonoids and phenolic acids. Thus, 13 compounds (Fig. 1), namely, gallic acid (1) , protocatechuic acid (2) , vanillic acid (3) , p-coumaric acid (4) , rutin (5) , ferulic acid (6) , isoquercitrin (7) , astragalin (8) , quercetin (9) , luteolin (10) , apigenin (11) , tricin (12) , and kaempferol (13)  were identified using previously published literature. Although none of the 13 compounds isolated from CE are new, this is the first report to describe their isolation from CE. Therefore, our results can be used as a reference for further research on CE.
Antioxidant activities of compounds from CE
The DPPH and ABTS assays were conducted to evaluate the antioxidant ability of the 13 isolated compounds, and the IC50 values were compared with that of ascorbic acid, a representative antioxidant. The results are shown in Table 2. Overall, phenolic acid tended to show higher antioxidant ability than flavonoids. In the DPPH assay, vanillic acid, p-coumaric acid, and quercetin demonstrated stronger radical-scavenging activity than ascorbic acid. In particular, vanillic acid was the most effective DPPH radical scavenger from CE. In the ABTS assay, most phenolic acids, quercetin, and luteolin exhibited better radical-scavenging activity than ascorbic acid, and gallic acid and quercetin exhibiting the greatest ability to quench the ABTS radicals (Tables 2, 3).
Eight flavonoids and five phenolic acids isolated from CE were analyzed quantitatively using HPLC–DAD. Quantitative analysis was also conducted to compare CE grown in Ganghwa and Hampyeong, which are the two main CE-growing regions. The compound peaks were successfully separated with the corresponding retention times with high resolution (Fig. 2). Good linearity was obtained within the tested concentration range with a correlation coefficient (r2) of 0.9990–1.0000.
In general, flavonoids are used as marker compounds to evaluate the pharmacological value of plant materials. Flavonoids have been reported to be mainly responsible for biological activity . Quantitative analysis of flavonoids and phenolic acid in CE indicated that their contents varied notably by the region of production (Table 4).
In CE from Hampyeong and Ganghwa, astragalin (4.72 mg/g and 2.85 mg/g, respectively) and tricin (3.00 mg/g and 2.08 mg/g, respectively) were the main flavonoids, and p-coumaric acid (4.30 mg/g and 3.39 mg/g, respectively) was the major phenolic acid in the EtOH extract of CE. Overall, the Hampyeong CE had a high phytochemical content, but in terms of ratio, it exhibited a similar pattern to that of Ganghwa CE. The growing environment can be assumed to influence differences in the contents of the active compounds of CE, but it does not change the fundamental constituents (Fig. 3). These findings are consistent with the results of previous studies in which the production of active substances did not differ significantly in the absence of extreme environmental changes .
In addition, the existing component analysis studies of CE and its related species have provided limited data. Therefore, a component analysis of 13 chemicals isolated from CE was performed for four species (MeOH extracts of C. difformis, C. microiria, C. sanguinolentus, and C. cyperoides) including CE. Astragalin was not included, but phytochemicals such as quercetin, isoquercitrin, and luteolin, which were isolated from CE, were also included in the four Cyperus species extracts (Table 5).
The results of the content analysis showed that the CE extract had the highest content of phenolic acids and flavonoids among the extracts of the five Cyperus species (Fig. 4). In the literature, C. difformis and several Cyperus species have been reported to contain high levels of phenolic compounds, particularly luteolin .
Thirteen compounds were identified from the EtOAc and n-BuOH fractions of CE. Most of the isolated compounds have been reported to have various biological activities. Among the isolated compounds, astragalin and isoquercitrin were isolated from the genus Cyperus for the first time in the present study; similarly, the remaining 11 compounds have not been reported to have been isolated from CE. An HPLC analytical method that can simultaneously analyze 13 compounds isolated from CE was established, and a content comparison of CE by the region of production and a comparative analysis of four related species were conducted. This study demonstrated the academic value of CE, which has not been sufficiently explored. Furthermore, our findings may serve as a reference for using CE as functional material to treat oxidative stresses as well as for expanding our understanding of its antioxidant properties and phytochemical components (Additional file 1: Figs. S1–S26).
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This research was supported by grants from the Natural Product Institute of Science and Technology (2021), Anseong, Korea and the Basic Science Research Program through the National Research Foundation of Korea (NRF) funded by the Ministry of Education (2018R1A6A1A03025159).
On behalf of all authors, the corresponding author states that there is no competing interests.
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Fig. S1. 1H-NMR spectrum of compound 1 (Gallic acid). Fig. S2. 13C-NMR spectrum of compound 1 (Gallic acid). Fig. S3. 1H-NMR spectrum of compound 2 (Protocatechuic acid). Fig. S4. 13C-NMR spectrum of compound 2 (Protocatechuic acid). Fig. S5. 1H-NMR spectrum of compound 3 (Vanillic acid). Fig. S6. 13C-NMR spectrum of compound 3 (Vanillic acid). Fig. S7. 1H-NMR spectrum of compound 4 (P-coumaric acid). Fig. S8. 13C-NMR spectrum of compound 4 (P-coumaric acid). Fig. S9. 1H-NMR spectrum of compound 5 (Rutin). Fig. S10. 13C-NMR spectrum of compound 5 (Rutin). Fig. S11. 1H-NMR spectrum of compound 6 (Ferulic acid). Fig. S12. 13C-NMR spectrum of compound 6 (Ferulic acid). Fig. S13. 1H-NMR spectrum of compound 7 (Isoquercetin). Fig. S14. 13C-NMR spectrum of compound 7 (Isoquercetin). Fig. S15. 1H-NMR spectrum of compound 8 (Astragalin). Fig. S16. 13C-NMR spectrum of compound 8 (Astragalin). Fig. S17. 1H-NMR spectrum of compound 9 (Quercetin). Fig. S18. 13C-NMR spectrum of compound 9 (Quercetin). Fig. S19. 1H-NMR spectrum of compound 10 (Luteolin). Fig. S20. 13C-NMR spectrum of compound 10 (Luteolin). Fig. S21. 1H-NMR spectrum of compound 11 (Apigenin). Fig. S22. 13C-NMR spectrum of compound 11 (Apigenin). Fig. S23. 1H-NMR spectrum of compound 12 (Tricin). Fig. S24. 13C-NMR spectrum of compound 12 (Tricin). Fig. S25. 1H-NMR spectrum of compound 13 (Kaempferol). Fig. S26. 13C-NMR spectrum of compound 13 (Kaempferol).
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Choi, J., Lee, HD., Cho, H. et al. Antioxidative phenolic compounds from the aerial parts of Cyperus exaltatus var. iwasakii and their HPLC analysis. Appl Biol Chem 66, 61 (2023). https://doi.org/10.1186/s13765-023-00820-3