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. 2024 Mar 11;9(11):13284–13297. doi: 10.1021/acsomega.3c10083

Molecular Structure, Antioxidant Potential, and Pharmacokinetic Properties of Plant Flavonoid Blumeatin and Investigating Its Inhibition Mechanism on Xanthine Oxidase for Hyperuricemia by Molecular Modeling

Cisem Altunayar-Unsalan †,‡, Ozan Unsalan §,*
PMCID: PMC10956095  PMID: 38524493

Abstract

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Hyperuricemia, which usually results in metabolic syndrome symptoms, is increasing rapidly all over the world and becoming a global public health issue. Xanthine oxidase (XO) is regarded as a key drug target for the treatment of this disease. Therefore, finding natural, nontoxic, and highly active XO inhibitors is quite important. To get insights into inhibitory potential toward XO and determine antioxidant action mechanism depending on the molecular structure, plant flavonoid blumeatin was investigated for the first time by Fourier transform infrared (FTIR) spectroscopy, density functional theory (DFT), ADME/Tox (absorption, distribution, metabolism, excretion, and toxicity) analysis, and molecular docking study. Theoretical findings indicated that blumeatin has high radical scavenging activity due to its noncoplanarity and over twisted torsion angle (−94.64°) with respect to its flavanone skeleton could explain that there might be a correlation between antioxidant activity and planarity of blumeatin. Based on the ADME/Tox analysis, it is determined that blumeatin has a high absorption profile in the human intestine (81.93%), and this plant flavonoid is not carcinogenic or mutagenic. A molecular docking study showed that Thr1010, Val1011, Phe914, and Ala1078 are the main amino acid residues participating in XO’s interaction with blumeatin via hydrogen bonds.

1. Introduction

Hyperuricemia (HUA) is the fourth most prevalent metabolic disease following hyperglycemia, hyperlipidemia, and hypertension.1 It is described as a serum urate level that surpasses the usual range of 2.5–7.0 mg/dL in men and 1.5–6.0 mg/dL in women.2 HUA arises from hepatic overproduction and/or renal underexcretion of uric acid (UA), which leads to UA deposition at the kidneys and joints, appearing mainly as kidney stone-related diseases and gout, respectively.3 HUA is not only the molecular base for gout, but it is also linked to the onset and progression of diabetes, hypertension, coronary heart disease, and other illnesses.4

One of the most significant HUA treatment approaches is to successfully inhibit UA production. Xanthine oxidase (XO) is an essential target for the therapy of HUA.1 XO is a multifunctional molybdoflavoprotein enzyme and found in many mammalian tissues, particularly the intestine tract and liver.5 It catalyzes the oxidation of hypoxanthine and xanthine to generate UA and reactive oxygen species (ROS) such as superoxide anion radical and hydrogen peroxide, which plays an essential role in HUA development.6 Allopurinol, an efficient XO inhibitor, is still the primary drug for HUA, but it has a number of adverse effects, including Stevens-Johnson syndrome, hypersensitivity syndrome, renal toxicity, vasculitis, and ROS-induced diseases.7 As a result, the search for new XO inhibitors with fewer adverse effects and greater therapeutic action as alternatives to current drugs for the avoidance and therapy of HUA is critical.5 Naturally derived XO inhibitors have emerged as one of the research areas for drugs owing to the high safety of naturally derived substances.8

Flavonoids are an extensive group of polyphenolic natural products found in a variety of plant-based foods and beverages.9 They possess a numerous range of biological activities such as anticancer,10,11 antiviral,12−15 antifungal,16 antibacterial,17 and antioxidant activities.18−21 Their bioactivity arises from structural characteristics which provide them powerful ability to inhibit enzymes, decrease oxidized chemical entities, scavenge free radicals, and bring metal chelation.22 However, finding both robust experimental and theoretical links/correlations between the structure and activities of flavonoids is still a challenging ongoing process. In our previous studies, we focused on the experimental and theoretical vibrational spectral results of the molecular structures of some flavonoid derivatives.23−32 While some flavonoids show activity with the presence of both a double bond (C2–C3) and a catechol moiety, and additionally C3 hydroxyl, some others lacking the C3–OH group still show activity.33 From this point of view, a highly systematic theoretical and experimental effort is crucial to further explain how such structural parameters affect the antioxidant and radical scavenging activities of these natural plant-based compounds. While the presence and number of hydroxyl groups at certain positions were considered in some previous studies,34,35 the interaction between the groups was not taken into account. Thus, this study targets one step further in the evaluation of these correlations by investigating a very rarely studied flavonoid family member, blumeatin. Blumeatin is a naturally occurring flavonoid found in the sembung plant (Blumea balsamifera L.).36 It has hydroxyl groups at positions C-3′ and 5′ and lacks a C2–C3 double bond (Figure 1). Compared to flavanol tamarixetin, blumeatin has a higher level of antioxidant activity.37 Blumeatin also possesses a number of biological properties, including superoxide radical scavenging, hepatoprotective, antioxidant, and antityrosinase activities.36,38,39 However, no detailed studies have been reported to elucidate the molecular mechanism of action of blumeatin so far. To clarify the relationship between the structure and molecular activity of this compound, first, attenuated total reflection-Fourier transform infrared (ATR-FTIR) spectroscopy and density functional theory (DFT) calculations were carried out. Thus, blumeatin’s quantum chemical descriptors such as chemical potential (μ = −χ), global chemical hardness (η), electrophilicity index (ω), electronegativity (χ), global softness (s), and nucleophilicity (ε) were found. Second, ADME/Tox (absorption, distribution, metabolism, excretion, and toxicity) analysis on this flavonoid was performed for the earlier prediction of pharmacokinetic properties, and it was evaluated in terms of selected molecular descriptors including drug-likeness, tumorigenicity, mutagenicity, and toxicity. Then, molecular docking studies against XO were conducted. The interaction of blumeatin as an inhibitor with XO is critical to understanding drug–protein interactions and possible therapeutic uses. These interactions can also give important knowledge about the mechanism of binding of blumeatin to XO at the molecule level. Finally, it is clear that further understanding the molecular structure and biological activity relationship of such particular flavonoids would help to design potential similar compounds that could exhibit enhanced antioxidant activity and give people suffering from HUA hope.

Figure 1.

Figure 1

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Blumeatin and its corresponding optimized geometric structure.

2. Materials and Methods

2.1. FTIR Spectroscopy

Blumeatin was purchased from Cayman Chemical (Ann Arbor, MI, USA). 1 mg of blumeatin was inserted on the attenuated total reflection (ATR) diamond crystal which is located in the PerkinElmer spectrum two FTIR spectrometer (PerkinElmer Inc., Waltham, MA, USA). This spectrometer has a deuterium triglycine sulfide (DTGS) detector. FTIR spectra for blumeatin were collected in 4000–400 cm–1 range at ambient temperature. An average of 64 scans were taken during the spectral collection process. The spectral resolution of the spectrometer was adjusted to 2 cm–1 to better distinguish the FTIR spectral peaks. Water and carbon dioxide contributions were eliminated from the spectra by subtracting the initial background spectrum. The spectra were evaluated by the spectrometer’s PerkinElmer Spectrum Version 10.5.4. software (PerkinElmer Inc., Waltham, MA, USA), and ATR-FTIR spectra were plotted with OriginPro 2022 b (Academic Version).40

2.2. Computational Details

Quantum chemical computation on blumeatin was performed by Gaussian 0941 using the B3LYP functional that consists of Becke’s gradient exchange correction;42,43 the Lee, Yang, and Parr correlation functional;44 and the Vosko, Wilk, and Nusair45 correlation functional with the 6-311++G(d,p) basis set.46 Computed wavenumbers were scaled by double factors of 0.967 and 0.955 for the wavenumbers below and over 1800 cm–1, respectively. Optimizations were performed by the geometry direct inversion of the invariant subspace method.47 The optimized molecular structure of blumeatin was found to be a minimum energy conformation. Moreover, for the conformational flexibility determination purpose, a scanning procedure was performed over the dihedral angle (C2–C1–C15–C20) between rings C and B. For this purpose, the same level of basis set and correlation functionals mentioned above were used with 12 steps of 30° increments.

2.3. ADME/Tox Analysis

To obtain the ADME/Tox data for blumeatin, the graph-based structural signatures concept was utilized.48 For this purpose, SMILES (simplified molecular-input line-entry system) code (COC1=CC(=C2C(=O)CC(OC2=C1)C3=CC(=CC(=C3)O)O)O) for blumeatin was introduced into pkCSM, and the ADME/Tox results of blumeatin were obtained.

2.4. Molecular Docking

For molecular docking study, blumeatin and XO were prepared according to the procedure that was previously reported by Altunayar-Unsalan et al.31 The X-ray crystallographic structure of bovine XO with the complex of flavonoid quercetin with 2.0 Å resolution was downloaded from the protein data bank (PDB) (https://www.rcsb.org/) (PDB ID: 3NVY). Optimized molecular structure of blumeatin was used for docking this flavonoid into the site where the cocrystallized ligand (quercetin) of XO resided naturally. Afterward, whole XO protein was sterilized by removing other natural ligands and water using PyMol Molecular Visualization software.49 As the next step, both blumeatin and XO were converted to “pdbqt” file format in PyRx 0.850 to get these structures prepared for docking. In order to visualize and make a conclusion which interactions and residues would be involved for blumeatin, BIOVIA Discovery Studio Visualizer v21.1.0.2029851 was used. After these interaction residues were determined for blumeatin, they were selected in the search grid box in PyRx by manually toggling them one by one manually. The active site of the protein containing amino acids interacting with blumeatin was predicted by using this grid box. Then, the docking procedure was started by the Auto Dock Vina algorithm which was embedded in PyRx. Binding affinities were obtained in kcal/mol, and resultant docking poses of blumeatin that bound to the previously removed natural ligand’s (quercetin) vicinity was saved in pdb file format for further visualization. In addition, we also applied a homology modeling study using the human protein sequence (NCBI Accession number: NP_000370) and carrying out the molecular docking study with the human 3D structure for the PDB code 2E1Q. Our findings are presented in the Supporting Information as Figures 1–9 and Table S4.

