Bioactivity-Guided Isolation of Potential Antidiabetic and

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Bioactivity-Guided Isolation of Potential Antidiabetic and Antihyperlipidemic Compounds from Trigonella stellata Safa M. Shams Eldin,†,‡ Mohamed M. Radwan,†,‡ Amira S. Wanas,†,§ Abdel-Azim M. Habib,‡ Fahima F. Kassem,‡ Hala M. Hammoda,‡ Shabana I. Khan,† Michael L. Klein,⊥,∥ Khaled M. Elokely,⊥,∥,¶ and Mahmoud A. ElSohly*,†,# †

National Center for Natural Products Research, School of Pharmacy, University of Mississippi, University, Mississippi 38677, United States ‡ Department of Pharmacognosy, Faculty of Pharmacy, Alexandria University, Alexandria 21521, Egypt § Department of Pharmacognosy, Faculty of Pharmacy, Minia University, Minia 61519, Egypt ⊥ Department of Chemistry and ∥Institute for Computational Molecular Science, College of Science and Technology, Temple University, Philadelphia, Pennsylvania 19122, United States ¶ Department of Pharmaceutical Chemistry, Faculty of Pharmacy, Tanta University, Tanta 31527, Egypt # Department of Pharmaceutics and Drug Delivery, School of Pharmacy, University of Mississippi, University, Mississippi 38677, United States S Supporting Information *

ABSTRACT: The in vitro antidiabetic and antihyperlipidemic activities of an alcoholic extract of Trigonella stellata were evaluated in terms of the activation of PPARα and PPARγ in human hepatoma (HepG2) cells. The extract was investigated phytochemically, aiming at the isolation of the most active compounds to be used as a platform for drug discovery. Three new isoflavans, (3S,4R)-4,2′,4′-trihydroxy)-7-methoxyisoflavan (1), (3R,4S)-4,2′,4′-trihydroxy-7-methoxy-4′-O-β-D-glucopyranosylisoflavan (2), and (2S,3R,4R)-4,2′,4′-trihydroxy-2,7-dimethoxyisoflavan (3), were isolated and characterized along with the five known compounds p-hydroxybenzoic acid (4), 7,4′-dihydroxyflavone (5), dihydromelilotoside (6), quercetin-3,7-O-α-Ldirhamnoside (7), and soyasaponin I (8). The structures of 1−3 were elucidated using various spectroscopic techniques including HRESIMS and 1D and 2D NMR. The absolute stereochemistry of the new isoflavans (1−3) was determined using both experimental and calculated electronic circular dichroism as well as DP4 calculations. The isolated compounds were tested for their PPARα and PPARγ activation effects in HepG2 cells.

T

agents is growing considerably, and it accounts for over 10% of the total healthcare costs in several developed countries.4 Peroxisome proliferator-activated receptors (PPARs) are a group of nuclear receptor proteins that function as transcription factors regulating the expression of genes.5,6 PPARs are composed of three subtypes, α, β/δ, and γ, which play key roles in glucose and lipid metabolism as well as energy homeostasis.7 Of the three isoforms, PPARα is expressed

ype 2 diabetes mellitus (T2DM) is a metabolic disease characterized by insulin deficiency resulting from inadequate β-cell insulin secretion or insulin resistance. The worldwide prevalence of T2DM is rapidly increasing, and the number of patients is projected to become approximately 550 million by 2030.1 Increased blood glucose levels in T2DM are closely associated with other metabolic disorders such as hypertension, atherosclerosis, and cardiovascular disease.2 Furthermore, chronic hyperglycemia of diabetes leads to long-term organ damage, especially the eyes, kidneys, nervous system, and heart.2,3 Accordingly, the market for oral T2DM © XXXX American Chemical Society and American Society of Pharmacognosy

Received: August 18, 2017

A

DOI: 10.1021/acs.jnatprod.7b00707 J. Nat. Prod. XXXX, XXX, XXX−XXX

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Chart 1

Trigonella stellata Forrsk. (Leguminosae) is a member of the genus Trigonella, which includes fenugreek (T. foenumgraecum), used as a traditional medicine as an antidiabetic and antihyperlipidemic.21 These plants contain diverse chemical constituents such as phenolics and flavonoids, of which isoflavonoids are predominant. Recent studies reported that soy isoflavones are effective antihyperlipidemics with no reported adverse reactions when tested on menopausal women.22 Moreover, a study testing the effect of soy isoflavones on ovariectomized rats emphasized the antioxidant properties and antidiabetic effects of soy isoflavones and suggested the use of these natural phytoestrogens as a strategy for relieving oxidative stress and insulin resistance in postmenopausal women.23 The present investigation was undertaken to evaluate the potential antidiabetic and antihyperlipidemic effects of the crude extracts and fractions of T. stellata, in terms of their ability to activate PPARα and PPARγ in human hepatoma (HepG2) cells. This work resulted in the isolation of compounds 1−8 from the active chromatographic fractions (Table S1, Supporting Information). Their activities in PPARα and PPARγ activation assays are reported.

