Antioxidant and Antidiabetic Activities of Flavonoid Derivatives from the Outer Skins of Allium cepa L.

Article abstract of DOI:10.1021/acs.jafc.0c02122

The onion, known as the bulb onion or common onion, is not only a key ingredient in many tasty and healthy vegetarian meals but also many traditional medicines. Nine new flavonoids [cepaflavas A, B (5, 6), cepadials A-D (7-9 and 14), and cepabiflas A-C (10-12)] and six known compounds (1-4, 13, 15) were obtained from the outer skins of Allium cepa L. Among them, compounds 5, 6, and 9 might be artificial products formed during extraction and isolation. New compounds were structurally elucidated using various spectroscopy/spectrometry techniques, including NMR and HRMS, and computational methods. Their absolute configurations were determined using time-dependent density functional theory calculations, combined with ECD spectroscopy, optical rotation calculation, and statistical procedures (CP3 and DP4 analysis). The free radical scavenging assays revealed that the new compounds 10-12 possessed considerable antioxidant activities with IC50 values of 4.25-8.88 and 7.12-8.14 μM against DPPH and ABTSa +, respectively. Compounds 13-15 showed substantial inhibitory activities against both α-glucosidase and protein tyrosine phosphatase 1B (PTP1B), with IC50 values of 0.89-6.80 and 1.13-6.82 μM, respectively. On the basis of molecular docking studies, 13 and 15 were predicted to have high binding capacity and strong affinity toward the active site of PTP1B.

Full text of DOI:10.1021/acs.jafc.0c02122

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Bioactive Constituents, Metabolites, and Functions
Antioxidant and antidiabetic activities of flavonoid
derivatives from the outer skins of Allium cepa L.
Ngoc Khanh Vu, Chung Sub Kim, Manh Tuan Ha, Thi Quynh Mai Ngo, Se Eun
Park, Haeun Kwon, Dongho Lee, Jae Sue Choi, Jeong Ah Kim, and ByungSun Min
J. Agric. Food Chem., Just Accepted Manuscript • DOI: 10.1021/acs.jafc.0c02122 • Publication Date (Web): 30 Jun 2020
Downloaded from pubs.acs.org on June 30, 2020
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Antioxidant and antidiabetic activities of flavonoid derivatives from the outer skins
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of Allium cepa L.
†
,⊥
‡,⊥
†
§
‖
Ngoc Khanh Vu, Chung Sub Kim, Manh Tuan Ha, Quynh-Mai Thi Ngo, Se Eun Park, Haeun
‖
*,¶
*,†
∇
∇
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6
Kwon, Dongho Lee, Jae Sue Choi, Jeong Ah Kim, and Byung Sun Min
†
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College of Pharmacy, Drug Research and Development Center, Daegu Catholic University, Gyeongbuk
38430, Republic of Korea
‡
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0
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School of Pharmacy, Sungkyunkwan University, Suwon 16419, Republic of Korea
§
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College of Pharmacy, Hai Phong University of Medicine and Pharmacy, 72A Nguyen Binh Khiem, Hai
Phong 180000, Viet Nam
∇Department of Biosystems and Biotechnology, College of Life Sciences and Biotechnology, Korea
University, Seoul 136-713, Republic of Korea
‖
14
Department of Food and Life Science, Pukyong National University, Busan 48513, Republic of Korea
¶
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College of Pharmacy, Research Institute of Pharmaceutical Sciences, Kyungpook National University,
Daegu 41566, Republic of Korea
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ABSTRACT: The onion, known as the bulb onion or common onion, is not only key ingredients in
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many tasty and healthy vegetarian meals but also many traditional medicines. Nine new flavonoids
[cepaflavas A, B (5, 6), cepadials A–D (7–9, and 14), and cepabiflas A–C (10–12)] and six known
compounds (1–4, 13, 15) were obtained from the outer skins of Allium cepa L.. Among them, compounds
5, 6, and 9 might be artificial products formed during extraction and isolation. New compounds were
structurally-elucidated using various spectroscopy/spectrometry techniques, including NMR and HRMS,
and computational methods. Their absolute configurations were determined using time-dependent
density functional theory calculations, combined with ECD spectroscopy, optical rotation calculation,
and statistical procedures (CP3 and DP4 analysis). The free radical scavenging assays revealed that the
new compounds 10–12 possessed considerable antioxidant activities with IC values of 4.25–8.88 and
50
•
+
7.12–8.14 μM against DPPH and ABTS , respectively. Compounds 13–15 showed substantial inhibitory
activities against both α-glucosidase and protein tyrosine phosphatase 1B (PTP1B), with IC values of
50
0.89–6.80 and 1.13–6.82 μM, respectively. Based on molecular docking studies, 13 and 15 were
predicted to have high binding capacity and strong affinity toward the active site of PTP1B.
KEYWORDS: Allium cepa skins, antioxidant, α-glucosidase, PTP1B, kinetic, molecular docking
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INTRODUCTION
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Allium genus is a member of the Alliaceae family, formerly considered a part of Liliaceae, which includes
more than 700 species widely distributed through temperate and warm temperate climate and boreal
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zones of the northern hemisphere. Red onion (Allium cepa L.) is a perennial plant which has opposite
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leaves and globe-shaped bulbs formed from several leaf bases encased in thin, dry skins. Owing to a
typical pungent smell associated with sulfur-containing compounds, and its rich source of several
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phytonutrients, onion is an essential kitchen vegetable in many dishes. It is also a folk remedy for various
1
diseases of the heart, and headaches, bites, worms, and tumors. According to prior investigations, A.
cepa possesses various biological activities including anticancer,4 antioxidant,5 antiplatelet,6
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antidiabetic, and anti-inflammatory activities. Among the diverse secondary metabolites in A. cepa,
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sapogenins, saponins, and flavonoids are known as the major components. Recently, Li et al.
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concentrated total flavonoids from onion peels and obtained 90.25% flavonoids content extract. The
outer skins of onion, a waste product from the food industry, is a natural source contains high-value
functional ingredients, should be searched for possible ways for their utilization.
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Along with cancer, cardiovascular, and chronic lung diseases, diabetes is one of the four priority
noncommunicable diseases. Diabetes, which accounts for 1.6 million deaths in 2015, has emerged as a
major public health problem.11, 12 Not only is diabetes a serious chronic disorder that is the main cause
of kidney failure, heart attack, blindness, and stroke, but it is also a global economic burden, incurring
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up to 1.31 trillion USD in 2015 alone. Type 2 diabetes comprises more than 90% of all diabetes cases,
usually occurs in adults, and is strongly related to overweight, obesity, and physical inactivity.12, 14 The
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body does not use insulin properly in type 2 diabetes, causing abnormally high blood glucose levels. In
general, increased levels of glucose induce insulin secretion by releasing insulin from pancreatic β-
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cells. Insulin binds to its receptor at the plasma membrane, activating an intrinsic tyrosine-protein
kinase, which results in receptor autophosphorylation on tyrosine residues and triggers downstream
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signaling events.15 By tyrosine dephosphorylation of the insulin substrate and its receptor, PTP1B
disrupts insulin signaling pathways.16, 17 Thus, PTP1B is considered a negative regulator of the leptin and
insulin signaling pathways that are directly related to obesity and diabetes.17 In vitro, PTP1B
overexpression in rat adipose cells dramatically impaired insulin-stimulated translocation of GLUT4.18
Moreover, in a mouse model, knockout of the PTP1B gene ameliorated insulin sensitivity and promoted
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weight loss on a high-fat diet. Therefore, compounds that inhibit PTP1B are promising therapeutic
agents to treat obesity and related metabolic disorders. Another approach for controlling postprandial
hyperglycemia is to reduce the carbohydrate absorption rate in the gastrointestinal tract by inhibiting α-
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glucosidase. In the brush border of the small intestine, α-glucosidase inhibitors bind competitively to a
carbohydrate-binding region of α-glucosidase enzymes, which consequently delays carbohydrate
breakdown and alleviates postprandial hyperglycemia.20 However, marketed drugs targeting α-
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glucosidase enzymes cause some side effects on gastrointestinal system. Thus, new α-glucosidase
inhibitors need to be discovered for the development of antidiabetic drugs. In this work, phytochemical
investigations of the outer skins of A. cepa were undertaken to discover more naturally occurring
compounds with undescribed structures and their anti-diabetic activities.