3. Results and Discussion

3.1. Molecular Structure, Antioxidant Properties, and Quantum Chemical Descriptors

Blumeatin is composed of a flavanone skeleton structure with three OH moieties at three carbon positions (5′, 3′, and 5) and a methoxy group located at the seventh carbon. Blumeatin is composed of three rings (A, C, and B, respectively), and the C1 symmetry group was obtained for this compound. There are six different types of atom distances (CC, OC, CH, OO, OH, and HH) and 597 distances in total for blumeatin (Table S1). Computed optimized molecular structure of blumeatin is presented in Figure 1, and related data are given in Table S2. A and C rings are connected together with a C4–C5 double bond, and the C ring has a single C=O (C3–O11) group attached to the C3 atom. One OH group is attached to the C10 atom in ring A. The B ring has two OH groups which make blumeatin support possible H-bond acceptors and donors via the O21 and O22 atoms. Blumeatin has only one methyl group connected via the O12 atom in ring A. van Acker et al.52 emphasized that the existence of an OH group at 3-carbon position, C2=C3 double bond (C1–C2 single bond for blumeatin’s numbering scheme), and catechol moiety is responsible for the increasing radical scavenging activity of flavonoids. The authors in that study also focused on the torsion angles of ring B and tried to explain the structure–activity relation based on the planarity. They also questioned why this double bond without a hydroxyl group exhibits increasing activity by means of quantum chemical investigation. In this study, the B and C rings of blumeatin are not in perfect planar orientation. The torsion angles are −36.27° (O6–C1–C15–C20) for ring B and 51.28° (C5–O6–C1–C2) for ring C. Geometrically, rings A and C of flavonoids favor to be in coplanar conformation. For blumeatin, the torsion angle (C4–C3–C2–C1) of ring C was computed to be 33.62° as distorted from planarity. However, the torsion angle (C8–C7–C15–C16) of ring B compared to the plane where rings C and A located was computed to be −76.78° (Figure 2A) compared to the plane where rings A and B located. This deviation from coplanarity with regard to rings A and C could be an explanation or responsible for blumeatin’s better antioxidant activity as explained by previous papers,52−55 and it is obvious that further clarification by systematic theoretical and experimental studies data is required. A computational and crystal diffraction study53 is carried out where the most stable conformations were found approximately at 140° for morin and myricetin. Based on the conformation analysis performed on blumeatin, the scanned torsion angle (C2–C1–C15–C20) of ring B was found to be −94.64° as it is given in Figure 2B. This deviation from the coplanarity is much more than previously reported53 dihedral angles that vary between −42.73 (naringenin) and 35.64° ((+)-catechin), and blumeatin’s better antioxidant activity could also be interpreted by this higher deviation with respect to the most flavonoids.53 It was previously discussed that 3-OH portion is important in the antioxidant activity of flavones by its interaction with ring B via a H bond,53 and researchers in that study stated that this moiety fixes the position of ring B as coplanar with the C and A rings by forming such a H bond. Blumeatin has only two CH bonds in this moiety, and construction of such a hydrogen bond does not seem possible. Although blumeatin does not have a 3-OH moiety, it is interesting to note that blumeatin exhibits better antioxidant activity compared to most flavonoids. Moreover, it is not possible to suggest at the moment why blumeatin shows such a high activity even it lacks both 3-OH moiety and C2=C3 double bond which are known as extreme active scavengers together.55

Figure 2.

Figure 2

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Optimized dihedral angle (C8–C7–C15–C16) between rings A and B of blumeatin (A). Scanned dihedral coordinate (C2–C1–C15–C20) of blumeatin (B).

A recent investigation on the antioxidant activities of Alyssum virgatum by Koç et al.56 demonstrated that ionization potential, bond dissociation energies, single electron transfer followed by proton transfer (SET-PT), hydrogen atom transfer (HAT), and proton dissociation enthalpies are indicators of antioxidant reactivities. In that work, hydroxy radicals in ferulic acid have a better antioxidant property than those in cinnamic acid determined in A.. virgatum. As it was mentioned in their work, DFT-extracted parameters were shown to be effective in understanding the antioxidant mechanism of compounds under study.56 These mechanisms were also investigated for five monophenols (thymohydroquinone, thymol, carvacrol, para-cymene-2,3-diol, and dihydroxyphenol) because of their well-known antioxidant characteristics depending on the locations of OH groups in the compound, and these structures were planar.57 To date, there has not been any specific work dealing with the correlation of antioxidant activity with the planarity of the flavonoids.

The most detailed correlation by means of evaluation of hydroxyl positions and bond dissociation energies in both aqueous solution and the gas phase was studied on myricetin. It was shown by Hu-Jun et al.58 that myricetin and its antioxidant activity are due to the high antioxidation activity of the 4′-OH group in myricetin. This was attributed to very stable H-deleted radical species, conjugation phenomena, and delocalization as well as by an internal hydrogen bond, constructed between the adjacent hydroxyl groups and radicalized O·. For (−)catechin, it was reported that intramolecular hydrogen bonds occurred between OH moieties of the ring B and OH groups of 3- and 5-OH groups with the 4-keto group of ring C.52 However, our results revealed no intramolecular hydrogen bonds for blumeatin, and this compound does not have a 3-OH group. Interestingly, it exhibits more radical scavenging activity (RSA) compared to morin, foeniculin, procyanidin, and gallic acid even with the lack of this specific group. Unlikely to the results of van Acker et al.52 that suggested 3-OH moiety takes a dominant role in the antioxidant activity via a hydrogen bond, it is obvious that more intense work is required to explain this contraction, particularly the correlation of the existence of the 3-OH group and intramolecular hydrogen bonds.

A large number of flavonoids were investigated for their theoretical and experimental radical scavenging activities (RSA) (in %) by developed the quantitative structure–activity relationship (QSAR) model and equation.33 They used the correlation between the number of free phenolic OH groups and the indicator variable, I, developed by Lien and co-workers.59 According to the updated equation by Amić et al.33 (given below), indicator I was included that contains the existence (I = 1) of I3′, 4′-diOH or 3-OH or I5-OH or 2,3-double bond, otherwise (I = 0). This equation was applied to blumeatin and found the RSA to be 88.403%. Our finding is an exact match (88.403%) of 11 flavonoids (quercetin, taxifolin, morin, galangin, kaempferol, luteolin 7-glycosyl, rutin, laricytrin quercetin 3,7-diglycosyl, and laricytrin-3′-glycosyl, myricetin) where at least one of those three criteria met. In contrast, we were not able to compare our prediction to the experimental RSA, but this is planned for our future work. Amić and his co-workers33 also reported that the C2–C3 double bond might not be required for high activity, but a 3-hydroxyl group highly improves the antioxidant activity. Blumeatin lacks both portions but has 3′-5′-diOH groups instead. This contributes to its higher predicted RSA and is in agreement with previous findings.

3.1.

van Acker et al.52 suggested that flavonoids (diosmin, kaempferol, galangin, and apigenin) with a 3-OH moiety are planar, whereas flavonoids lacking 3-OH are twisted. As was suggested in that study, blumeatin is also twisted as it lacks a 3-OH moiety. In their frontier work, they demonstrated that the ring B torsion angle with the rest of the compound is in correlation with the scavenging activity owing to the enhanced conjugation. In their work, they presented that rutin with a sugar moiety makes ring B lose its coplanarity, and this was proposed as an explanation of rutin’s less active scavenging profile because it is not able to use its full delocalization potential compared to quercetin which is almost in planar orientation. They also explained that luteolin exhibits a good antioxidant profile because it possesses a twisted torsional angle of 16.29° without a 3-OH moiety. Blumeatin also lacks the 3-OH moiety and twisted torsional angle and exhibited acceptable computed RSA.

Sadasivan and co-workers60,61 discussed the intramolecular hydrogen bonds between ring A and the keto group in ring C and the planarity of mearnsetin and myricetin (in their both neutral and radical forms) which is an indication of possible extended conjugation and their antioxidant activity. Based on the bond dissociation energy (BDE) computations, they found that H atom transfer (HAT) from the B ring is easier compared to that from the A ring (C5- and C7–OH) and C ring (C3–OH) for mearnsetin and myricetin. It was also shown that HAT is energetically more favorable from the B ring compared to the C and A rings.61 The uniform charge density distribution among the rings was also explained by the planar arrangement of the rings. To the best of our knowledge, their work is the only detailed and systematic work that performed the conformational analysis together with the discussion of a possible correlation between the planarity, charge distribution, BDEs, HAT mechanism, and spin densities of selected flavonoids.