primarily in the liver, kidney, and heart and enhances fat degradation.8 Activation of PPARα promotes the uptake, utilization, and catabolism of fatty acids by upregulation of genes involved in fatty acid transport and fatty acid βoxidation.9 It is activated primarily through ligand binding. PPARα-selective agonists such as fenofibrate and gemfibrozil are efficacious in improving dyslipidemia by effectively lowering serum triglycerides and raising serum HDL-cholesterol levels.10 However, they do not have sufficient therapeutic efficacy to serve as useful antidiabetic agents.11 The PPARγ isoform is highly expressed in the adipocytes, which can improve insulin resistance and glucose tolerance in target tissues.12,13 PPARγ agonists are used clinically to counteract hyperglycemia.14 The thiazolidinediones (TZDs), such as ciglitazone and rosiglitazone, are representative PPARγ agonists described as potent insulin-sensitizing drugs for the treatment of T2DM.15 However, administration of TZDs has been associated with unexpected side effects such as weight gain, edema, and hepatotoxicity.16,17 Natural products are a rich source of ligands for nuclear receptors and have been used in treating and preventing diabetes for a long time.18,19 Thus, there has been a continuous interest to find new PPAR modulators from natural products that may not have the property of adipogenesis.20 B

DOI: 10.1021/acs.jnatprod.7b00707 J. Nat. Prod. XXXX, XXX, XXX−XXX

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Table 1. NMR Spectroscopic Data of Compound 1 (500 and 125 MHz, CDCl3, δ in ppm), Compound 2 (400 and 100 MHz, DMSO-d6, δ in ppm), and Compound 3 (600 and 150 MHz, DMSO-d6, δ in ppm) 1

2

3

δH (J in Hz)

δC

δH (J in Hz)

δC

δH (J in Hz)

δC

66.6

5.07 (d, 1H, J = 4.2)

101.0

39.6

3.68 (d, 1H, J = 6.4, Hα-2) 4.29 (bd, 1H, J = 6.0, Hβ-2) 3.67 (d, 1H)

65.9

3

3.61 (dd, 1H, J = 11.5, 11.0, Hα-2) 4.22 (dd, 1H, J = 11.0, 5.0, Hβ-2) 3.51 (ddd, 1H, J = 11.5, 7.0, 5.0)

38.8

44.6

4 5 6

5.49 (d, 1H, J = 7.0) 7.12 (d, 1H, J = 8.5) 6.45

78.9 124.9 106.4

5.61 (d, 1H, J = 6.0) 7.25 (d, 1H, J = 8.0) 6.45 (dd, 1H, J = 2.4, 8.0)

77.7 125.3 106.1

3.71 (dd, 1H, J = 8.4, 4.2) 5.70 (d, 1H, J = 8.4) 7.21 (d, 1H, J = 8.4) 6.45 (dd, 1H, J = 2.4, 8.4)

position 2

7 8 9 10 1′ 2′ 3′ 4′ 5′ 6′ glucose 1″ 2″ 3″ 4″ 5″ 6″ 2-OCH3 7-OCH3

■

6.57 (dd, 1H, J = 2.5, 8.5)

161.2 97.0 156.7 112.0 119.4 158.1 103.8 160.8 110.2

7.35 (d, 1H, J = 8.5)

132.1

6.43

6.43 (d, 1H, J = 2.5)

55.6

6.72 (dd, 1H, J = 2.4, 8.4)

161.0 96.3 160.0 114.1 119.2 156.5 104.0 158.0 110.4

7.39 (d, 1H, J = 8.4)

131.9

4.84 (d, 1H, J = 7.6) 3.23 3.19 3.16 3.31 3.43, 3.67 (2H)

100.3 73.3 76.4 69.7 77.0 60.6

6.43 (d, 1H, J = 2.4)

6.56 (d, 1H, J = 2.4)

3.70 (s, 3H)

6.37 (d, 1H, J = 2.4)

6.26 (d, 1H, J = 2.4) 6.48 (dd, 1H, J = 2.4, 8.4) 7.24 (d, 1H, J = 8.4)

3.40 (s, 3H) 3.76 (s, 3H)

78.6 126.1 106.7 161.1 96.6 153.1 118.6 113.0 159.5 104.1 160.7 110.5 131.7

56.3 55.2

7.35 (1H, d, J = 8.5 Hz, H-6′)]. The 13C NMR and DEPT spectra (Table 1) showed 16 signals, namely, for one methyl, one methylene, eight methines, and six quaternary carbons. In the 13C NMR spectrum, four oxygenated quaternary aromatic carbons were assigned to C-7 (δC 161.2), C-9 (δC 156.7), C-2′ (δC 158.1), and C-4′ (δC 160.8) on a biogenetic basis.25 The chemical shift of the methoxy group (δC 55.6) indicated it is not ortho-flanked.26 The downfield shift of C-7 suggested the attachment of the methoxy group to position C-7. The location of the two hydroxy groups at C-2′ and C-4′ and the position of the methoxy group at C-7 were confirmed by analysis of the HMBC spectrum (Figure 3).