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MATERIALS AND METHODS
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General Experiment Procedures. A JASCO P-2000 digital polarimeter (JASCO Corporation,
Tokyo, Japan) was used for measuring optical rotations. UV and electronic circular dichroism (ECD)
spectra were obtained using a J-1500 circular dichroism spectrophotometer (JASCO Corporation, Tokyo,
Japan). IR spectra were measured by a JASCO FT/IR-4100 spectrometer (JASCO Corporation, Tokyo,
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Japan). Varian Unity Inova 400 MHz (ASW probe, 400 MHz for H NMR, 100 MHz for C NMR)
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(Varian, Inc., California, USA) and Bruker Ascend 500 MHz (Flow probe, 500 MHz for H NMR,
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125 MHz for C NMR) (Buker Corporation, Massachusetts, USA) spectrometers were used to acquire
the NMR spectra. High-resolution electron spray ionization-mass spectrometry (HRESIMS) was
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performed using an AB SCIEX TripleTOF 5600 spectrometer (AB Sciex Pte. Ltd., Ontario, Canada).
High-resolution electron impact-mass spectrometry (HREIMS) was undertaken using a JEOL JMS-AX
300L spectrometer (Jeol, Akishima, Tokyo, Japan). RP-18 F254s and Merck precoated silica gel F254 plates
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were used to perform thin-layer chromatography (TLC). (RP)-C (Merck, 75 mesh) and silica gel 60
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(Merck, 230–400 mesh) were used for column chromatography (CC). Preparative high-performance
liquid chromatography (HPLC) was carried out using a YMC Pak ODS-A column (20 × 250 mm, 5 μm
particle size; YMC Co., Ltd., Japan) and a Waters 2487 controller system with a UV detector (UV/VIS
- 156).
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Plant Materials. The outer skins used in this study were obtained from red onion (A. cepa) purchased
at the Central Market of Daejeon, Republic of Korea, in May 2017, and identified by Professor Byung
Sun Min (one of the authors). A voucher specimen (CUD-0814-1) was deposited at the Herbarium of the
College of Pharmacy, Daegu Catholic University, Korea.
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Extraction and Isolation. Air-dried outer skins of A. cepa (10 kg) were extracted using MeOH (10
01 L × 3), and the extracts were evaporated in vacuo until dryness. The MeOH extract (0.8 kg) was dissolved
02 in water (1 L) and successively partitioned with n-hexane, CH Cl , EtOAc, and n-BuOH. The EtOAc-
2
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03 soluble fraction (405 g) was subjected to silica gel CC (CH Cl :MeOH, 10:1, v/v) to generate 30 fractions
2 2
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04 (E1–E30). Fraction E23 (1.0 g) was chromatographed on a silica gel column (n-hexane:EtOAc, 20:1,
05 v/v) to yield 2 (50 mg). Fraction E30 (80 g) was fractionated by silica gel CC (n-hexane:EtOAc, 3:1, v/v)
06 to yield 15 subfractions (E30.1–E30.15). Fraction E30.5 (2.5 g) was purified using RP-18 silica gel CC
07 (MeOH:H O, 1:2, v/v) to afford 1 (150 mg). Compound 5 (10 mg) and 6 (15 mg) were obtained from
2
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08 fraction E30.4 (510 mg) using a silica gel column (n-hexane:acetone, 2:1, v/v). Fraction E30.8 (3 g) was
09 fractionated by silica gel CC (CH Cl :MeOH, 20:1, v/v) to produce 20 subfractions (E30.8.1–E30.8.20).
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10 Fraction E30.8.8 (136 mg) was separated using a RP-18 silica gel column (MeOH:H O, 1:1, v/v) to
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11 obtain 3 (10 mg) . Fraction E30.11 (6 g) was subjected to silica gel CC (n-hexane:acetone, 1:1, v/v) to
12 yield 4 (50 mg) and 13 (80 mg). Further purification of E30.11.18 (450 mg) by HPLC using MeOH:H O
2
13 (60:40, v/v) at a flow rate of 5 mL/min as the mobile phase, yielded 10 (3 mg), 11 (4 mg), and 12 (6 mg).
14 Fraction E30.12 (2.5 g) was fractionated by silica gel CC (CH Cl :MeOH, 15:1, v/v) to yield 20
2
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15 subfractions (E30.12.1–E30.12.20). Compounds 7 (3 mg), 8 (6 mg), 9 (4 mg), 14 (10 mg), and 15 (20
16 mg) were isolated from subfraction E30.12.6 (450 mg) by HPLC using MeOH:H O (55:45, v/v) as the
2
17 mobile phase at a flow rate of 5 mL/min.
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Cepaflava A (5): pale yellow, amorphous powder; [α] D –17.8 (c 0.08, MeOH); UV (MeOH) λmax
19 (log ε) 291 (2.28), 252 (1.75), 198 (2.57) nm; ECD (MeOH) λmax (Δε) 200 (2.89), 234 (–0.01), 261 (0.20),
1
13
20 299 (–0.18); IR (KBr) vmax 3324, 2941, 2831, 1637, 1448, 1024; H (400 MHz, methanol-d
4
) and C
+
21 NMR (100 MHz, methanol-d
4
) data, see Table 1; HREIMS m/z 390.0948 ([M] , calcd for C H O ,
1
9
18
9
22 390.0951).
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Cepaflava B (6): pale yellow, amorphous powder; [α] D –28.5 (c 0.09, MeOH); UV (MeOH) λmax
24 (log ε) 290 (2.31), 252 (1.69), 200 (2.66) nm; ECD (MeOH) λmax (Δε) 200 (1.40), 236 (–0.35), 260 (–
1
25 0.17), 293 (–0.61); IR (KBr) vmax 3318, 2942, 2832, 1636, 1448, 1116, 1024; H (400 MHz, methanol-
1
3
+
1
1
26
d
4
) and C NMR (100 MHz, methanol-d ) data, see Table 1; HREIMS m/z 390.0954 ([M] , calcd for
4
27 C H O 390.0951).
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Cepadial A (7): brown, amorphous powder; [α] D –24.6 (c 0.09, MeOH); UV (MeOH) λmax (log
29 ε) 292 (2.46), 268 (2.20), 209 (2.69) nm; ECD (MeOH) λmax (Δε) 299 (0.03), 263 (0.29), 243 (0.25), 203
1
13
30 (3.04); IR (KBr) vmax 3324, 2945, 2831, 1733, 1450, 1116, 1036; H (500 MHz, methanol-d