A systematic report on calculations of antioxidant action mechanisms of selected nine isoflavones focused primarily on revealing ionization potentials (IPs), proton dissociation enthalpies, proton affinities, hydroxyl bond dissociation enthalpies (BDEs), and electron-transfer enthalpies in both gas and solution phases.62 In that study, authors demonstrated that the lowest BDEs were determined for 3′- and 4′-OH groups in ring B. For the ionization potentials, it was emphasized that ring B possesses an essential role, particularly if it has a 4′-O-methyl group.62

Attempts have been made to the determination of hydrogen abstraction energies for selected flavonols by DFT methods together with experimental techniques for antioxidant and antiradical activities using ABTS [2,2′-azino-bis(3-ethylbenzothiazoline-6-sulfonic acid)] and DPPH (1,1-diphenyl-2-picrylhydrazyl) radicals and ferric ions (FRAP) tests.63 Sroka and co-workers63 focused on the 10 different flavonols including luteolin, kaempferol, isorhamnetin, myricetin, rhamnetin, quercetin, apigenin, genkwanin, diosmetin, and chrysoeriol. In their study, they showed that the most active antiradical compounds with a C3–OH group were present. Flavones without that group were found to be less active. They discussed the positions of the atoms and their activity in detail and suggested that the positions related to the activity were mostly the positions of the OH groups located at C4′, C3, and C3′. Based on the work published by Rice-Evans and coinvestigators,64 Sroka et al.63 agreed that OH groups in ring A which are located at C5 and C7 are not important.

Semiempirical methods like PM6 (Parameterization Method 6) implemented in MOPAC (Molecular Orbital PACkage) software were used to demonstrate the correlation between the electrochemical oxidation potential (EOP), antioxidant activity, and spin population of flavonoid radicals in water. The author tried to show the correlation between flavonoid and antioxidant activity by checking atomic orbital spin populations over the skeleton atoms of a radical molecule and stated that there is an excellent correlation.65 In a study where hydrogen bonding formation was discussed, a series of aminophenyl derivatives of kaempferol were synthesized, and intermolecular hydrogen bonding was shown to play a crucial role in primary aminomethyl product formation of kaempferol. Moderate to potent cytotoxic activity against HeLa, HCC1954, and SK-OV-3 human cancer cell lines has also been demonstrated in that work.66

A group of flavonoids’ ability to scavenge the DPPH radical was elucidated by means of DFT-based quantum chemical descriptors such as frontier electron density, group philicity, and hardness. It was stated that the obtained QSAR by DFT showed that the lower the global hardness, the higher the activity. Furthermore, four new flavonoids were proposed in that study based on a QSAR model they suggested.67 According to the literature, there is no study attempting to correlate the antioxidant profiles together with atomic charges, bond lengths, and bond angles. Previous studies demonstrated that the structural properties are crucial for the high antioxidant activity of flavonoids.54 The radical scavenging ability of flavonoids is known to depend on the structures and substituents of the heterocyclic A ring. Furthermore, major determinants of antioxidant activities are the existence of a double bond between C2 and C3 conjugated with the C4-oxo group carbonyl group at C4 and the C3 hydroxyl group present in flavonols.54,68,69

Global quantum chemical descriptors are chemical potential (μ = −χ), global chemical hardness (η), electrophilicity index (ω), electronegativity (χ), global softness (s), and nucleophilicity (ε). These descriptors are calculated from EHOMO (energy of the highest occupied molecular orbital) and ELUMO (energy of the lowest unoccupied molecular orbital) values obtained by quantum chemical computations.70−75Table 1 presents the comparison of blumeatin’s quantum chemical descriptors with other selected flavonoids. If calculated band gap energies are small, this suggests that they are highly reactive.57

Table 1. Comparison of Quantum Chemical Descriptorsa (in eV) of Blumeatin to Various Flavonoids.

compd EHOMO ELUMO ΔE I A χ η μ ω ε s ref
blumeatin –6.56 –1.73 –4.84 6.32 1.48 3.90 2.42 –3.90 3.14 –9.44 0.21 this work
quercetin –5.03 –2.49 –2.54 5.03 2.49 3.76 1.27 –3.76 5.57 –4.78 0.39 (77)
quercetin –5.88 –2.10 –3.78 5.88 2.10 3.99 1.89 –3.99 4.21 –7.54 0.26 (78)
taxifolin –5.44 –2.46 –2.98 5.44 2.46 3.95 1.49 –3.95 5.24 –5.89 0.34 (78)
luteolin –5.93 –1.67 –4.26 5.93 1.67 3.80 2.13 –3.80 3.39 –8.09 0.23 (79)
gallic acid –1.23 0.24 –1.47 1.23 –0.24 0.50 0.74 –0.50 0.17 –0.36 0.68 (80)
betulinic acid –6.47 0.17 –6.64 6.47 –0.17 3.15 3.32 –3.15 1.49 –10.46 0.15 (80)
lupeol –6.31 0.76 –7.07 6.31 –0.76 2.78 3.54 –2.78 1.09 –9.81 0.14 (80)
procyanidin –0.04 0.02 –0.06 0.04 –0.02 0.01 0.03 –0.01 0.00 0.00 16.67 (80)
foeniculin –0.34 –0.16 –0.18 0.34 0.16 0.25 0.09 –0.25 0.35 –0.02 5.56 (80)
morin –7.17 –0.68 –0.49 7.17 6.68 6.93 0.25 –6.93 97.87 –1.70 2.04 (81)
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a

I = −EHOMO; A = −ELUMO, stability measure: ΔE = EHOMO – ELUMO, χ = (I + A)/2, chemical potential: μ = −(I + A)/2, electrophilicity index: ω = μ2/2η, global hardness: η = −1/2 (EHOMO-ELUMO), and global softness: s = 1/η.

HOMO–LUMO gap energy for blumeatin was found to be −4.84 eV. Based on the computational results, it can be safely suggested that blumeatin exhibits better antioxidant activity than some of the previously reported flavonoids and flavonoid-related derivatives in decreasing order as follows: lupeol > betulinic acid > blumeatin > luteolin > quercetin > taxifolin > gallic acid > morin > foeniculin > procyanidin. However, chemical potential μ is a measure of escaping nature of an electron; thus, the more negative the μ, the more difficult to lose an electron. According to Table 1, blumeatin has higher stability than morin and taxifolin (thus less reactive) but is less stable (more reactive) than the rest of the compounds given in Table 1.

Electronegativity (χ) corresponds to the electron attraction capability of compounds, and based on the results in Table 1, blumeatin has the fourth biggest electronegativity property among other compounds. Hardness (η) and softness (s) descriptors are of importance particularly in defining the behavior of chemical systems since hard compounds have larger energy gap, whereas soft compounds have small energy gap.76 As a consequence, soft compounds are more polarizable. Blumeatin has the third highest hardness value (η = 2.42 eV) compared to the first two hardest compounds (lupeol (3.54 eV) and betulinic acid (3.32 eV)). Furthermore, based on the s values, lupeol is the softest compound (0.14 eV), whereas procyanidin has the highest s value (16.67 eV) and blumeatin has the third least softness (0.21 eV) according to our findings and comparison. Another descriptor electrophilicity (ω) is a measure of the energy stabilization upon saturation by the electrons from the external surrounding. It is a hint for the charge-donating capability. A good and more reactive nucleophile is defined by a lower value of ω, whereas the higher values show the existence of a good electrophile. Our comparison suggests that blumeatin exhibits medium nucleophile character due to its electrophilicity value of 3.14 eV among others. The most nucleophile compound is procyanidin, whereas the highest ω value is morin.

3.2. Vibrational Assignments

Blumeatin consists of 36 atoms with corresponding 102 vibrational wavenumbers. Normal modes of blumeatin have been assigned based on the individual detailed motion of the atoms. Computed unscaled and scaled FTIR wavenumbers and their assignments for blumeatin are given in Table 2. Experimental and computational IR spectra of blumeatin were plotted as in Figures 3 and 4, respectively. Normal modes used to determine the individual contributions for the vibrational modes are listed in Table S3.

Table 2. Vibrational Wavenumbers (cm–1), Mode Assignmentsa, and Potential Energy Distributions (%) (PED) of Blumeatin.