RESULTS AND DISCUSSION Three new 4-hydroxyisoflavans (1−3) along with five known compounds (4−8) were isolated, and their structures determined through 1H and 13C NMR and 2D spectroscopic and HR-ESIMS analysis. Compound 1 was isolated as an optically active ([α]25D −132 (c 0.1, MeOH)) buff powder with a molecular formula of C16H16O5, which was inferred from the HRESIMS [m/z 271.0972 (M − H2O + H)+ in the positive-ion mode and at m/ z 269.0811 (M − H2O − H)− in the negative-ion mode]. The UV spectrum showed only one maximum at 285 nm, characteristic for an isoflavan skeleton.24 This inference was confirmed from the 1H NMR spectrum (Table 1), which exhibited a complex ABMX system for the four aliphatic protons of a hydroxylated isoflavan heterocyclic ring. The occurrence of this system was supported by the presence of a doublet of doublets at δH 3.61 (1H, dd, J = 11.5, 11.0 Hz), a doublet of doublets at δH 4.22 (1H, dd, J = 11.0, 5.0 Hz), a doublet of doublet of doublets at δH 3.51 (1H, ddd, J = 11.5, 7.0, 5.0 Hz), and a doublet at δH 5.49 (1H, d, J = 7.0 Hz); these signals were thus assigned for H-2α, H-2β, H-3, and H-4, respectively, of a 4-hydroxyisoflavan skeleton. The corresponding carbons were identified from the HMQC spectrum as two methines and one methylene carbon at δC 66.6, 78.9, and 39.6, respectively. In the 1H NMR spectrum, several signals were also observed, showing the presence of a methoxy group (singlet at δH 3.76), an ABX system of the a C-7-substituted ring A [δH 7.21 (1H, d, J = 8.5 Hz, H-5), 6.44 (2H, H-6 and H-8)], and another ABX system for a 2′,4′-disubstituted ring B [δH 6.43 (1H, d, J = 2.5 Hz, H-3′), 6.57 (1H, dd, J = 2.5, 8.5 Hz, H-5′),

Figure 1. Electronic circular dichroism (ECD) spectra of compound 1 recorded in methanol in the 350−200 nm range. Calculated ECD spectra as shown in red compared to the experimental values shown in black. C

DOI: 10.1021/acs.jnatprod.7b00707 J. Nat. Prod. XXXX, XXX, XXX−XXX

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Figure 2. Key HMBC correlations of compounds 1−3.

455.1331 [M − H2O + Na]+ in the positive-ion mode and at m/z 467.1120 [M − H2O + Cl]− in the negative-ion mode. The UV spectrum showed a maximum at 285 nm. On comparing its 1 H NMR and 13C NMR spectra with those of compound 1, compound 2 was inferred as being a trihydroxy-7-methoxyisoflavan derivative. However, in contrast to compound 1, the 1H NMR spectrum displayed signals of a hexose sugar (Table 1). The presence of a sugar moiety was shown by the appearance of an anomeric proton doublet at δH 4.84 (J = 7.6 Hz), which correlated to its anomeric carbon signal at δC 100.4 in the HMQC spectrum. Complete assignments of the 1H and 13C NMR signals of the sugar portion were accomplished by HMQC and HMBC experiments, which allowed the unambiguous assignments of all proton signals of the sugar and to identify a glucopyranosyl unit.29,30 The identity was confirmed by acid hydrolysis and TLC on comparing with authentic glucose. The glycosylation site at C-4′ of the aglycone of compound 2 was determined by a HMBC experiment (Figure 2), which showed a long-range correlation between the anomeric proton signal at δH 4.84 (H-1′′Glc) and the carbon resonance at δC 158.0 (C-4′) (Table 1) in addition to the presence of a correlation between δH 4.84 (H-1′′Glc) and H-3′ and H-5′. The absolute stereochemistry of C-3 and C-4 was determined using an ECD experiment. Compound 2 showed a positive Cotton effect at ∼240 nm and a negative Cotton effect at ∼290 nm, in an opposite manner to compound 1, suggesting that it has a (3R,4S) absolute configuration.27,28 Compound 2 exhibited well-matched calculated and experimental ECD spectra (Figure 3) and was assigned as the (3R,4S) diastereomer based on the ECD matching and the DP4 probability. The calculated ECD spectrum of the (3R,4S) diastereomer showed a positive Cotton effect at ∼240 nm similar to the experimental one and a negative Cotton effect at ∼265 nm (about 30 nm apart from the experimental peak). The DP4 probability supported this assignment using 96.3% compared to other diastereomers. On the basis of the above data and extensive 2D NMR experiments (COSY, HMQC, and HMBC), the structure of compound 2 was assigned as (3R,4S)4,2′,4′-trihydroxy-7-methoxyisoflavan-4′-O-β-D-glucopyranoside. Compound 3 was isolated as an optically active ([α]25D −64 (c 0.1, MeOH)) buff white, amorphous powder with a molecular formula of C17H18O6, as inferred from the HRESIMS [m/z 301.1073 (M − H2O + H)+ in the positive-ion mode]. The UV spectrum showed only one maximum at 285 nm, characteristic of an isoflavan skeleton,24 and this was confirmed from the 1H NMR spectrum, which exhibited a complex ABMX system for the three aliphatic protons of a hydroxylated isoflavan heterocyclic ring. This system was indicated by the presence of a doublet at δH 5.07 (1H, d, J = 4.2 Hz), a doublet of doublets at δH 3.71 (1H, dd, J = 8.4, 4.2 Hz), and a doublet at δH 5.70 (1H, d, J = 8.4 Hz); these signals were assigned to H2, H-3, and H-4, respectively, of a 4-hydroxyisoflavan skeleton. The corresponding carbons were identified using the HMQC