4
) and C
+
1
1
31 NMR (125 MHz, methanol-d
4
) data, see Table 2; HRESIMS m/z 477.0436 ([M + Na] , calcd for
32 C H O Na, 477.0434).
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Cepadial B (8): brown, amorphous powder; [α] D –8.5 (c 0.08, MeOH); UV (MeOH) λmax (log ε)
34 292 (2.32), 268 (2.05), 209 (2.54) nm; ECD (MeOH) λmax (Δε) 299 (–0.51), 261 (–0.14), 248 (–0.21),
1
35 203 (3.94); IR (KBr) vmax 3324, 2945, 2831, 1733, 1450, 1116, 1036; H (500 MHz, methanol-d
4
) and
1
3
+
1
1
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C NMR (125 MHz, methanol-d ) data, see Table 2; HRESIMS m/z 477.0436 ([M + Na] , calcd for
4
37 C H O Na, 477.0434).
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Cepadial C (9): brown, amorphous powder; [α] D –10.2 (c 0.10, MeOH); UV (MeOH) λmax (log
39 ε) 290 (2.58), 252 (2.31), 200 (2.80) nm; ECD (MeOH) λmax (Δε) 295 (–0.95), 206 (4.62); IR (KBr) vmax
1
13
1
1
1
40 3324, 2945, 2831, 1733, 1450, 1116, 1036; H (500 MHz, methanol-d
4
) and C NMR (125 MHz,
+
41 methanol-d
4
) data, see Table 2; HRESIMS m/z 491.0588 ([M + Na] , calcd for C H O Na, 491.0590)
2
3
16 11
+
42 and m/z 469.0763 ([M + H] , calcd for C H O 469.0771).
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,
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Cepabifla A (10): bronze-yellow, amorphous powder; [α] D –26.3 (c 0.10, MeOH); UV (MeOH)
44 λmax (log ε) 299 (2.51), 260 (2.46), 203 (2.90) nm; ECD (MeOH) λmax (Δε) 319 (7.37), 288 (–2.90), 254
1
45 (0.04), 234 (–2.44); IR (KBr) vmax 3316, 2946, 2831, 1450, 1112, 1024; H (400 MHz, methanol-d
4
) and
1
3
+
1
1
46
C NMR (100 MHz, methanol-d
4
) data, see Table 3; HRESIMS m/z 787.1119 ([M + Na] , calcd for
+
47 C H O Na, 787.1122), and m/z 765.1329 ([M + H] , calcd for C H O 765.1303).
3
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28 19
36 29 19
,
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1
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Cepabifla B (11): bronze-yellow, amorphous powder; [α] D +60.3 (c 0.1, MeOH); UV (MeOH)
49 λmax (log ε) 303 (2.39), 255 (2.40), 201 (2.85) nm; ECD (MeOH) λmax (Δε) 304 (7.02), 252 (–1.40), 239
1
50 (–2.32), 218 (2.10); IR (KBr) vmax 3316, 2946, 2831, 1450, 1112, 1024; H (400 MHz, methanol-d
4
) and
1
3
+
1
1
51
C NMR (100 MHz, methanol-d
4
) data, see Table 3; HRESIMS m/z 787.1105 ([M + Na] , calcd for
+
52 C H O Na, 787.1122), and m/z 765.1329 ([M + H] , calcd for C H O 765.1303).
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36 29 19
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Cepabifla C (12): bronze-yellow, amorphous powder; [α] D –30.0 (c 0.1, MeOH); UV (MeOH) λmax
54 (log ε) 303 (2.38), 255 (2.40), 201 (2.83) nm; ECD (MeOH) λmax (Δε) 304 (–4.79), 276 (1.79), 251 (0.69),
1
55 239 (1.28), 223 (–2.14), 199 (3.05); IR (KBr) vmax 3316, 2946, 2831, 1450, 1112, 1024; H (400 MHz,
1
3
1
1
56 methanol-d
4
) and C NMR (100 MHz, methanol-d
4
) data, see Table 3; HRESIMS m/z 787.1115 ([M +
+
+
57 Na] , calcd for C H O Na, 787.1122), and m/z 765.1329 ([M + H] , calcd for C H O 765.1303).
3
6
28 19
36 29 19
,
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1
1
1
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Cepadial D (14): bronze-yellow, amorphous powder; [α] D –23.5 (c 0.1, MeOH); UV (MeOH) λmax
59 (log ε) 299 (2.53), 264 (2.26), 202 (2.87) nm; ECD (MeOH) λmax (Δε) 303 (0.26), 269 (0.49), 241 (0.43),
1
13
60 203 (3.15); IR (KBr) vmax 3324, 2945, 2832, 1684, 1449, 1037; H (500 MHz, methanol-d
4
) and C
+
61 NMR (125 MHz, methanol-d
4
) data, see Table 1; HRESIMS m/z 791.0848 ([M + Na] , calcd for
+
62 C H O Na, 791.0860), and m/z 769.1047 ([M + H] , calcd for C H O 769.1041).
38
24 18
38 25 18
,
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Acid Hydrolysis of 10–12. For hydrolyzation, 1 mg of each compound was weighed, then 3 mL of
°
64 2N CF COOH was added and boiled for 3 h at 95 C. Methylene chloride (3 × 3 mL) was used to partition
3
65 the solution. The water layer was dried, analyzed by TLC (MeOH:CHCl :H O, 5:8:1), and then compared
3
2
66 with the standard. Preparative TLC (MeOH:CHCl :H O, 5:8:1) was used to yield the monosaccharide
3
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2
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67 part. The sugar moiety was verified as D-glucose (positive [α] ) by measuring the optical rotation.
D
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Computational Method. Systematic conformational searches for 5, 6, 11, and 12 were accomplished
69 under the molecular mechanics force field (MMFF) in Spartan'14 software (Wavefunction, Inc., Irvine,
70 CA, USA). The appropriate conformers were selected for optimization by density functional theory
71 (DFT) calculations at the B3LYP/6-31+G(d,p) level using the Gaussian 09 package (Gaussian, Inc.,
72 Wallingford, CT, USA), and time-dependent DFT ECD calculations of the optimized conformers were
73 carried out at the CAM-B3LYP/TZVP level with a conductor polarized continuum (CPCM) solvent
74 model in MeOH. The calculated ECD spectra of different conformers were simulated with a half
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75 bandwidth of 0.2−0.3 eV and ECD curves were generated by SpecDis 1.64 software. The ECD spectra
76 of 5, 6, 11, and 12 were weighted by Boltzmann distribution after UV correction.
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Conformers of 5−10 and 14 used in this study were found using “Mixed torsional/Low-mode
78 sampling” in the MMFF force field with the MacroModel (version 2015-2; Schrödinger LLC, New York,
79 NY, USA) module. To explore all potential conformers, the gas phase with a 5 kJ/mol energy window
80 limit and 10,000 maximum number of steps were implemented. To minimize conformers, the Polak–
81 Ribière conjugate gradient method was carried out with 10,000 iterations and a 0.001 kJ (mol Å)−1
82 convergence threshold on the root mean square (RMS) gradient. For CP3 (for 5−8) and DP4 (for 14)
83 analysis, chemical shifts of the conformers were calculated without geometry optimization at the
84 mPW1PW91/6-311G* level in the polarizable continuum model (PCM; MeOH) using the Gaussian 16
85 package (Gaussian Inc.). The calculated NMR properties were averaged based upon their respective
86 Boltzmann populations and analyzed by the applets available at http://www-jmg.ch.cam.ac.uk/tools/nmr
87 for CP3 and DP4 analysis. For the ECD calculation, all the conformers were geometry-optimized in the
88 gas phase at the BP86/SVP level and subjected to the calculation of excitation energies, rotatory strength,
89 and oscillator strength at the BP86/aug-cc-pVDZ level in the PCM (MeOH). Based on the calculated
90 Gibbs free energy of each conformer and visualized utilizing SpecDis software (version 1.71) with a σ/γ
91 value of 0.6 eV, the ECD spectra were Boltzmann-averaged. The molecular energies for the cis- and
92 trans- form of 7 were calculated at MM2 level from the ChemDraw 19.0 and B3LYP/6-31+G(d,p) level
93 for the conformer with the lowest energy after a conformational search using Macromodel.