mode no. wavenumbers (computed) wavenumbers (scaled) wavenumbers (experimental) notation assignment with PED (%)b
1 14 14 – τring ring torsion CCCC (22) (ring C–B) + torsion OCCC (21) (ring C–B) + torsion HCC = C (11) (ring C–B) + torsion HCCC (11) (ring C–B)
2 36 35 – τring ring torsion (12) (ring C–B)
3 54 52 – τring ring torsion (19) (ring C–B)
4 59 57 – τring ring torsion CCOC (12) (ring A-methyl) + torsion COCH (10) (methoxy)
5 81 78 – τring ring torsion (26) (ring C)
6 105 102 – γmethyl (CH3 twist + ring torsion) (11) (ring A) + ring torsion (10) (A-C)
7 162 157 – τring ring torsion (21) (ring C)
8 180 174 – ωCHb CH2 wagging (ring A)
9 189 183 – γCH CH2 twist (methyl) (27) (ring A)
10 212 205 – γCH CH2 twist (methyl) (12) (ring A)
11 222 215 – γOH OH o.o.p. bending (20) (ring B) + ring torsion (16) (ring B)
12 233 225 – τring ring torsion (87) (ring B)
13 236 228 – γCH CH2 twist (methyl) (27) (ring A)
14 248 240 – τring ring torsion (23) (ring B)
15 255 247 – τring ring torsion (21) (ring A-C) + methyl torsion (15) (ring A)
16 290 280 – τring CCOH torsion (15) (ring A)
17 297 287 – τring CCOH torsion (10) (ring A)
18 310 300 – γOH OH o.o.p. bending (69) (ring B)
19 329 318 – γOH OH o.o.p. bending (52) (ring B)
20 338 327 – δCOH COH bending (16) (ring B)
21 348 337 – τringb CCOH torsion (ring A)
22 392 379 – γOH OH o.o.p. bending (46) (ring A)
23 417 403 419 τmethyl (methyl + CCOH) torsion (11) (ring A)
24 455 440 – δring ring bending (12) (ring C)
25 478 462 459 τCH CH2 twist (16) (ring C)
26 522 505 499 τringb ring torsion (ring B)
27 526 509 507 τringb ring torsion (ring A + B)
28 536 518 – γCHb CH2 twist (ring C) + ring torsion (ring C)
29 546 528 528 γCHb CH2 twist (ring C)
30 561 542 553 γCHb CH2 twist (ring C)
31 594 574 563 ωCH3b CH3 wagging (ring A) + ring torsion (ring B)
32 615 595 597 γCH CH o.o.p. bending (ring B) (30) + ring torsion (17) (ring B)
33 620 600 582 γCH CCH o.o.p. bending (14) (ring A) + ring torsion (11) (ring A)
34 631 610 606 γCHb CCH o.o.p. bending (ring B)
35 638 617 621 γCHb CCH o.o.p. bending (ring A)
36 652 630 631 τring ring torsion (17) (ring A)
37 671 649 656 τringb ring torsion (ring C)
38 687 664 664 γCH CH o.o.p. bending (26) (ring B)
39 713 689 688 γCH CCH o.o.p. bending (12) (ring B)
40 746 721 715 γCH CCH o.o.p. bending (16) (ring A)
41 781 755 759 γCH CCH o.o.p. bending (13) (ring A)
42 792 766 – γCH CCH o.o.p. bending (21) (ring A)
43 819 792 – γCH CH o.o.p. bending (41) (ring B)
44 834 806 – γCH CH o.o.p. bending (42) (ring B)
45 844 816 815 γCH CH o.o.p. bending (28) (ring A)
46 875 846 858 γCH CH o.o.p. bending (38) (ring A)
47 896 866 874 γCH CH2 twist (20) (ring C)
48 974 942 939 τringb ring torsion (ring C) + ring bending (ring A) + OC stretching (methyl)
49 996 963 964 τringb ring torsion (ring C) + OC stretching (methyl)
50 1007 974 – δringb ring bending (ring B)
51 1011 978 977 δringb ring bending (ring B)
52 1027 993 998 δCCHb CCH bending (ring B) + ring bending (ring B)
53 1054 1019 1028 νringb ring breathing (ring A) + OC stretching (methyl)
54 1079 1043 – νCCb CC stretching (ring C) + CO stretching (ring C)
55 1088 1052 – νCOb C–O stretching (methyl) + CCH bending (ring A) + CC stretching (ring C) + CCH bending (ring C)
56 1097 1061 1068 δCCHb CCH bending (ring A) + COH bending (ring A) + CO stretching (ring C)
57 1165 1127 1118 δCCHb CCH bending (ring B) + COH bending (ring B)
58 1167 1128 – δCCH CH rocking (methyl) (36) (ring A)
59 1172 1133 – δCOH CCH bending (23) (ring B) + COH bending (11) (ring B)
60 1178 1139 – δCCH CCH bending (19) (ring A + C)
61 1189 1150 1151 δCCH CCH + COH bending (15) (ring B)
62 1201 1161 – δCCH CCH bending (24) (ring A + C + B)
63 1208 1168 – δCOHb COH bending (ring A)
64 1215 1175 – δCOH COH bending (19) (ring B)
65 1220 1180 1189 δCCH CCH bending (21) (ring A + C)
66 1244 1203 1204 δCCH CCH bending (28) (ring A + C + B)
67 1278 1236 1244 δHCH CH wagging (19) (ring C)
68 1293 1250 1257 δCCHb CH bending (ring A)+ OH bending (ring A) + ring breathing (ring A)
69 1314 1271 1271 δCOH COH bending (41) (ring B)
70 1345 1301 1302 δCCHb CH bending (ring C + ring B) + ring stretching (ring B)
71 1356 1311 – δOCH CH bending (19) (ring C)
72 1366 1321 – νringb ring stretching (ring A) + OCH bending (ring C) + ring stretching (ring B) + COH bending (ring B)
73 1372 1327 1331 δOCHb OCH bending (ring C) + COH bending (ring A) + COH bending (ring B)
74 1384 1338 1346 δOCHb OCH bending (ring C) + CO stretching (ring A) + methyl umbrella
75 1403 1357 1361 δCCH CH bending (10) (ring C)
76 1454 1406 1407 δCCH CH2 scissoring (22) (ring C)
77 1463 1415 1419 δCCH methyl umbrella (24) + OCH bending (25) (ring A)
78 1477 1428 1436 δCCH methyl umbrella (49)
79 1493 1444 1439 δHCH HCH bending (methyl) (82) (ring A)
80 1502 1452 1450 δCCH CCH bending (14) (ring B) + HCH bending (methyl) (10) (ring A)
81 1503 1453 – δHCH HCH bending (methyl) (24) (ring A) + COCH torsion (13) (ring A)
82 1521 1471 – δCCH CCH bending (17) (ring A) + OCH bending (10) (ring A)
83 1533 1482 1499 δCCH CCH bending (23) (ring B) + ring stretching (12) (ring B)
84 1604 1551 1553 νring ring stretching (18) (ring B)
85 1645 1591 1559 νCC CC stretching (11) (ring B)
86 1649 1595 1601 νCC CC stretching (11) (ring A)
87 1660 1605 – δCCH CCH bending (19) (ring B) + CC stretching (12) (ring B)
88 1744 1686 1615 νC=O C=O stretching (15) (ring C) + CC stretching (10) (ring C)
89 2997 2862 2851 νCH CH stretching (52) (ring C)
90 3011 2876 2872 νCH methyl symmetric stretching (79) (ring A-methyl)
91 3033 2897 2909 νCH CH2 symmetric stretching (69) (ring C)
92 3073 2935 2944 νCH methyl asymmetric stretching (81) (ring A-methyl)
93 3110 2970 2980 νCH CH asymmetric stretching (58) (ring C)
94 3140 2999 2997 νCH CH stretching (79) (ring B)
95 3152 3010 – νCH CH stretching (70) (ring B)
96 3175 3032 3023 νCH CH stretching (74) (ring B)
97 3190 3046 – νCH CH stretching (71) (ring A)
98 3214 3069 – νCH CH stretching (73) (ring B)
99 3217 3072 3076 νCH CH stretching (74) (ring A)
100 3818 3646 – νOH OH stretching (90) (ring A)
101 3835 3662 – νOH OH stretching (91) (ring B)
102 3836 3663 – νOH OH stretching (91) (ring B)
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a

ν: stretching; δ: in-plane bending; γ: out-of-plane bending; τ: torsion; o.o.p.: out-of-plane; and ω: wagging.

b

Individual PED contributions ≤10% were not taken into consideration since their individual contributions are ≤1–3% and do not reach 10% when summed. In this case, tentative assignments that were visually obtained by GaussView were presented.

Figure 3.

Figure 3

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Experimental FTIR spectrum of blumeatin.

Figure 4.

Figure 4

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Computed IR spectrum of blumeatin.

Three O–H stretching vibrations of blumeatin were computed at 3663, 3662, and 3646 cm–1, respectively. These wavenumbers were previously reported to be between 3645 and 3600 cm–1 (narrow) for nonbonded hydroxy groups, 3645 and 3630 cm–1 for primary alcohols, 3635 and 3620 cm–1 for secondary alcohols, 3620 and 3540 cm–1 for tertiary alcohols, and 3640 and 3530 cm–1 for phenols.82 The wavenumber for O–H stretching computed at 3646 cm–1 for ring A was seen to be in agreement for hydroxy groups (3645 cm–1). Nevertheless, we could not observe the O–H stretching vibrations in the experimental IR spectrum of blumeatin. However, we observed a band at 3367 cm–1 as a shoulder profile, and we attributed this band to OH stretching due to the nonbonded hydroxyl group. The experimental 3189 cm–1 peak was assigned as C–H stretching, and this is in agreement with the unscaled wavenumber computed at 3190 cm–1. It should be kept in mind that since scale factors do not sufficiently predict the experimental wavenumbers for these types of flavonoids, some researchers proposed different scale factors for different regions, and thus, some scaled wavenumbers are not consistent with the experimental ones as in our case. For example, Merrick and co-workers83 suggested four different scale factors for such chromone derivatives. Thus, such overestimation between the theoretical and experimental data of blumeatin for the O–H vibrations might be overcome by using multiscaling factors, which is not in our scope in this study. In addition, our computational findings did not yield any intramolecular hydrogen bonding motif.