Figure 3. ECD (left) spectra of compound 2 recorded in methanol in the 350−200 nm range. Calculated ECD spectra are shown in red compared to the experimental values shown in black.

The absolute configuration of C-3 and C-4 in 1 was established by electronic circular dichroism (ECD). Previous reports have indicated that (3R,4S)-4-hydroxyisoflavan has positive and negative Cotton effects, in turn, at 220−250 nm and 250−300 nm.27,28 However, (3S,4R)-4-hydroxyisoflavan has shown an opposite ECD pattern. The experimental ECD spectrum of compound 1 exhibited a negative Cotton effect at ∼240 nm and a positive one at ∼290 nm, suggesting its absolute configuration as (3S,4R). This assignment was confirmed with computational calculations of the ECD spectra and NMR chemical shifts. The calculated ECD spectrum of the (3S,4R) isomer of compound 1 well matched the experimental one out of the four diastereomers (Figure 1). The calculated ECD spectrum exhibited a strong negative Cotton effect at ∼240 nm and a weak positive value at ∼265 nm. To confirm the assignment mode, DP4 methods were applied using the original DP4 database and the Student’s t-test. The DP4 probability was computed by scaling the calculated chemical shifts with the experimental values. The DP4 probability is the product of dividing the chemical shift error probabilities of each diastereomer by the sum product of the probabilities of all diastereomers. The calculated chemical shifts of the protons and carbons were compared with their closest experimental values. The results of the four diastereomers indicated that (3S,4R) is the most significant diastereoisomer, showing a probability of 97.4% based on both the 13C and 1H NMR chemical shifts. On the other hand, (3R,4S), (3R,4R), and the (3S,4S) diastereomers showed probabilities of 2.5%, 0%, and 0%, respectively. Relying on each 13C or 1H NMR chemical shift, the DP4 probability was still in favor of a (3S,4R) diastereomer with a probability of 72% using the 13C NMR data and 93.8% for the 1H NMR data. On the basis of the above data and extensive 2D NMR experiments (COSY, NOESY, HMQC, and HMBC), the structure of compound 1 was elucidated as (3S,4R)-4,2′,4′-trihydroxy-7-methoxyisoflavan. Compound 2 was isolated as an optically active buff powder ([α]25D −64 (c 0.1, MeOH)), with a molecular formula of C22H26O10, which was determined by HRESIMS at m/z D

DOI: 10.1021/acs.jnatprod.7b00707 J. Nat. Prod. XXXX, XXX, XXX−XXX

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Figure 4. Calculated ECD spectra (left) of four stereoisomers of compound 3 with the experimental ECD spectrum (black); calculated ECD spectrum of the isomer (right) with the experimental values recorded in methanol in the 400−200 nm range.