1
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DPPH Radical Scavenging Assay. This assay was conducted using the previously described method
95 (Supporting Information).22
•
+
•+
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96
Peroxyl (ABTS ) Radical Scavenging Assay. ABTS radical cation was produced by reacting
97 ABTS with potassium persulfate following the method previously described (Supporting Information).22
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98
PTP1B Inhibitory Assay. PTP1B inhibitory assays were conducted using p-nitrophenyl phosphate
99 (p-NPP) as the substrate following the protocol previously published (Supporting Information).23
2
2
2
2
00
α-Glucosidase Inhibitory Assay. By applying the previously published procedure,24 the α-
01 glucosidase inhibition was assayed using p-nitrophenyl α-D-glucopyranoside (p-NPG) as the substrate.
02 The release of p-NPG was measured at 405 nm using a microplate spectrophotometer (Molecular
03 Devices) for determining the α-glucosidase activity. Acarbose served as a positive control.
2
2
2
04
Enzyme Kinetic Analysis with PTP1B. Lineweaver–Burk and Dixon plots were utilized for
05 determining the mechanism by which the most active compounds 13–15 inhibit PTP1B.25-27 The PTP1B
06 enzymatic procedures have been described previously (Supporting Information).23
2
2
2
2
07
Molecular Docking Simulation in PTP1B Inhibition. Molecular docking simulations using
08 AutoDock 4.2 software were implemented to clarify the binding mode and interaction of the active
09 compounds (13 and 15) with PTP1B. The docking protocol was conducted with the AutoDock 4.2
10 program following the method previously described with modifications (Supporting Information).
211 RESULTS AND DISCUSSION
2
2
2
2
2
2
2
12
Phytochemical Study. The outer skins of A. cepa were extracted using MeOH to obtain a crude
Cl , EtOAc, and n-BuOH. The EtOAc
13 extract, which was partitioned successively with n-hexane, CH
2
2
14 fraction (405 g) was separated over silica gel and HPLC to obtain nine new flavonoids [cepaflavas A, B
15 (5, 6), cepadials A–D (7–9, and 14), and cepabiflas A–C (10–12)] and six known compounds (1–4, 13,
16 15) (Figure 1).
17
Cepaflava A (5) was isolated as a pale yellow, amorphous powder with a molecular formula of
18 C H O , which was deduced from the HREIMS that displayed a molecular ion peak at m/z 390.0948
19
18
9
+
2
2
19 [M] (calcd for C H O 390.0951). The UV spectrum exhibited absorption maxima at 198, 252, and
19 18 9
,
1
−
1
−1
20 291 nm. The IR spectrum displayed the hydroxyl (3324 cm ), carbonyl groups (1637 cm ). The H and
1
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2
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2
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21
22 J = 8.2, 1.9 Hz, H-6'), and 6.80 (1H, d, J = 8.2 Hz, H-5')], two meta-coupled doublets [δ
23 = 2.0 Hz, H-8) and 5.94 (1H, d, J = 2.0 Hz, H-6)], and a carbonyl signal at δ 196.4 (C-4), which were
C NMR spectra of 5 showed an ABX coupling system [δ
H
7.01 (1H, d, J = 1.9 Hz, H-2'), 6.87 (1H, dd,
H
5.98 (1H, d, J
C
24 typically assignable to a flavanone skeleton. There was a methyl signal (δ
25 group [δ 3.05 (1H, d, J = 16.0 Hz, H-1''a) and 2.25 (1H, d, J = 16.0 Hz, H-1''b)], and a carbonyl carbon
26 (δ 214.2, C-2''), accounting for a CH
H
2.23, s, H
3
-3''), a methylene
H
2
–CO–CH
3
fragment. In addition, a singlet δ 3.12 (3H, s, 2-OCH3)
H
C
1
13
2
2
2
2
2
2
2
2
2
2
2
2
2
2
2
27 for a methoxy group was detected in the H NMR spectrum of 5. The C NMR spectrum of 5 presented
28 19 signals, comprising two carbonyl [δ 214.2 (C-2'') and 196.4 (C-4)], a ketal (δ 108.8, C-2), an
C
C
29 oxygenated (δ 79.9, C-3), a methoxy (δ 51.4, 2-OCH
3
), a methyl (δ 31.9, C-3''), a methylene (δ 41.3,
C
C
C
C
2
8
30 C-1''), and 12 quaternary carbons. A comparison of the NMR data of 5 to erigeroflavanone revealed
31 their close structural similarities, except for the methoxy group (2''-OCH ) in erigeroflavanone was
32 replaced by a methyl group (2''-CH ) in 5. This inference was confirmed by the downfield chemical shift
3
3
33 of C-2'' (δ 214.2) in 5 in comparison with C-2'' (δ 171.4) in erigeroflavanone, and the HMBC
C
C
34 correlations from the methyl protons H
3
-3'' (δ 2.23) to C-2'' (δ 214.2) and C-1'' (δ 41.3), as well as
H
C
C
35 from H-1''a (δ 3.05) and H-1''b (δ 2.25) to C-2'' (δ 214.2) (Figure 2). In addition, the positions of the
H
H
C
36 methoxy group at C-2 and the acetonyl group at C-3 were confirmed by the HMBC correlations from H-
37 2' (δ 7.01), H-6' (δ 6.87), and 2-OCH (δ 3.12) to C-2 (δ 108.8), as well as the correlations between
H
H
3
H
C
38 H-1''a (δ 3.05) and H-1''b (δ 2.25) and C-4 (δ 196.4), C-2 (δ 108.8), and C-3 (δ 79.9), respectively
H
H
C
C
C
39 (Figure 2). Based on the above analysis, the planar structure of 5 was deduced.