C–H stretching vibrations are generally known to be observed between 2800 and 3100 cm–1,84 and particularly, in-plane bending C–H vibrations were previously reported in the 1100–1500 cm–1 region while out-of-plane bendings are generally observed in the 800–1000 cm–1 region.85,86 Based on the computational infrared data of blumeatin (Table 2), we observed C–H stretching vibrations between 3072 and 2999 cm–1 and 3076 and 2851 cm–1, respectively, in the computed and experimental IR spectra, which are in line with the previous findings for C–H vibrations.85,86 Only the C=O stretching vibration of blumeatin is expected to have a very strong intensity band around 1615 cm–1, but instead, we observed two satellite peaks (Figure 5) at 1635 and 1603 cm–1 at both sides of a split peak at 1615 cm–1. This was attributed to a Fermi resonance which occurs when an overtone or a combination band has the same wavenumber or a similar wavenumber to a fundamental vibration.87 This splitting here is not a consequence of any fundamental band that might be associated with ν, 2ν, or 3ν that gives rise to 1615 cm–1. Instead, we found that two combination bands observed at 1089 and 528 cm–1 (1089 + 528 = 1617 cm–1) contributed to this Fermi resonance that caused a split at ∼1615 cm–1 and yielded two satellites at 1635 and 1603 cm–1, respectively.

Figure 5.

Figure 5

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Fermi resonance with its satellite peaks in blumeatin’s FTIR spectrum.

C–H bending vibrations were computed at 1482, 1471, 1453, 1452, 1444, 1428, 1415, 1406, 1357, 1311, 1301, 1250, 1203, 1180, 1161, 1150, and 1139 cm–1 in the predicted IR spectrum. Experimental IR spectra (Table 2) of these computed bending modes can be seen in good agreement with the reported spectra.85,86 Methyl asymmetric and symmetric bending vibrations were computed at 1471 (1475 cm–187) and 1406 cm–1 (1380 cm–187), respectively.

In the literature, the ring in-plane C–C bending modes cause relatively weak bands below 1000 cm–1.88 The ring in-plane-bending modes (ring B) for blumeatin were experimentally observed at 977 cm–1 and computed at 978 cm–1. Besides, the ring out-of-plane modes mostly appear as a group of weak bands in the experimental IR spectral range 700–100 cm–1.88 The bands below ∼800 cm–1 were mostly attributed to torsional movements of the atoms in blumeatin. We observed the γCH out-of-plane modes between 874 and 528 cm–1 in the experimental IR spectrum, and these data were found to be in agreement compared to our computed vibrational wavenumbers. Our results demonstrated that in-plane and out-of-plane C–H bands agree with each other based on the previous research.85,86 Ring torsions were computed below 509 cm–1 together with the other torsional modes presented in Table 2.

3.3. Pharmacokinetic Properties of Blumeatin

Blumeatin was evaluated for its pharmacokinetics properties, including Lipinski’s rule of five,89,90 drug-likeness, and ADME/Tox analysis. No violation of Lipinski’s rule of five parameters (the molecular weight is less than 500 g/mol, MLOGP (Moriguchi octanol–water partition coefficient) ≤ 4.15, the number of oxygen atoms is less than 10, and the number of NH or OH groups is less than 5) was found for blumeatin. Its molecular weight is 302.28 g/mol; it has six oxygens and has no NH group and less than five OH groups. pkCSM web tool48 and SwissADME interface91 were utilized to obtain the ADME/Tox data for blumeatin, and the results are presented in Table 3.

Table 3. ADME/Tox Evaluation and Molecular Descriptors for Blumeatin (*: for Human).

3.3.

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3.3.1. Absorption

Absorption was computed via Caco2 permeability, water solubility, skin permeability, human intestinal absorption, and whether the molecule is a P-glycoprotein substrate or an inhibitor. The water solubility of the compound reflects at 25 °C. The LogS (aqueous solubility) value of blumeatin was calculated as −3.53, which shows that this compound is moderately soluble in water (LogS between −3 and −4).91 The ultimate bioavailability (a drug having a value of more than 0.90 is considered readily permeable) is determined by Caco2 permeability and human intestinal absorption. Blumeatin showed poor permeability (0.48). The drugs are known to be absorbed primarily in the human intestine. Hydrophilic compounds absorbed rapidly, and a compound with greater than 30% absorbance is regarded as readily absorbed. The human intestine absorption value for blumeatin was calculated to be 81.93%, and it can be considered that blumeatin has a high absorption profile in the human intestine. Blumeatin is the substrate for P-glycoprotein; however, this flavonoid was found as neither P-glycoprotein I nor P-glycoprotein II inhibitors.

3.3.2. Distribution

pKCSM tool allows us to calculate distribution by the human volume of distribution, central nervous system permeability, blood–brain barrier, and human fraction unbound in plasma. The volume of distribution is a theoretical volume that describes the drug’s overall dose. The more the VDss, the higher a drug is distributed in tissue. For antibiotics and antivirals, a more extensive tissue dispersion is preferred. VDss is low if the log VDss value is less than −0.15, whereas the value higher than 0.45 is regarded as high.92−94 This value for blumeatin is 0.17, and this indicates that blumeatin exhibits a high VDss effect.48,93 In the case of serum proteins, many plasma drugs will appear in symmetry between the unbound and bound states. Fraction unbound to human plasma must be between 0.2 and 1.0. Blumeatin exhibits a value of 0.05, which is not in this region. The more the compounds bind to the plasma, the less effective they can transverse cellular membranes or diffuse. This means that blumeatin does not bind to the plasma, and thus it cannot transverse cellular membranes or diffuse.48

3.3.3. Metabolism

The drug metabolism depends on the compound being a cytochrome p450 substrate or an inhibitor. Cytochrome p450 exists in the liver and acts as a detoxifying enzyme. It is of importance to confirm whether a molecule is capable of inhibiting cytochrome p450. This cytochrome p450s inhibitors’ isoforms are CYP2C9, CYP1A2, CYP2D6, CYP2C19, and CYP3A4.48,93 Blumeatin is the only inhibitor of CYP1A2 and CYP2C19 cytochrome enzymes, which demonstrates that it will be metabolized by the enzyme’s action, suggesting that they will not be hampered through the body’s biological transformation. Blumeatin is a noninhibitor of the rest of the cytochrome enzymes given in Table 3.

3.3.4. Excretion

Excretion is expressed by total clearance and whether a compound is a renal OCT2 substrate. Organic cation transporter 2 (OCT2) is a renal uptake transporter that deposits drugs into the kidney. Based on our results, blumeatin is not a renal OCT2 substrate, and it showed a total renal clearance of 0.52 which is less than 1 mL/min/kg, and it is found not to be a renal OCT2 substrate.

3.3.5. Toxicity

The AMES test with a negative value shows that the compound is nonmutagenic and noncarcinogenic. Blumeatin showed no AMES toxicity. Besides, the maximum recommended tolerance dose (MRTD) gives a prediction for the toxic dose in humans. MRTD ≤ log 0.477 (mg/kg/day) is known as low48 and blumeatin was found to have low toxicity to humans (Table 3). hERG (human ether-a-go-go gene) is in charge of blocking potassium channels.48,93,95 Blumeatin is a noninhibitor of both hERG1 and hERG2. It is nonskin sensitive, and it also does not induce hepatotoxicity. For a certain compound, the LD50 value corresponds to the amount that kills 50% of the test animals. Blumeatin showed an oral rat acute toxicity of 2.56 mol/kg, which was considered to be not toxic.

3.4. Molecular Docking Study of Blumeatin–Xanthine Oxidase System

It is recognized that hydrogen bonding is critical for secondary and tertiary structural protein structures. In molecular dynamics calculations, hydrogen bond interaction between a ligand and a protein region reflects the binding ability of a drug toward a protein target; therefore, the bigger the amount of H-bonds, the greater the interactions.96,97 Blumeatin has six hydrogen bond acceptors and three bond donors, and it is a relatively good candidate for potential hydrogen bonding source. Data obtained from molecular docking investigation are given in Table 4. Detailed docking orientation of blumeatin with its neighbor residues is shown in Figure 6. The two-dimensional representation of the docking environment of blumeatin into XO is demonstrated in Figure 7. Binding affinity for blumeatin to XO is found to be −9.2 kcal/mol, and this is in line with and close to several flavonoids and their binding affinities for XO (−9.993, −9.987, −9.563, −9.289, −9.252, −9.221, −9.158, −9.145, −8.857, −8.842, −8.791, −8.775, and −8.860 kcal/mol for quercetin, luteolin, tectochrysin, kaempferol, naringenin, genkwanin, myricetin, acacetin, apigenin, hesperetin, luteolin-3′-methyl ether, quercetin-3-methyl ether, and eriodictyol, respectively).98 Any interactions with crucial catalytic residues such as Glu1261 were not observed. This finding supports that the previously reported observation states that the existence of Glu1261 is crucial to block the XO activity.99−102

Table 4. Comparison of the Polar (Hydrogen Bonds) and Nonpolar Interactions (Hydrophobic, van der Waals Contacts) of Blumeatin and Quercetin (Natural Ligand) with the Residues from the Active Site of XO.

ligand conventional hydrogen bonds, bond lengths (in Å in parentheses), and typesa residues in van der Waals contact or hydrophobic interaction
blumeatin via O atom of Thr1010: 2.70:C=O and 3.26:O–H; via N atom of Thr1010: 2.93:C=O and 2.97:O–H; Val1011 (3.08:C=O); Arg880 (2.03:O–H); Phe914 (3.48:π–π and 4.04:(O–H) π–σ); Phe1009 (π–π); Ala1078 (3.58:O–H and 5.22:π–alkyl); Ala1079 (3.96:π–σ); Leu1014 (3.47:π–σ); Leu873 (4.74:π–alkyl), Leu648 (5.05:π–alkyl), Ala910 (4.37:π–alkyl) Glu802, Asn768, Ser1008, Glu1261, Phe1013, Phe1150, Ser876, Lys771, Thr803
quercetin Arg880, Thr1010, Glu802, Phe914, Ala1079, Phe1009, Leu873, Leu1014, Leu648, Val1011 Ser1008, Phe1150, Ser876, Phe1013, Lys771, Pro1076, Ala1078
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a

Bold text denotes mutual residues that are in interaction with both blumeatin and quercetin.