spectrum as three methine carbons at δC 101.0, 44.6, and 78.6, respectively. In the 1H NMR spectrum, several signals were also observed, showing the presence of two methoxy groups (two singlets at δH 3.40 and 3.68), an ABX system of a C-7substituted ring A [δH 7.21 (d, 1H, J = 8.4 Hz, H-5), 6.45 (dd, 1H, J = 2.4, 8.4 Hz, H-6), 6.37 (d, 1H, J = 2.4 Hz, H-8)], and another ABX system of a 2′,4′-disubstituted ring B [δH 6.26 (d, 1H, J = 2.4 Hz, H-3′), 6.48 (dd, 1H, J = 2.4, 8.4 Hz, H-5′), 7.24 (d, 1H, J = 8.4 Hz, H-6′)]. The 13C NMR and DEPT spectra showed 17 signals: two methyls, nine methines, and six quaternary carbons. In the 13C NMR spectrum, the four oxygenated quaternary aromatic carbons were assigned to C-7 (δC 161.1), C-9 (δC 153.1), C-2′ (δC 159.5), and C-4′ (δC 160.7), on a biogenetic basis.25 The chemical shifts of the two methoxy groups (δC 56.3 and 55.8) indicated they are not ortho-flanked.26 The downfield shift of C-7 suggested the attachment of the methoxy group to position C-7. The location of the two hydroxy groups at C-2′ and C-4′ and the position of the methoxy group at C-7 were confirmed by analysis of the HMBC spectrum (Figure 2). The absolute configurations of C-3 and C-4 were established by the ECD spectrum (Figure 4), showing a negative Cotton effect at 237 nm and a positive Cotton effect at 285 nm, suggesting a β-oriented H-3 and an α-oriented H-4. The ECD spectra were computed for the eight possible isomers of compound 3 and compared with the experimental spectrum. The (2R,3R,4R), (2R,3R,4S), (2R,3S,4R), and (2R,3S,4R) isomers were found to have ECD curves with some similarities to the experimental one (Figure 4). The (2R,3S,4R) showed the best match, but the results were not that conclusive. Thus, the computed and experimental NMR chemical shifts were compared with regression analysis, calculating mean absolute errors and DP4 probability calculation. The results were in favor of a (2R,3S,4R) isomer with an absolute sum of errors of 49.6 (13C NMR) and 1.76 (1H NMR) and mean absolute errors of 3.3 (13C NMR) and 0.195 (1H NMR). The DP4 probability based on both the 13C and 1H NMR spectra supported the assignment of compound 3 as (2R,3S,4R) with 73.4% probability, compared to 26.6% for (2S,3R,4S), 0.1% for (2S,3R,4R), and 0% for the remaining isomers. On the basis of the above data, involving 2D NMR experiments (HMQC and

HMBC) and ECD and computed NMR chemical shifts, the absolute configuration of 3 was proposed as (2R,3S,4R). Thus, the chemical structure of compound 3 was assigned as (2R,3S,4R)-4,2′,4′-trihydroxy-2,7-dimethoxyisoflavan. Generally, the geometry of the 4-hydroxyisoflavan unit is known to exhibit two main conformations based on the C3− C2−O1−C1a dihedral angle (Figure 5). The low-energy

Figure 5. Geometric features of the hydroxyisoflavan 1.

conformer set (1a) has shown a dihedral angle of about −45° (Figure 5, panel A), while the high-energy set (1b) showed a value of about 65° (Figure 5, panel B). This observation stimulated us to run the ECD and DP4 calculations for the two sets of conformers despite the fact that they are more than 10 kJ/mol apart. Of the two geometrical isomers, compounds 1−3 were expected to exist in the 1a conformer (data set 1, Supporting Information) based on the matching between the calculated and experimental ECD spectra. The ECD and NMR calculations supported the assignment of the absolute configuration of compounds 1 and 2. In addition, p-hydroxybenzoic acid (4)31 and 7,4′-dihydroxyflavone (5)32,33 were isolated from the EtOAc fraction and were identified by comparison with published data. Fractionation of the n-BuOH fraction resulted in the isolation of dihydromelilotoside (6),34,35 quercetin 3,7-O-α-LE

DOI: 10.1021/acs.jnatprod.7b00707 J. Nat. Prod. XXXX, XXX, XXX−XXX

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dirhamnoside (7),36,37 and 3-O-[α-L-rhamnopyranosyl-(1‴→ 2″)-β-D-galactopyranosyl-(1″→2′)-β-D-glucuronopyranosyl]soyasapogenol B (soyasaponin I) (8).38 This represents the first reports of the isolation of compound 4 and compound 6 from the genus Trigonella and of compound 7 from the family Fabaceae, while compound 8 was isolated for the first time from T. stellata. The in vitro antidiabetic and antihyperlipidemic activity of the extract, the different fractions, and the eight isolated compounds was tested in terms of PPARα and PPARγ activation through a reporter gene assay. The alcoholic extract did not show any significant activation of PPARα and PPARγ (1.3-fold at 50 μg/mL). The ethyl acetate fraction showed activation of both PPARα (1.5-fold at 50 μg/mL) and PPARγ (1.8-fold at 50 μg/mL) receptors (Table S1, Supporting Information). Among the isolated compounds, 1−3 showed an increase of PPARα activity, while only compound 2 showed an ability to activate the PPARγ receptor as well (Table 2). In addition, compounds 4 and 8 showed mild to moderate activation of PPARγ receptors (Table 2).