40 Cepaflava B (6) was obtained as a pale yellow, amorphous powder, which has the same molecular
41 formula of C H O as 5 based on its molecular ion peak at m/z 390.0954 in the HREIMS spectrum. The
1
9
18
9
1
13
2
2
42
H and C NMR data of 6 were very similar to those of 5 (Table 1), except for the differences in chemical
43 shift of H
2
-1'' [δ 2.42 and 2.56 (6) and 2.25 and 3.05 (5)], C-4 [δ 199.0 (6), 196.4 (5)], C-1'' [δ 49.8
H
C
C
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44 (6), 41.3 (5)], and C-2'' [δ 209.3 (6), 214.2 (5)], suggesting that 5 and 6 may be stereoisomers. The
C
1
3
2
2
2
2
2
2
2
2
2
2
2
2
2
2
2
2
2
2
45 relative configuration of 2-OCH and 3-OH groups in 5 and 6 was determined by comparing their C
3
46 NMR chemical shifts with those of stereoisomers: trans-2,5-dihydroxy-6/8-methyl-7-methoxyflavanone
2
9
13
47 (trans-DMM) and cis-DMM, which possess a hydroxyl group at both C-2 and C-3. The C NMR
48 chemical shifts values at C-2, C-3, and C-4 of trans-DMM (δ 103.7/103.4, 75.4/75.2, and 197.5/197.3)
C
49 were smaller than those of cis-DMM (δ 105.4/104.8, 77.4, and 197.8/197.6). The same pattern was
C
50 observed in 5 and 6 [δ 108.8, 79.9, and 196.4 (5) and δ 109.4, 80.0, and 199.0 (6)] which indicated the
C
C
51 relative configuration of 5 and 6 as the trans- and cis-form, respectively. This initial assignment was
52 supported by the calculation of the chemical shift followed by statistical analysis (CP3 probability
1
13
53 method). The H and C NMR chemical shifts of both trans- and cis-forms of 5 and 6 were calculated
54 and compared with the experimental values, and CP3 analysis showed a 100% probability for 5 as trans-
55 and 6 as cis-isomers (Figure S62, Supporting Information). The ECD spectra of 5 and 6 (Figure 4) were
3
0
31
56 similar to those of mattucinol-7-O-β-D-glucopyranoside, naringin, and (2R, 3R)-taxifolin-6-C-β-D-
57 glucopyranoside32 displaying a negative Cotton effect at ~300 nm. Accordingly, the absolute
58 configuration of C-2 in both 5 and 6 was assigned as R. Calculated ECD spectra of 5 and 6 supported
59 their 2R configuration (Figure 4). Therefore, the absolute configurations of 5 and 6 were determined to
60 be (2R, 3R) and (2R, 3S), and given the trivial names cepaflavas A and B, respectively (Figure 1).
61
Cepadials A (7) and B (8) were isolated as brown, amorphous powders. The molecular formulae
62 of 7 and 8 were determined to be the same, C H O , based on the sodium adduct molecular ion peaks
2
2
14 11
1
2
2
2
2
2
63 in their HRESIMS spectra. The H NMR data of 7 revealed a similar pattern to 8, with two sets of ABX
64 coupling systems [δ 7.676 (1H, dd, J = 9.0, 2.0 Hz, H-6''), 7.682 (1H, d, J = 2.0, H-2''), and 7.02 (1H,
H
65 d, J = 9.0 Hz, H-5'')] for D-ring and [δ 7.23 (1H, d, J = 2.3 Hz, H-2'), 7.05 (1H, dd, J = 8.5, 2.3 Hz, H-
H
66 6'), and 6.69 (1H, d, J = 8.5 Hz, H-5')] for B-ring in 7, and [δ 7.70 (1H, dd, J = 8.5, 1.8 Hz, H-6''), 7.61
H
67 (1H, d, J = 1.8, H-2''), and 7.12 (1H, d, J = 8.5 Hz, H-5'')] for D-ring and [δ 7.24 (1H, d, J = 2.2 Hz, H-
H
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68 2'), 7.07 (1H, dd, J = 8.5, 2.2 Hz, H-6'), and 6.71 (1H, d, J = 8.5 Hz, H-5')] for B-ring in 8, and two meta-
69 coupled proton signals [δ 5.93 (1H, d, J = 2.1 Hz, H-8), 5.94 (1H, d, J = 2.1 Hz, H-6)] in 7 and [δ 5.96
H
H
1
3
2
2
2
2
2
2
2
2
2
2
2
2
2
2
2
2
2
2
2
2
2
2
70 (2H, s, H-6 and H-8)] in 8. Analysis of the C NMR data presented total 22 carbon resonances, including
71 an ester carbonyl signal at C-1''' (δ 169.1 in 7 and 8), a ketone carbonyl signal at C-4 (δ 189.6 in 7 and
C
C
72 189.7 in 8), characteristic signals of two quaternary carbons [C-3 (δ 92.1 in 7 and 91.8 in 8) and C-2 (δC
C
73 101.8 in 7 and 101.9 in 8)], and 18 aromatic carbons. These two characteristic quaternary carbon signals
74 (C-2 and C-3) resembled those of the 2,3-dioxygenated flavanone structure in 5 and 6. However, C-3 (δC
75 92.1 in 7 and 91.8 in 8) was deshielded in comparison with those in 5 (δ 79.9) and 6 (δ 80.0), likely
C
C
76 because of the stronger inductive effect caused by the change to a hemiketal carbon of C-3 in 7 and 8.
77 These NMR data of 7 and 8 were almost identical to those of two regioisomers reported previously,33
78 which are supposedly condensed compounds derived from quercetin and protocatechuic acid. However,
79 in that study, the precise planar structures, as well as their stereochemistry of those two regioisomers
80 have not yet determined.33 An ABX coupling system, a carboxyl group, along with the HMBC
81 correlations from H-2'' (δ 7.682 in 7 and 7.61 in 8) and H-6'' (δ 7.676 in 7 and 7.70 in 8) to δ 169.1,
H
H
C
82 H-5'' (δ 7.02 in 7 and 7.12 in 8) to C-3'' (δ 141.7 in 7 and 142.0 in 8) and C-4'' (δ 146.6 in 7 and 146.1
H
C
C
83 in 8) confirmed the existence of the protocatechuic acid moiety in both 7 and 8. Oxidative coupling of
84 the conjugated olefinic linkage (C-2/C-3) of the C-ring of quercetin and the ortho-dihydroxy group (C-
85 3''/C-4'') of protocatechuic acid would give rise to two regioisomers with either a C2-C4''/C3-C3'' or C2-
86 C3''/C3-C4'' dioxane linkage with more favorable cis-form because of spatial arrangement (syn-addition,
87 Figure 3). This cis-favorability was supported by lower calculated molecular energy of cis-isomer
88 (22.8898 kcal/mol and -1674.064271 Hartree) than that of trans-isomer (25.8620 kcal/mol and -
89 1674.051275 Hartree) of compound 7 (Figure S63, Supporting Information), and by referring the
90 previous research, in which only cis-isomer was reported to be produced from quercetin and a 1,2-
3
4
91 benzoquinone derivative under reflux and microwave condition. Therefore, a total of four cis-isomers
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2
2
92 (7a, 7b, ent-7a, and ent-7b) are possible for 7 and 8 (Figure 5A and Figure S55, Supporting Information).
93 Considering that HMBC correlations cannot distinguish between 7a and 7b (or ent-7a and ent-7b), we
1
13
94 calculated the H and C NMR chemical shifts and compared them with the experimental values,
95 revealing that the major differences are at the H-2'' and H-5''. As shown in Figure 5B, the experimental
96 ∆δH-2''/H-5'' were 0.67 for 7 and 0.50 for 8, while the calculated ones were 0.61 for 7a and 0.54 for 7b
97 (Table S3, Supporting Information), suggesting that the most plausible structures for 7 and 8 are
98 structures of 7a and 7b (or ent-7a and ent-7b), respectively. The proposed structures of 7 and 8 were
99 supported by comprehensive statistical analyses using the CP3 probability method on the entire
1
13
3
3
3
3
3
3
00 calculated and experimental H and C NMR chemical shifts, with 98% probability (Table S3, Figure
01 S56, Supporting Information). Despite our efforts to assign the absolute configuration of compounds 7
02 and 8 by simulation of ECD and calculation of optical rotation values, the resultant data were not
03 conclusive. Hence, the structures of 7 and 8 were demonstrated, as shown in Figure 1, and named
04 cepadials A and B, respectively.