Figure 6.

Figure 6

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Docked pose of blumeatin into XO.

Figure 7.

Figure 7

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Binding mode of blumeatin on XO and its interaction with the surrounding residues.

As can be seen in Table 4, the natural ligand (quercetin) of XO exhibits a hydrophobic bond type with Ser1008, Phe1150, Ser876, Phe1013, Lys771, Pro1076, and Ala1078 residues, whereas it is in interaction with Arg880, Thr1010, Glu802, Phe914, Ala1079, Phe1009, Leu873, Leu1014, Leu648, and Val1011. It is interesting to note that no hydrogen-bonded interaction was observed between quercetin and Ala 910 and Ala1078, whereas blumeatin is in contact with these two residues by pi-alkyl type interaction. According to Table 4, eight conventional hydrogen bonds formed between Thr1010 (C=O and O–H), Val1011 (C=O), Arg880 (O–H), Phe914 (O–H), and Ala1078 (O–H) are 2.70, 3.26, 2.93, 2.97, 3.08, 2.03, 4.04, and 3.58 Å, respectively. Strong (almost covalent), moderate (mostly electrostatic), and weak electrostatic hydrogen bonds refer to the distances between 2.2 and 2.5, 2.5–3.2, and 3.2–4.0 Å, respectively.103,104 Based on these definitions, blumeatin binds to XO with very strong (almost covalent) bonds (2.03 Å) via Arg880, moderate hydrogen bonds (2.70, 2.93, and 2.97 Å) via Thr1010, and weak hydrogen bonds (3.26, 3.08, 4.04, and 3.58 Å) via Thr1010, Val1011, Phe914, and Ala1078, respectively. Finally, the rest of the residues in XO surrounding blumeatin formed hydrophobic interactions via Glu802, Asn768, Ser1008, Glu1261, Phe1013, Phe1150, Ser876, Lys771, and Thr803 (mainly van der Waals and hydrophobic).

Besides, based on our human 3D structure (2E1Q) study, the first nine binding structures and the interactions of the surrounding residues for blumeatin are also given in Figures S1–S9 and Table S4. Accordingly, blumeatin pose 1 (hereafter, blumeatin 1) interacts with its surroundings via two conventional hydrogen bonds over Lys1046 and Ala1080. In this docking conformation, it is also in contact with various forms of interaction like alkyl and pi-alkyl as shown in Figures S1 and S2. Blumeatin 1 has a binding affinity of −9.6 kcal/mol, which is the best docked ligand for 2E1Q. Blumeatin 2 shows conventional hydrogen bonds over Gln112, Arg913, and Ser1083. Its docking geometry and other interactions as depicted in Figures S3 and S4 are via Gly1040, Phe799, Met1039, Cys150, Gln1195, and Gln1041 residues with the binding affinity of −9.4 kcal/mol. Blumeatin 2 also has two pi–sulfur interaction types over Met1039 and Cys 150. This is the only docked pose among nine poses by pi-sulfur, amide-pi stacked, and pi–pi stacked interaction types we found for blumeatin, and even this configuration is not the best binding affinity. At this point, we have to admit that pi-sulfur, amide-pi stacked, and pi–pi stacked types of interactions are subject of a different type of study that requires more attention. Blumeatin 3 (Figure S5) is in interaction with Asn261 and Leu404 hydrogen bonds and other types of interactions by Ile353, Ile264, Leu257, Val259, Gly350, and Ser347 residual neighbors (Figure S6). We found its binding affinity to be −9.2 kcal/mol. Fourth hit after our docking effort is blumeatin 4, and this ligand exhibits conventional hydrogen bonds over Leu404 and Ile264 residues. Other types of interactions for this pose of blumeatin 4 were seen between Val259, Ile353, Leu257, val259, and Glu402 residues with a binding affinity of −9.2 kcal/mol. Blumeatin 5 (Figure S9) was observed to construct hydrogen bonds with Gly260, Asn261, and other bonds via Leu257, Ile403, Ile 353, Val259, Ala346, Pro281, Val258, Ser347, and Thr262 residues (Figure S10). It binds to the natural ligand’s pocket of 2E1Q with a binding affinity of −9.0 kcal/mol. Our findings demonstrated that blumeatin 6 (Figure S11) is in interaction with the binding affinity of −9.0 kcal/mol with its environment via hydrogen bonds over Leu404, Lys256, Lys249, Gly350, and Ala301 but other interaction types with Val259, Gly349, Glu402, Ser399, Ile353, and Leu257 residues (Figure S12). This ligand conformation is the one that has the largest number of hydrogen bond contacts among other docked poses. Besides, when blumeatin 7 (−9.0 kcal/mol of binding affinity) is considered (Figure S13), it was seen that it binds to the protein via conventional hydrogen bonding over Leu404, Glu263, and Gly260. Other residues that are shown in Figure S14 (Thr262, Val259, Asn261, Ala346, Ser347, Gly350, Ley257, Ile353, Leu287, Pro281, and Ile403) were seen to be in contact with the pi-donor hydrogen bond, alkyl, pi-alkyl, and van der Waals types of interactions. Blumeatin 8 (Figure S15) exhibits three hydrogen bonding interactions and six various types of interactions as given in Figure S16. These residues are given in Table S4. This conformation’s binding affinity was found to be −8.8 kcal/mol. Two hydrogen bonds were seen to form between Val1260 and Gln1041 for blumeatin 9 (Figure S17) docking pose with a binding affinity of −8.7 kcal/mol. Ala1079, Gln1195, Arg913, Gly1261, Lys1046, Leu1403, Met1039, Phe799, and Ala1084 residues are in different types of interactions rather than hydrogen bonding as presented in Figure S18.

In addition to the discussion above, recent studies showed that blumeatin was also studied by docking strategies, and it was shown that it was docked with the highest score against N-myristoyltransferase as an inhibitor together with other various flavonoids including naringin. In their study, Hadi and Nastiti also demonstrated that blumeatin interacted with Tyr, Leu, Asn, and Cys residues.103 Another study focused on a detailed review on selected particular flavonoids including blumeatin and their chemical constituents, and bioactivities were evaluated, but their study did not give any detailed information on blumeatin.104 On the other hand, two selected flavonoids (blumeatin and luteolin) were also docked against caspase-1 by optimizing these two ligands by semiempiric optimization method (AM1), and it was considered that blumeatin and luteolin were suggested to be anti-inflammatory agents.105 Another recent publication by Xia et al. in 2023 stated that blumeatin was synthesized and confirmed its hydroxyl groups at positions C-3′ and C-5′ by experimental NMR spectroscopy, and they only reported selected IR peaks rather than detailed individual vibrational modes as we present here. They also mentioned that the isolation procedure of this flavonoid from plants has limitations of prolonged duration and high cost.106

4. Conclusions

In this study, the first computed and experimental IR spectra of blumeatin were presented. Computational results indicated no intramolecular hydrogen bonding for blumeatin. Many investigations separately focused on the geometric structures of flavonoids and tried to reveal structure–activity relationships only based on selected several structural parameters and charge-transfer mechanisms. To the best of our knowledge, there are not many attempts to correlate the planarity, antioxidant activities, and experimental and theoretical IR spectra at the same time. Blumeatin showed better theoretical radical scavenging activity compared with several flavonoids. Our attempt on constructing a link between the geometric structure and the activity of blumeatin would provide a stimulus for further systematic and detailed experimental and theoretical research that is required to better reveal the robust connections between action mechanisms and geometric descriptors of flavonoids. To figure out the roles of several quantum chemical descriptors/parameters in the antioxidant activities of both neutral and radical flavonoids and better understand their action mechanism and nature, some questions are crucial to ask. How do (i) the positions of the hydroxyl and other functional groups (methyl, carbonyl, etc.), (ii) spin densities and localizations on the individual atoms in the flavonoid, and (iii) existing intra/intermolecular bonding and the numbers of these bonds affect the nature of the action of flavonoids? For these purposes, both systematic theoretical (conformational analyses, determination of spin densities, HOMO–LUMO study, etc.) and experimental future work are required to unravel the whole picture.

Acknowledgments

No funding was received for the performed study.

Supporting Information Available

The Supporting Information is available free of charge at https://pubs.acs.org/doi/10.1021/acsomega.3c10083.

  • Distance types and distances in blumeatin, optimized geometric structure of blumeatin, normal modes used in the determination of vibrational modes and their contribution, comparison of the polar (hydrogen bonds), and nonpolar interactions (hydrophobic, van der Waal’s contacts) of blumeatin with the residues from the active site of 2E1Q PDF

Author Contributions

Cisem Altunayar-Unsalan contributed to conceptualization, methodology, formal analysis, investigation, data curation, writing—original draft, and writing—review and editing. Ozan Unsalan contributed to conceptualization, methodology, formal analysis, investigation, resources, data curation, writing—original draft, and writing—review and editing.

The authors declare no competing financial interest.