compounds was confirmed by HPLC (Waters, PDA detector, C18(2) analytical Luna Phenomenex columns, 150 mm × 4.6 mm, 5 μm). Plant Material. The aerial parts and roots of Trigonella stellata were collected from Daba, Matrouh, Egypt, in May 2012. The plant was identified by Prof. Dr. Salama El-Darier, Faculty of Science, Botany and Microbiology Department, Alexandria University, Alexandria, Egypt. A voucher specimen (No. AU/03/TS 0501) has been deposited in the herbarium of the Department of Botany and Microbiology, Faculty of Science, Alexandria University. ACS-grade solvents, methanol (MeOH), methylene chloride (CH2Cl2), petroleum ether, ethyl acetate (EtOAc), n-butanol (n-BuOH), and hexanes were purchased from Fisher Scientific (Chicago, IL, USA). Extraction and Isolation. The dried plant material (8 kg) was ground and extracted exhaustively with 90% alcohol at room temperature for 10 days. The alcoholic extract was filtered and evaporated under reduced pressure to yield 160 g of dried extract. The alcoholic extract was fractionated with petroleum ether, CH2Cl2, EtOAc, and n-BuOH, successively. The EtOAc-soluble extract was evaporated in vacuo to yield a dried residue (16 g), of which 10 g was chromatographed over a silica gel column (600 g, 60 × 5 cm). Elution was performed initially using CH2Cl2; then the polarity was increased by gradual addition of MeOH. Thirty fractions, each of 500 mL, were collected and monitored chromatographically, using CH2Cl2−MeOH (9:1) as solvents, with similar fractions combined to yield five subfractions (E1−E5). Subfraction E2 (550 mg, eluted with 5−7% MeOH−CH2Cl2) was chromatographed over a silica gel column (35 g, 27 × 2 cm), eluted initially with 40% EtOAc−hexanes; then the polarity was raised gradually by increasing the amounts of EtOAc. Fifty fractions, 25 mL each, were collected and monitored chromatographically using a mixture of EtOAc−hexanes (80:20). Similar fractions were combined to obtain 13 subfractions (E2A−E2M). Subfraction E2C (40% EtOAc−hexanes, 30.5 mg) was crystallized from MeOH to yield 7.8 mg of an amorphous, white residue designated as compound 4. Fraction E2H (70% EtOAc−hexanes, 58.6 mg) was applied on a C18-SPE column (10 g) eluted with 50% MeOH−H2O, where 15 fractions were collected (20 mL each). Fraction 6 showed a yellow spot (7.4 mg) of a white residue, designated as compound 5. Fraction 8 showed a single purplish-pink spot (10.5 mg) of a white, amorphous powder designated as compound 1. Fraction 9 showed a pink spot (13.2 mg) of buff powder designated as compound 2. Subfraction E4 (12−20% MeOH− CH2Cl2) showed one major spot with Rf 0.41 in the solvent system EtOAc−MeOH−H2O−HOAc (30:5:4:0.5). An aliquot of the E4 residue (2.8 g) was subjected to size-exclusion chromatography using a Sephadex LH-20 column (37 cm × 3 cm). The column was eluted with 100% methanol. Thirty-five fractions, 3 mL each, were collected and monitored chromatographically using a mixture of EtOAc− MeOH−H2O−HOAc (30:5:4:0.5) as solvent. Fractions (E4(20)− E4(29)) afforded 20 mg of a buff powder, designated as compound 3. The n-BuOH-soluble fraction was evaporated in vacuo to yield a dried residue (35 g), of which 10 g was chromatographed over a silica gel column (300 g, 120 × 4 cm), eluted with CH2Cl2, with the polarity increased by gradual addition of MeOH. Altogether, 75 fractions, each 100 mL, were collected and monitored chromatographically, using EtOAc−MeOH−H2O−HOAc (30:5:4:0.5), and similar fractions were combined and concentrated in vacuo to yield five subfractions (B1− B5). Fraction B3 (150 mg, 32−45% MeOH−CH2Cl2) showed one major blue-green spot, which was rechromatographed over a Sephadex LH-20 column (30 × 2 cm). The column was eluted with a 50% MeOH−H2O mixture, with the percentage of MeOH increased gradually to 100% MeOH. Thirty-five fractions, 2 mL each, were collected and screened by TLC, and similar fractions were combined to give six subfractions (S1−S6). Subfraction S3 (50% MeOH−H2O) afforded 22 mg of yellow residue. This residue was subjected to preparative TLC on fluorescent silica gel plates, using as developing system EtOAc−MeOH−H2O−HOAc (30:6:4:0.5), to yield 9 mg of a yellow, amorphous powder, designated as compound 6. Fraction S4 (150 mg, 75% MeOH−H2O) showed one major yellow spot that gave a bright yellow color upon exposure to ammonia in addition to minor undifferentiated spots. The residue left after the

Table 2. Activation of PPARα and PPARγ by Compounds Isolated from Trigonella stellataa fold induction in PPARα activity

fold induction in PPARγ activity

compound

50 μg/ mL

25 μg/ mL

12.5 μg/ mL

50 μg/ mL

25 μg/ mL

12.5 μg/ mL

1 2 3 4 5 6 7 8

2.25 1.67 NA NA NA NA NA NA

1.88 1.32 2.00 NA NA NA NA NA

1.29 1.14 1.93 NA NA NA NA NA

NAb 1.74 NA 1.35 NA NA NA 1.79

NA 1.36 NA 0.90 NA NA NA 1.36

NA 1.22 NA 1.00 NA NA NA 1.22

Ciprofibrate (1.45 μg/mL) caused a 2.2-fold induction in PPARα activity, and rosiglitazone (1.79 μg/mL) caused a 3.5-fold induction in PPARγ activity. bNA = no activation. a