05
Cepadial C (9) was isolated as a brown, amorphous powder. The molecular formula of 9 was
+
3
3
06 established as C H O based on the sodium adduct molecular ion peak at m/z 491.0588 [M + Na]
23 16 11
1
13
07 (calcd for C H O Na, 491.0590) by HRESIMS. The H and C NMR data of 9 were similar to those
2
3
16 11
3
3
3
3
3
3
3
08 of 8, except for the presence of resonances of a methoxy group [δ 3.84 (3H, s), δ 52.6] and a shielded
H
C
09 carboxyl signal at C-1''' in 9 (δ 169.1 in 8 and 167.8 in 9). These differences indicated that 9 was a
C
10 methyl ester derivative of 8. The HMBC correlations between the carbon at C-1''' (δ 167.8) and methoxy
C
11 proton δ 3.84 confirmed the location of the methoxy group at the carboxyl group of the protocatechuic
H
12 acid moiety. The relatively small ∆δH-2''/H-5'' value of 9 (0.47 ppm) indicated that the location of the
13 carboxylic acid is the same as that of 8 (Figure 5C). Thus, the structure of 9 (cepadial C) was interpreted
14 as shown in Figure 1.
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Cepabifla A (10) was obtained as a bronze-yellow, amorphous powder and its HRESIMS showed
+
16 a sodium adduct molecular ion peak at m/z 787.1119 [M + Na] (calcd for C H O Na, 787.1122),
36
28 19
1
3
3
3
3
3
3
3
3
17 consistent with the molecular formula C H O . The H NMR spectrum of 10 displayed signals for two
36 28 19
18 sets of trisubstituted benzene rings [δ 6.95 (1H, s, H-2'), 6.86 (1H, d, J = 8.4 Hz, H-6'), 6.73 (1H, d, J =
H
19 8.4 Hz, H-5') for B-ring, and δ 7.95 (s, 1H, H-2'''), 7.85 (1H, d, J = 8.2 Hz, H-6'''), 7.29 (1H, d, J = 8.2
H
20 Hz, H-5''') for B'-ring]; two meta-coupled proton signals at δ 5.90 (1H, s, H-6) and 5.85 (1H, s, H-8) for
H
21 A-ring, an aromatic proton signal at δ 6.54 (1H, s, H-6'') for A'-ring, and an anomeric proton at δ 4.92
H
H
22 (1H, d, J = 8.6 Hz, H-1''''). Acid hydrolysis of 10 by CF
3
COOH yielded D-glucose, which was identified
23 by comparison of TLC migration and specific rotation with those of standard compound. The β-
24 configuration of the D-glucose moiety was deduced from the large coupling constant (J = 8.6 Hz) of the
1
3
3
3
3
3
3
3
3
3
3
3
3
3
3
25 anomeric proton (H-1''''). The C NMR spectrum of 10 presented 30 resonances correlated to the
26 aglycone motif, including three characteristic signals at δ 119.4 (C-2), 81.8 (C-3), and 192.6 (C-4). The
C
27 comparison of these spectroscopic data with those of a known compound Lawsonia biflavone A
3
5
28 suggested that 10 could be a biflavonoid. The structural differences between 10 and Lawsonia biflavone
29 A were the presence of a D-glucose moiety and a trisubstituted benzene ring (B-ring) in 10 instead of a
3
5
30 disubstituted benzene ring in Lawsonia biflavone A. The HMBC correlation from the anomeric proton
31 H-1'''' (δ 4.92) with C-4''' (δ 148.1) confirmed the linkage between the sugar group and the aglycone
H
C
32 (Figure 2). Similar to the more favorable syn-addition between quercetin and protocatechuic acid in the
33 proposed biosynthetic pathway of compounds 7–9 (Figure 3), the coupling of two quercetin molecules
34 could occur through syn-addition, resulting in the cis-form of 10 (Figure S60, Supporting Information).
35 The calculated ECD spectrum of 10 with (2S, 3S)-configuration, showed a good fit with the experimental
36 result, confirming the absolute configuration of 10 (Figure 4). Hence, the structure of 10, a new
37 biflavonoid glycoside (cepabifla A), was characterized, as shown in Figure 1.
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38
Cepabiflas B (11) and C (12) were both obtained as bronze-yellow, amorphous powders. These
39 compounds shared the same molecular formula C H O based on their HRESIMS data (m/z 787.1105
36
28 19
+
1
13
3
3
40 for 11 and 787.1115 for 12 [M + Na] , calcd for C H O Na, 787.1122). The H and C NMR data of
36 28 19
41 11 and 12 were remarkably similar to each other, but their ECD spectra were mirror images (Figure 4),
1
3
3
3
3
3
3
3
3
42 suggesting that they could be a pair of stereoisomers. Inspection of the H NMR spectra of 11 and 12
43 revealed signals derived from two sets of ABX coupling systems [δ 6.97 (1H, d, J = 2.1 Hz, H-2'), 6.86
H
44 (1H, dd, J = 8.5, 2.1 Hz, H-6'), 6.82 (1H, d, J = 8.5 Hz, H-5') for B-ring and δ 7.80 (1H, d, J = 2.2 Hz,
H
45 H-2'''), 6.33 (1H, dd, J = 8.8, 2.2 Hz, H-6'''), 7.02 (1H, d, J = 8.8 Hz, H-5''') for B'-ring in 11, and δ 6.94
H
46 (1H, d, J = 1.8 Hz, H-2'), 6.81 (1H, dd, J = 8.4, 1.8 Hz, H-6'), 6.76 (1H, d, J = 8.4 Hz, H-5') for B-ring
47 and δ 7.78 (1H, s, H-2'''), 6.13 (1H, d, J = 8.8 Hz, H-6'''), 6.99 (1H, d, J = 8.8 Hz, H-5''') for B'-ring in
H
48 12]; two meta-coupled proton signals [δ 5.99 (1H, d, J = 2.1 Hz, H-6) and 6.02 (1H, d, J = 2.1 Hz, H-
H
49 8) for A-ring in 11, and δ 5.93 (1H, s, H-6) and 5.97 (1H, s, H-8) for A-ring in 12], and an aromatic
H
1
3
3
3
3
3
3
3
3
3
3
3
3
3
50 proton signal [δ 6.44 (1H, s, H-6'')] in 11 and δ 6.39 (1H, s, H-6'') in 12]. The C NMR spectra of 11
H
H
51 and 12 showed a carbonyl carbon (δ 186.9, C-4 in 11 and 12), an oxygenated carbon (δ 103.2 in 11
C
C
52 and 104.3 in 12, C-3), and a highly deshielded ketal carbon (δ 93.7 in 11 and 93.6 in 12, C-2). By
C
53 comparing these NMR data with the literature, 11 and 12 were confirmed to be two stereoisomers of 2-
54 [4-(β-D-glucopyranosyloxy)-3-hydroxyphenyl]-3,5,7-trihydroxy-9-[2-(3,4-dihydroxyphenyl)-2,3-
55 dihydro-2,3-epoxy-5,7-dihydroxy-4H-1-benzopyrane-4-on-8-yl]-4 H-1-benzopyrane-4-one.33 In this
56 previous communication, the individual stereoisomers were not purified, and the absolute configurations
57 were not determined. To assign the absolute configurations of the 2,3-epoxide present in 11 and 12, the
58 experimental ECD spectra of 11 and 12 were compared with their simulated ECD spectra. Based on the
59 results shown in Figure 4, the absolute configurations of 11 and 12 were deduced as (2S, 3S) and (2R,
60 3R) forms, respectively. Thus, the structure of compounds 11 and 12 was established and named
61 cepabiflas B and C, respectively (Figure 1).