Supplementary Material

ao3c10083_si_001.pdf (2.9MB, pdf)

References

  1. Wang Y.; Zhang S.; Zhang Y.; Yao F.; Zhao G.; Wang J.; Liu L.; Yang Y.; Li X.; Sun Y.; Hu Y.; Bai Z.; Wang P.; Li R.; Xu X. Screening and Evaluation of Flavonoids as Xanthine Oxidase Inhibitors for Reducing Uric Acid through Combined Cell Biology and Molecular Simulation. ACS Food Sci. Technol. 2021, 1 (11), 2182–2191. 10.1021/acsfoodscitech.1c00348. [DOI] [Google Scholar]
  2. Nutmakul T. A Review on Benefits of Quercetin in Hyperuricemia and Gouty Arthritis. Saudi Pharm. J. 2022, 30 (7), 918–926. 10.1016/j.jsps.2022.04.013. [DOI] [PMC free article] [PubMed] [Google Scholar]
  3. Ajala O. S.; Ayeleso A. O.; Owolabi M.; Akinleye M. O.; Ukpo G. Xanthine Oxidase Inhibitory Potentials of Flavonoid Aglycones of Tribulus Terrestris : In Vivo, in Silico and in Vitro Studies. Future J. Pharm. Sci. 2022, 8 (1), 58. 10.1186/s43094-022-00448-y. [DOI] [Google Scholar]
  4. Zeng N.; Zhang G.; Hu X.; Pan J.; Gong D. Mechanism of Fisetin Suppressing Superoxide Anion and Xanthine Oxidase Activity. J. Funct. Foods 2019, 58, 1–10. 10.1016/j.jff.2019.04.044. [DOI] [Google Scholar]
  5. Wang J.; Sun S.; Zhao K.; Shi H.; Fan J.; Wang H.; Liu Y.; Liu X.; Wang W. Insights into the Inhibitory Mechanism of Purpurogallin on Xanthine Oxidase by Multiple Spectroscopic Techniques and Molecular Docking. J. Mol. Struct. 2021, 1228, 129772. 10.1016/j.molstruc.2020.129772. [DOI] [Google Scholar]
  6. Ou R.; Lin L.; Zhao M.; Xie Z. Action Mechanisms and Interaction of Two Key Xanthine Oxidase Inhibitors in Galangal : Combination of in Vitro and in Silico Molecular Docking Studies. Int. J. Biol. Macromol. 2020, 162, 1526–1535. 10.1016/j.ijbiomac.2020.07.297. [DOI] [PubMed] [Google Scholar]
  7. Lin S.; Zhang G.; Liao Y.; Pan J. Inhibition of Chrysin on Xanthine Oxidase Activity and Its Inhibition Mechanism. Int. J. Biol. Macromol. 2015, 81, 274–282. 10.1016/j.ijbiomac.2015.08.017. [DOI] [PubMed] [Google Scholar]
  8. Zhang M.; Li Y.; Zhu T. The Theoretical Investigation of Monohydroxy Flavone : A Combined DFT and Molecular Docking Study. J. Mol. Struct. 2022, 1250, 131823. 10.1016/j.molstruc.2021.131823. [DOI] [Google Scholar]
  9. Wang Y.; Zhang G.; Pan J.; Gong D. Novel Insights into the Inhibitory Mechanism of Kaempferol on Xanthine Oxidase. J. Agric. Food Chem. 2015, 63, 526–534. 10.1021/jf505584m. [DOI] [PubMed] [Google Scholar]
  10. Li S.; Dong P.; Wang J.; Zhang J.; Gu J.; Wu X.; Wu W.; Fei X.; Zhang Z.; Wang Y.; Quan Z.; Liu Y. Icariin, a Natural Flavonol Glycoside, Induces Apoptosis in Human Hepatoma SMMC-7721 Cells via a ROS/JNK-Dependent Mitochondrial Pathway. Cancer Lett. 2010, 298 (2), 222–230. 10.1016/j.canlet.2010.07.009. [DOI] [PubMed] [Google Scholar]
  11. Li H.; Huang J.; Yang B.; Xiang T.; Yin X.; Peng W.; Cheng W.; Wan J.; Luo F.; Li H.; Ren G. Mangiferin Exerts Antitumor Activity in Breast Cancer Cells by Regulating Matrix Metalloproteinases, Epithelial to Mesenchymal Transition, and β-Catenin Signaling Pathway. Toxicol. Appl. Pharmacol. 2013, 272 (1), 180–190. 10.1016/j.taap.2013.05.011. [DOI] [PubMed] [Google Scholar]
  12. Reutrakul V.; Ningnuek N.; Pohmakotr M.; Yoosook C.; Napaswad C.; Kasisit J.; Santisuk T.; Tuchinda P. Anti HIV-1 Flavonoid Glycosides from Ochna Integerrima. Planta Med. 2007, 73 (7), 683–688. 10.1055/s-2007-981538. [DOI] [PubMed] [Google Scholar]
  13. Xiao J.; Capanoglu E.; Jassbi A. R.; Miron A. Advance on the Flavonoid C-Glycosides and Health Benefits. Crit. Rev. Food Sci. Nutr. 2016, 56 (sup1), S29–S45. 10.1080/10408398.2015.1067595. [DOI] [PubMed] [Google Scholar]
  14. Khalil H.; Abd El Maksoud A. I.; Roshdey T.; El-Masry S. Guava Flavonoid Glycosides Prevent Influenza A Virus Infection via Rescue of P53 Activity. J. Med. Virol. 2019, 91 (1), 45–55. 10.1002/jmv.25295. [DOI] [PubMed] [Google Scholar]
  15. Cherrak S. A.; Merzouk H.; Mokhtari-Soulimane N. Potential Bioactive Glycosylated Flavonoids as SARS-CoV-2 Main Protease Inhibitors: A Molecular Docking and Simulation Studies. PLoS One 2020, 15, e0240653 10.1371/journal.pone.0240653. [DOI] [PMC free article] [PubMed] [Google Scholar]
  16. Tasdemir D.; Kaiser M.; Brun R.; Yardley V.; Schmidt T. J.; Tosun F.; Rüedi P. Antitrypanosomal and Antileishmanial Activities of Flavonoids and Their Analogues: In Vitro, in Vivo, Structure-Activity Relationship, and Quantitative Structure-Activity Relationship Studies. Antimicrob. Agents Chemother. 2006, 50 (4), 1352–1364. 10.1128/AAC.50.4.1352-1364.2006. [DOI] [PMC free article] [PubMed] [Google Scholar]
  17. Sati P.; Dhyani P.; Bhatt I. D.; Pandey A. Ginkgo Biloba Flavonoid Glycosides in Antimicrobial Perspective with Reference to Extraction Method. J. Tradit. Complementary Med. 2019, 9 (1), 15–23. 10.1016/j.jtcme.2017.10.003. [DOI] [PMC free article] [PubMed] [Google Scholar]
  18. Leong C. N. A.; Tako M.; Hanashiro I.; Tamaki H. Antioxidant Flavonoid Glycosides from the Leaves of Ficus Pumila L. Food Chem. 2008, 109 (2), 415–420. 10.1016/j.foodchem.2007.12.069. [DOI] [PubMed] [Google Scholar]
  19. Kumar M.; Ahmad A.; Rawat P.; Khan M. F.; Rasheed N.; Gupta P.; Sathiamoorthy B.; Bhatia G.; Palit G.; Maurya R. Antioxidant Flavonoid Glycosides from Evolvulus Alsinoides. Fitoterapia 2010, 81 (4), 234–242. 10.1016/j.fitote.2009.09.003. [DOI] [PubMed] [Google Scholar]
  20. Wen L.; Zhao Y.; Jiang Y.; Yu L.; Zeng X.; Yang J.; Tian M.; Liu H.; Yang B. Identification of a Flavonoid C -Glycoside as Potent Antioxidant. Free Radical Biol. Med. 2017, 110, 92–101. 10.1016/j.freeradbiomed.2017.05.027. [DOI] [PubMed] [Google Scholar]
  21. Bhardwaj V. K.; Singh R.; Sharma J.; Rajendran V.; Purohit R.; Kumar S. Identification of Bioactive Molecules from Tea Plant as SARS-CoV-2 Main Protease Inhibitors. J. Biomol. Struct. Dyn. 2021, 39 (10), 3449–3458. 10.1080/07391102.2020.1766572. [DOI] [PMC free article] [PubMed] [Google Scholar]
  22. Bitew M.; Desalegn T.; Demissie T.; Belayneh A.; Endale M.; Eswaramoorthy R. Pharmacokinetics and Drug-Likeness of Antidiabetic Flavonoids : Molecular Docking and DFT Study. PLoS One 2021, 16 (12), e0260853 10.1371/journal.pone.0260853. [DOI] [PMC free article] [PubMed] [Google Scholar]
  23. Xu S. B.; Chen W. F.; Liang H. Q.; Lin Y. C.; Deng Y. J.; Long K. H. Protective Action of Blumeatin against Experimental Liver Injuries. Acta Pharmacol. Sin. 1993, 14 (4), 376–378. [PubMed] [Google Scholar]
  24. Unsalan O.; Erdogdu Y.; Gulluoglu M. T. FT-Raman and FT-IR Spectral and Quantum Chemical Studies on Some Flavonoid Derivatives: Baicalein and Naringenin. J. Raman Spectrosc. 2009, 40 (5), 562–570. 10.1002/jrs.2166. [DOI] [Google Scholar]
  25. Erdoǧdu Y.; Ünsalan O.; Güllüoğlu M. T. Vibrational Analysis of Flavone. Turk. J. Phys. 2009, 33 (5), 249–259. 10.3906/fiz-0809-8. [DOI] [Google Scholar]