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EXPERIMENTAL SECTION

General Experimental Procedures. Optical rotations were obtained using a Research Analytical Autopol V polarimeter (Rudolf). UV spectra were obtained using a Varian Cary 50 Bio UV−visible spectrophotometer, and IR spectra were recorded using a Bruker Tensor 27 spectrophotometer. ECD spectra were obtained using a Cary-50 Bio spectrophotometer and a CD JASCO J-715 spectrometer. 1D and 2D NMR spectra were obtained on a Varian-Mercury AS 400 MHz, Bruker Avance III 500 (with a Prodigy cryogenic probe and z gradients), and Bruker Avance III 600 NMR spectrometer. Highresolution electrospray ionization mass spectrometry (HRESIMS) was conducted on a Bruker Bio Apex spectrometer in the ESI mode. Analytical TLC analysis was performed on precoated silica gel aluminum sheets 60 F254, 0.25 mm (20 × 20 cm), and reversed-phase RP-18 F254 (E. Merck, Germany). Detection was carried out under UV light (254 nm, 366 nm), and visualization effected with vanillin− H2SO4 reagent (1.0 g vanillin in 100 mL of 10% H2SO4 in EtOH) followed by heating. Column chromatographic separations were performed on silica gel G60 (60−120 mesh, Merck, and 63−200 μm) and Sephadex LH-20 (Mitsubishi Kagaku, Tokyo, Japan). Centrifugal preparative TLC (CPTLC, using a Chromatotron; Harrison Research, Inc. model 8924) was carried out using 4 mm silica gel P254 (Analtech) rotors. Solid-phase extraction (SPE) cartridges (Strata C18, 10 g) were employed. Purity of the isolated F

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removal of the solvent was purified using a Sephadex LH-20 column (30 × 2 cm). The column was eluted isocratically with 50% MeOH− H2O to give 12.3 mg of a yellow powder, designated as compound 7. Subfraction B4 (500 mg, 50−55% MeOH−CH2Cl2) showed one major spot. Crystallization from MeOH yielded 15 mg of crystalline, white residue, designated as compound 8. (3S,4R)-4,2′,4′-Trihydroxy-7-methoxyisoflavan (1): buff powder; [α]25D −132 (c 0.1, MeOH); ECD (c 3.47 × 10−4 M, MeOH) λext (Δε) 239 (−58), 291 (+20) nm; UV (MeOH) λmax (log ε) 205 (4.26), 230 (4.24), 285 (4.0) nm; IR νmax 3386, 2919, 1623, 1584 cm−1; 1H NMR (CDCl3, 500 MHz); 13C NMR (CDCl3, 125 MHz) data, see Table 1; HRESIMS m/z 269.0811 [M − H2O − H]− in the negative mode (calcd for C16H13O4, 269.0814) and m/z 271.0972 [M − H2O + H]+ in the positive mode (calcd for C16H15O4, 271.0970). (3R,4S)-4,2′,4′-Trihydroxy-7-methoxyisoflavan-4′-O-β-D-glucopyranoside (2): buff powder; [α]25D −64 (c 0.1, MeOH); ECD (c 2.22 × 10−4 M, MeOH) λext (Δε) 235 (+60), 288 (−22) nm; UV (MeOH) λmax (log ε) 205 (4.62), 215 (4.59), 285 (4.15) nm; IR νmax 3386, 2919, 1623, 1584 cm−1; 1H NMR (DMSO-d6, 400 MHz) and 13C NMR (DMSO-d6, 100 MHz) NMR data, see Table 1; HRESIMS m/z 467.1120 [M − H2O + Cl]− in the negative mode (calcd for C22H24O9Cl, 467.1109) and m/z 455.1331 [M − H2O + Na]+ in the positive mode (calcd for C22H24O9Na, 455.1318). (2S,3R,4R)-4,2′,4′-Trihydroxy-2,7-dimethoxyisoflavan (3): white, amorphous powder; [α]25D −64 (c 0.1, MeOH); ECD (c 2.22 × 10−4 M, MeOH) λext (Δε) 235 (−48), 285 (+18) nm; UV (MeOH) λmax (log ε) 205 (4.20), 215 (4.50), 285 (4.10) nm; IR νmax 3386, 2919, 1623, 1584 cm−1; 1H NMR (DMSO-d6, 600 MHz) and 13C NMR (DMSO-d6, 150 MHz) NMR data, see Table 1; HRESIMS m/z 301.1073 [M − H2O + H]+ in the positive mode (calcd 301.1076 for C17H17O5). Hydrolysis of Compound 2. Compound 2 (3 mg) was hydrolyzed with 2 M aqueous CF3COOH (5 mL) in a thermostatically controlled H2O bath at 80 °C for 3 h. After the solvent was removed in vacuo with MeOH, the residue was partitioned between EtOAc and H2O to give the aglycone of 2. The sugar identity was confirmed by using the aqueous layer obtained from the acid hydrolysis for TLC comparison with authentic sugars (D-glucose, D-galactose, and Lrhamnose) using n-BuOH−HOAc−H2O (5:1:1) as a mobile phase and vanillin sulfuric acid spray reagent. Reporter Gene Assay for the Activation of PPARα and PPARγ. Activation of PPARα and PPARγ was determined in human hepatoma (HepG2) cells as described previously.39 In brief, HepG2 cells were cultured in Dulbecco’s modified Eagle’s medium supplemented with 10% fetal bovine serum, 100 units/mL penicillin G sodium, and 100 μg/mL streptomycin. For the PPARα and PPARγ activation assay, HepG2 cells were transfected with pSG5-PPARα and PPRE X3-tk-luc or pCMV-rPPARγ and pPPREaP2-tk-luc plasmid DNA (25 μg/1.5 mL cell suspension), respectively, by electroporation at 160 V for a single 70 ms pulse using a BTX Electro Square Porator T 820. Transfected cells were plated at a density of 5 × 104 cells/well in 96-well tissue culture plates and grown for 24 h. After 24 h, the cells were treated with the extracts, fractions, or compounds. After incubation for 24 h, the cells were lysed and the luciferase activity was measured. The fold induction of luciferase activity in treated cells was calculated in comparison to the vehicle control. Ciprofibrate and rosiglitazone (both from Cayman Chemical, Ann Arbor, MI, USA) were used as positive controls. Molecular Modeling Study. The relative configurations at C-2, C-3, and C-4 of compound 3 and at C-3 and C-4 of compounds 1 and 2 were assigned by computational calculations of ECD and NMR chemical shifts. MM Conformational Sampling. The conformational space of each diastereoisomer was sampled using MacroModel of the Schrödinger suite.40 A mixed torsional/low-mode sampling search method was performed in the gas phase with intermediate torsion sampling, a 40 kJ/mol energy window, a 1000 maximum number of steps, and 100 steps per rotatable bond. Energy minimization was established using Polak-Ribiere conjugate gradient method, with an OPLS2005 force field with extended cutoff, 5000 maximum iterations,