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Cepadial D (14) was isolated as a bronze-yellow, amorphous powder. The molecular formula was
63 determined to be C H O by its HRESIMS that displayed a sodium adduct molecular ion peak at m/z
3
8
24 18
+
1
3
3
3
3
3
3
64 791.0848 [M + Na] (calcd for C H O Na, 791.0860), implying 27 degrees of unsaturation. The H
38 24 18
65 NMR spectrum of 14 revealed the presence of two sets of meta-coupled aromatic hydrogen resonances
66 at δ 5.93 (2H, d, J = 2.7 Hz, H-6, H-6'') and δ 6.00 (2H, d, J = 2.7 Hz, H-8, H-8''), a methoxy group at
H
H
67 δ 3.88 (3H, s), together with three sets of trisubstituted benzene rings [δ 7.71 (1H, d, J = 8.5 Hz, H-
H
H
68 6''''), 7.63 (1H, s, H-2''''), and 7.18 (1H, d, J = 8.5 Hz, H-5'''')] for D-ring, [δ 7.21 (1H, s, H-2'), 7.03 (1H,
H
69 d, J = 8.4 Hz, H-6'), and 6.69 (1H, d, J = 8.4 Hz, H-5')] for B-ring, and [δ 7.44 (1H, d, J = 2.0 Hz, H-
H
1
3
3
3
3
70 2'''), 7.34 (1H, dd, J = 8.5, 2.0 Hz, H-6'''), and 6.93 (1H, d, J = 8.5 Hz, H-5''')] for B'-ring. The C NMR
71 spectrum of 14 showed 38 carbon signals, including two carbonyl groups (δ 189.7 and 189.3), an ester
C
72 carbonyl group (δ 167.7), a methoxy group (δ 52.6), four characteristic quaternary carbon signals (δ
C
C
C
2
3
3
3
3
3
3
3
3
3
3
3
3
3
73 101.8, 101.5, 91.9, 91.7), 12 oxygenated aromatic carbons, and 18 sp methine carbons. These above
74 NMR data were similar to those of 9, except for the presence of an additional quercetin moiety in 14
75 (Tables 1 and 2, and Figure 5C). The chemical shifts of C-2/C-2'' or C-3/C-3'' in 14 were similar, while
76 the chemical shifts of C-3', C-4' were downfield shifted (~4 ppm) from those of C-3''', C-4''', respectively,
77 suggesting that the additional quercetin unit could be attached at C-3' and C-4'. The chemical shifts of C-
78 2'' (δ 101.5) and C-3'' (δ 91.7) were almost identical to those of C-2 (δ 101.8) and C-3 (δ 91.9),
C
C
C
C
79 respectively, and this similarity implied the additional quercetin unit was connected through the same
80 dioxane linkage as in C-2 and C-3. As described above for 7–9, all the dioxane linkages in 14 were
81 suggested as cis-forms. Based on these data, a total of eight isomers were possible (14a–14d and ent-
82 14a–14d; Figure S57, Supporting Information). To determine the planar structure and relative
83 configuration of 14, chemical shifts of 14a–14d (chemical shifts of enantiomers of each isomer were
84 identical) were calculated and subjected to DP4 analysis with the experimental values (Figure S57,
85 Supporting Information). The statistical results denoted the structural equivalence of 14 to 14b, with a
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3
3
3
3
3
3
3
3
3
3
3
3
3
4
4
4
4
4
4
4
4
4
86 100% probability (Figure S59, Tables S4, and S5, Supporting Information). The absolute configuration
87 of 14 was deduced by comparing its experimental ECD data with the calculated ECD of 14b and its
88 enantiomer. As shown in Figure 4, the experimental ECD of 14 well-matched those of 14b, suggesting
89 that the absolute configuration of 14 was 2R, 3R, 2''S, 3''S. From those data, compound 14 (cepadial D)
90 was determined, as shown in Figure 1.
91
On the basis of analysis of the spectroscopic data and by comparisons with the literature, the
3
6
37
92 structures of six known compounds were determined as quercetin (1), kaempferol (2), isorhamnetin
93 (3),38 quercetin 4'-O-β-D-glucopyranoside (4),36 1,3,11a-trihydroxy-9-(3,5,7-trihydroxy-4H-1-
94 benzopyran-4-on-2-yl)-5a-[4-(β-D-glucopyranosyloxy)-3-hydroxyphenyl]-5,6,11-hexahydro-5,6,11-
3
9
95 trioxanaphthacene-12-one (13), and 1,3,11a-trihydroxy-9-(3,5,7-trihydroxy-4H-1-benzopyran-4-on-2-
96 yl)-5a-[1,3,11a-trihydroxy-5a-(3,4-dihydroxyphenyl)-5,6,11-hexahydro-5,6,11-trioxanaphthacene-12-
3
3
97 on-9-yl]-5,6,11-hexahydro-5,6,11-trioxanaphthacene-12-one (15). Besides, the appearance of attached
98 acetonyl or methoxyl groups in compounds 5, 6, 9, and 14, gave rise to doubt about their artificial origin.
99 Thus, we analyzed the ethanol extract of original sample (onion skins) by LC-MS/MS. As a result, only
00 compound 14 was detected (Figure S64, Supporting Information) in the ethanol extract, so compounds
01 5, 6, and 9 might be artificial products formed during extraction and isolation.
02
Antioxidant Activity. The scavenging capacities of 5–15 were investigated against two free radical
•
+
03 DPPH and ABTS in concentration-dependent manners as shown in Table 4. Ascorbic acid and trolox
•
+
04 were used as reference compounds to validate the inhibition results in DPPH and ABTS scavenging
•
+
05 assays, respectively. All tested compounds presented good DPPH and ABTS inhibition with IC range
5
0
06 of 4.25–47.69 and 7.12–23.45 μM, respectively. Among the tested compounds, 10–12 were found to
07 possess significant scavenging properties. Compound 11 showed the strongest quenching capacities
•
+
08 against both DPPH and ABTS radicals with IC of 4.25 and 7.12 μM, respectively, followed by 10
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09 (IC of 6.27 and 7.84 μM). On comparing 7–9, it could be proposed that the position of the carboxylic
5
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10 group could affect the DPPH scavenging activity.
11 α-Glucosidase and PTP1B Inhibitory Activities. Inhibition of α-glucosidase and PTP1B
12 enzymes is an effective strategy to treat metabolic diseases such as diabetes and obesity. All isolated
13 compounds were assayed in vitro for their α-glucosidase and PTP1B inhibitory activities. The results are
14 presented in Table 4. Compounds 1–3 and 8–15 differed in their inhibition against PTP1B in a dose-
15 dependent manner, with an IC ranging from 1.13 to 62.42 μM. Especially, compounds 15 (IC = 1.13
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16 μM), 14 (IC = 1.68 μM), and 13 (IC = 6.82 μM) displayed strong PTP1B inhibition, more potent than
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17 both of two positive controls [ursolic acid (IC = 12.09 μM) and sodium orthovanadate (IC = 23.20
5
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50
18 μM)]. Compounds 10 and 12 also showed significant PTP1B inhibitory activity, with IC values of 17.01
5
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19 and 14.29 μM, respectively.
20 In addition, two compounds (4 and 6) displayed no activity against α-glucosidase, while the others
21 exhibited α-glucosidase inhibition with an IC range of 0.89–74.44 μM, which was noticeably higher
5
0
22 than that of the positive control acarbose (IC = 257.41 μM). Interestingly, the two most potent PTP1B
5
0
23 inhibitors compounds (14 and 15) exhibited outstanding α-glucosidase inhibition too and were found to
24 be 100- and 250-fold more active than acarbose, respectively. Compound 9, with one more methyl group,
25 was determined to be the esterification product of 8, and showed similar activities to 8 in both PTP1B
26 and α-glucosidase inhibitory assays, suggesting that the addition of a methyl group did not influence
27 these activities. Compound 7, which differed from 8 (IC = 22.55 μM) and 9 (IC = 22.33 μM) by the
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28 position of the carboxylic group, was inactive in the PTP1B inhibitory assay, implying that the position
29 of the carboxylic group played a key role in the PTP1B inhibitory activity in vitro. In contrast, the α-
30 glucosidase activities of compounds 7, 8, and 9 were not influenced by the alteration of the carboxylic
31 position.
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32
Enzyme Kinetics of PTP1B Inhibition. Lineweaver−Burk and Dixon plots were used to
33 determine the enzyme inhibition properties of compounds 13–15. As depicted in Figure 6, compounds
34 14 and 15 were identified as competitive inhibitors because the regression lines for each compound
35 intersected at the y-axis. Moreover, from Dixon plots, the inhibition constant (K ) values for compounds
i
36 14 and 15 were 1.25 and 2.18 μM, respectively (Table 4). In addition, increasing the concentration of
37 compound 13 resulted in a series of lines sharing a common y-axis intercept but with different gradients,
38 indicating that compound 13 was a mixed-typed inhibitor with a K
i
value of 3.72 μM (Table 4).
39 Molecular Docking Simulation of PTP1B Inhibition. AutoDock 4.2 software was employed to
40 dock compounds 13 and 15 into the binding sites of the crystallographic structure of PTP1B to understand
41 the binding modes and interaction of these active compounds. To optimize and validate the docking
42 procedure, we re-docked two native co-ligands, namely compound A (an allosteric inhibitor) and
43 compound C (a catalytic inhibitor), into the allosteric and catalytic sites of PTP1B, respectively. Figure
44 7 shows the binding pattern of the respective flavonoids in the active site cavity of the PTP1B enzyme,
45 and Table S1 (Supporting Information) presents the binding energies, along with the interacting residues.
46 A previous study indicated that the interference with the binding of helix α7 to helices α3–α6 of PTP1B
1
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47 could lead to allosteric inhibition of the enzyme activity. From our docking results, the PTP1B–13
48 inhibitor complex at the allosteric site displayed negative binding energy of –7.92 kcal/mol, with four H-
49 bond interactions and interacting residues of Ala189 (in the α3 helix), Glu276, Gly277, and Lys279 (in
50 the α6 helix) (Figure 7, Table S1, Supporting Information). In addition, 13 and compound A shared the
51 same allosteric residues, Glu276 (in the α6 helix) via hydrogen bond interactions, and Met282 via Van
52 der Waals interactions. Catalytic inhibition by 13 against PTP1B (–6.41 kcal/mol) showed six H-bonds
53 with Gln266, Thr263, Gly183, Arg221, Arg24, and Asp48 residues. Similar to compound C, 13 was also
54 involved in two H-bond interactions with key catalytic residues Arg221 and Asp48. Furthermore, 13
55 displayed hydrophobic interactions with Phe182, Ala27, Gly259, Val49, Ile219, Ala217, Tyr46, Gly220,
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56 Asp265, and Val184, thus consolidation the protein-ligand interaction between 13 and PTP1B. As
57 demonstrated in Figure 7, 15 was bound in the catalytic region of PTP1B with a –8.01 kcal/mol binding
58 energy, indicating a high affinity to PTP1B residues. The predicted binding mode showed that 15 formed
59 five hydrogen bond interactions with Arg221, Asp48, Gln 262, Arg24, and Gly183 residues through the
60 hydroxy groups. Among them, Arg221 is a signature motif of PTP1B, which is essential for catalytic
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61 inhibition. In addition, 15 and compound C shared the same residues, Gln266, Trp179, and Val49 via
62 hydrophobic interactions and Met258 residue via π-sulfur interactions. Thus, from the docking results,
63 13 and 15 were characterized as mixed and competitive inhibitors of PTP1B, respectively.
64
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66 ASSOCIATED CONTENT
67 Supporting Information
68 The Supporting Information is available free of charge on the ACS Publications website at DOI:
69
70 1D, 2D NMR, and HRMS spectra of cepaflavas A, B (5, 6), cepadials A–D (7–9, and 14), and cepabiflas
71 A–C (10–12). Optimized conformers of 7, 8, and 14 with DFT/B3LYP/6-31G in the gas phase. CP3
72 probability analysis results of 5–8 and DP4 of 14. Proposed biosynthesis of aglycone 10–12. Kinetic
73 studies for selected compounds 13 and 15 (PDF).
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74 AUTHOR INFORMATION
75 Corresponding Author
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76 *E-mail: jkim6923@knu.ac.kr. Tel: +82-53-950-8574. Fax: +82-53-950-8557 (J.A. Kim).
77 *E-mail: bsmin@cu.ac.kr. Tel: +82-850-3613. Fax: +82-53-850-3602 (B.S. Min).
478 ORCID
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79 Byung Sun Min: 0000-0002-8501-4289
80 Jeong Ah Kim: 0000-0001-7337-1812
81 Manh Tuan Ha: 0000-0001-7280-0654
82 Chung Sub Kim: 0000-0001-9961-4093
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83 Author Contributions
⊥
84
Ngoc Khanh Vu and Chung Sub Kim contributed equally to this article.
85 Notes
86 The authors declare no competing financial interest.
87 ACKNOWLEDGMENTS
88 This research was supported by the National Research Foundation of Korea (NRF) funded by
89 the Ministry of Education (NRF-2016R1D1A1B03930246) and the Ministry of Science, ICT and Future
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90 Planning (NRF-2015M3A9A5031271), Korea. We are thankful to the Korea Basic Science Institute
91 (KBSI) for mass spectrometric measurements.
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92 REFERENCES
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92 Legend of Figures
93 Figure 1. Chemical structures of compounds 1–15.
94 Figure 2. Key HMBC and COSY correlations for 5, 9, 10, 11, and 14.
95 Figure 3. Proposed biosynthetic pathway of 7 from two precursors, quercetin and protocatechuic acid.
96 cis-form is more favorable than trans-form because of spatial arrangement.
97 Figure 4. Calculated and experimental ECD spectra of compounds 5, 6, 10–12, and 14 in MeOH
98 Figure 5. Determination of the location of carboxylic acid in 7–9 and 14. (A) Structure of compounds
99 7 and 8 and two possible regioisomers 7a and 7b (see Supporting Information for the structure of their
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00 enantiomers). (B) Experimental H NMR spectra of 7 and 8 (left) and calculated H NMR spectra of 7a
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01 and 7b (right) with the δH-2''–H-5'' value. (C) Experimental H NMR spectra of 9 and 14 with the δH-2''–H-5''
02 value (left) and their structures (right).
03 Figure 6. Lineweaver-Burk plots for PTP1B inhibition of 13 (a), 14 (c), and 15 (e). Dixon plots for
04 PTP1B inhibition of 13 (b), 14 (d), and 15 (f).
05 Figure 7. Molecular docking of compounds 13 (A and D) and 15 (C and F) via catalytic inhibition and
06 13 (B and E) via allosteric inhibition of PTP1B. The 2D and 3D figures were generated using
07 Discovery Studio Visualizer and Pymol, respectively.
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