and a convergence threshold of 0.005. No constraints were imposed during the course of sampling. ECD Calculations. The conformations within 10 kJ of the global minimum structure were geometry optimized at the level of DFT B3LYP/6-31G** with the Gaussian 09 package.41 The conformers were calculated in a polarizable continuum DMSO solvent model. All optimized conformers were then used to calculate the ECD spectra using the time-dependent density functional theory method. The calculations were carried out at the B3LYP/6-31G** level with excited states of 80 and a PCM methanol solvent model. NMR Calculations. The gauge-including atomic orbital (GIAO) shielding constants of all conformers were calculated after geometry optimization using the B3LYP/6-311+G(2d,p) level in a chloroform solvent model with Gaussian 09.41 The Boltzmann-weighted conformer population was calculated based on the Gibbs free energy from the geometry optimization step. The chemical shifts were calculated for each conformer. Then, Boltzmann-weighted averages of the chemical shifts were calculated to scale them against the experimental values. The DP4 method42,43 (http://www-jmg.ch.cam.ac.uk/tools/ nmr/DP4/) was applied to compute the chemical shift errors. The error probability was determined using the Student’s t-test and the original DP4 database (a database of over 1700 13C and1H NMR chemical shifts). For compound 3, the geometry optimization and frequency calculations were carried out at the M06-2X/6-31+G(d,p) level, and the NMR shielding tensors were calculated at the B3LYP/6311+G(2d,p) level using the integrated equation formalism polarized continuum model.

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ASSOCIATED CONTENT

S Supporting Information *

The Supporting Information is available free of charge on the ACS Publications website at DOI: 10.1021/acs.jnatprod.7b00707. 1 H and 13C NMR spectroscopic data for 1−3 as well as the DEPT-135 NMR spectra for 1 and 3, the ROESY spectrum for 2, and the HMQC NMR spectra for 1 and 3 (PDF)

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AUTHOR INFORMATION

Corresponding Author

*Tel: +1 662 915 5928. Fax: +1 662 915 5587. E-mail: [email protected]. ORCID

Mahmoud A. ElSohly: 0000-0002-0019-2001 Notes

The authors declare no competing financial interest.

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ACKNOWLEDGMENTS This study was supported in part by the grant provided by the Egyptian government through the joint supervision program and the National Science Foundation, Alexandria, VA (CNS09-58854).

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REFERENCES

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