  26. Erdogdu Y.; Unsalan O.; Amalanathan M.; Hubert Joe I. Infrared and Raman Spectra, Vibrational Assignment, NBO Analysis and DFT Calculations of 6-Aminoflavone. J. Mol. Struct. 2010, 980 (1–3), 24–30. 10.1016/j.molstruc.2010.06.032. [DOI] [Google Scholar]
  27. Erdogdu Y.; Unsalan O.; Gulluoglu M. T. FT-Raman, FT-IR Spectral and DFT Studies on 6, 8-Dichloroflavone and 6,8-Dibromoflavone. J. Raman Spectrosc. 2010, 41 (7), 820–828. 10.1002/jrs.2520. [DOI] [Google Scholar]
  28. Erdogdu Y.; Unsalan O.; Sajan D.; Gulluoglu M. T. Structural Conformations and Vibrational Spectral Study of Chloroflavone with Density Functional Theoretical Simulations. Spectrochim. Acta, Part A 2010, 76 (2), 130–136. 10.1016/j.saa.2010.02.043. [DOI] [PubMed] [Google Scholar]
  29. Erdogdu Y.; Dereli O. ¨.; Sajan D.; Joseph L.; Unsalan O.; Gulluoglu M. T. Vibrational (FT-IR and FT-Raman) Spectral Investigations of 7-Aminoflavone with Density Functional Theoretical Simulations. Mol. Simul. 2012, 38 (4), 315–325. 10.1080/08927022.2011.632416. [DOI] [Google Scholar]
  30. Altunayar-Unsalan C.; Unsalan O.; Mavromoustakos T. Insights into Molecular Mechanism of Action of Citrus Flavonoids Hesperidin and Naringin on Lipid Bilayers Using Spectroscopic, Calorimetric, Microscopic and Theoretical Studies. J. Mol. Liq. 2022, 347, 118411. 10.1016/j.molliq.2021.118411. [DOI] [Google Scholar]
  31. Altunayar-Unsalan C.; Unsalan O.; Mavromoustakos T. Molecular Interactions of Hesperidin with DMPC/Cholesterol Bilayers. Chem. Biol. Interact. 2022, 366, 110131. 10.1016/j.cbi.2022.110131. [DOI] [PubMed] [Google Scholar]
  32. Ertan-Bolelli̇ T.; Bolelli̇ K.; Altunayar-Unsalan C.; Ünsalan O.; Yilmaz B. Effects of Flavonoids on SARS-CoV-2 Main Protease (6W63): A Molecular Docking Study. Turk. Comput. Theor. Chem. 2023, 7 (2), 34–57. 10.33435/tcandtc.1151841. [DOI] [Google Scholar]
  33. Amić D.; Davidović-Amić D.; Bešlo D.; Trinajstić N. Structure-Radical Scavenging Activity Relationships of Flavonoids. Croat. Chem. Acta 2003, 76 (1), 55–61. [Google Scholar]
  34. Cao G.; Sofic E.; Prior R. L. Antioxidant and Prooxidant Behavior of Flavonoids: Structure-Activity Relationships. Free Radical Biol. Med. 1997, 22 (5), 749–760. 10.1016/S0891-5849(96)00351-6. [DOI] [PubMed] [Google Scholar]
  35. Rice-Evans C.; Miller N.; Paganga G. Antioxidant Properties of Phenolic Compounds. Trends Plant Sci. 1997, 2 (4), 152–159. 10.1016/S1360-1385(97)01018-2. [DOI] [Google Scholar]
  36. Pratama I. P. A. A. C.; Putra I. M. H.; Pujasari L. W. S.; Dewi K. D. M. S.; Laksmiani N. P. L. The Potency of Blumeatin and Luteolin as Caspase-1 Inhibitor by Molecular Docking. Pharm. Rep. 2022, 2 (1), 22. 10.51511/pr.22. [DOI] [Google Scholar]
  37. Nessa F.; Ismail Z.; Mohamed N.; Haris M. R. H. M. Free Radical-Scavenging Activity of Organic Extracts and of Pure Flavonoids of Blumea Balsamifera DC Leaves. Food Chem. 2004, 88 (2), 243–252. 10.1016/j.foodchem.2004.01.041. [DOI] [Google Scholar]
  38. Xu J.; Jin D.; Liu C.; Xie C.; Guo Y.; Fang L. Isolation, Characterization, and NO Inhibitory Activities of Sesquiterpenes from Blumea Balsamifera. J. Agric. Food Chem. 2012, 60 (32), 8051–8058. 10.1021/jf302530u. [DOI] [PubMed] [Google Scholar]
  39. Nessa F.; Ismail Z.; Mohamed N. Xanthine Oxidase Inhibitory Activities of Extracts and Flavonoids of the Leaves of Blumea Balsamifera. Pharm. Biol. 2010, 48 (12), 1405–1412. 10.3109/13880209.2010.487281. [DOI] [PubMed] [Google Scholar]
  40. OriginPro Version 2022b. OriginLab Corporation, Northampton, MA, USA., 2022.
  41. Frisch M. J.; Trucks G. W.; Schlegel H. B.; Scuseria G. E.; Robb M. A.; Cheeseman J. R.; Scalmani G.; Barone V.; Mennucci B.; Petersson G. A.; Nakatsuji H.; Caricato M.; Li X.; Hratchian H. P.; Izmaylov A. F.; Bloino J.; Zheng G.; Sonnenberg J. L.; Hada M.; Ehara M.; Toyota K.; Fukuda R.; Hasegawa J.; Ishida M.; Nakajima T.; Honda Y.; Kitao O.; Nakai H.; Vreven T.; Montgomery J. A. Jr.; Peralta J. E.; Ogliaro F.; Bearpark M.; Heyd J. J.; Brothers E.; Kudin K. N.; Staroverov V. N.; Kobayashi R.; Normand J.; Raghavachari K.; Rendell A.; Burant J. C.; Iyengar S. S.; Tomasi J.; Cossi M.; Rega N.; Millam J. M.; Klene M.; Knox J. E.; Cross J. B.; Bakken V.; Adamo C.; Jaramillo J.; Gomperts R.; Stratmann R. E.; Yazyev O.; Austin A. J.; Cammi R.; Pomelli C.; Ochterski J. W.; Martin R. L.; Morokuma K.; Zakrzewski V. G.; Voth G. A.; Salvador P.; Dannenberg J. J.; Dapprich S.; Daniels A. D.; Farkas Ö.; Foresman J. B.; Ortiz J. V.; Cioslowski J.; Fox D. J.. Gaussian 09; Gaussian, Inc.: Wallingford, CT, 2009
  42. Becke A. D. Density-Functional Thermochemistry. III. The Role of Exact Exchange. J. Chem. Phys. 1993, 98 (7), 5648–5652. 10.1063/1.464913. [DOI] [Google Scholar]
  43. Becke A. D. Density-Functional Exchange-Energy Approximation with Correct Asymptotic Behavior. Phys. Rev. A 1988, 38 (6), 3098–3100. 10.1103/PhysRevA.38.3098. [DOI] [PubMed] [Google Scholar]
  44. Lee C.; Yang W.; Parr R. G. Development of the Colle-Salvetti Correlation-Energy Formula into a Functional of the Electron Density. Phys. Rev. B 1988, 37 (2), 785–789. 10.1103/PhysRevB.37.785. [DOI] [PubMed] [Google Scholar]
  45. Vosko S. H.; Wilk L.; Nusair M. Accurate Spin-Dependent Electron Liquid Correlation Energies for Local Spin Density Calculations: A Critical Analysis. Can. J. Phys. 1980, 58 (8), 1200–1211. 10.1139/p80-159. [DOI] [Google Scholar]
  46. Frisch M. J.; Head-Gordon M.; Pople J. A. A Direct MP2 Gradient Method. Chem. Phys. Lett. 1990, 166 (3), 275–280. 10.1016/0009-2614(90)80029-D. [DOI] [Google Scholar]
  47. Császár P.; Pulay P. Geometry Optimization by Direct Inversion in the Iterative Subspace. J. Mol. Struct. 1984, 114, 31–34. 10.1016/S0022-2860(84)87198-7. [DOI] [Google Scholar]
  48. Pires D. E. V.; Blundell T. L.; Ascher D. B. PkCSM: Predicting Small-Molecule Pharmacokinetic and Toxicity Properties Using Graph-Based Signatures. J. Med. Chem. 2015, 58 (9), 4066–4072. 10.1021/acs.jmedchem.5b00104. [DOI] [PMC free article] [PubMed] [Google Scholar]
  49. Lill M. A.; Danielson M. L. Computer-Aided Drug Design Platform Using PyMOL. J. Comput. Aided Mol. Des. 2011, 25 (1), 13–19. 10.1007/s10822-010-9395-8. [DOI] [PubMed] [Google Scholar]
  50. Dallakyan S.; Olson A. J.. Small-Molecule Library Screening by Docking with PyRx. In Chemical Biology: Methods and Protocols; Hempel J. E., Williams C. H., Hong C. C., Eds.; Springer New York: New York, NY, 2015, pp 243–250. [DOI] [PubMed] [Google Scholar]
  51. BIOVIA ; Dassault Systèmes, Discovery Studio Visualizer, V21.1.0.20298, San Diego: Dassault Systèmes, 2020.
  52. van Acker S. A. B. E.; de Groot M. J.; van den Berg D.-J.; Tromp M. N. J. L.; Donné-Op den Kelder G.; van der Vijgh W. J. F.; Bast A. A Quantum Chemical Explanation of the Antioxidant Activity of Flavonoids. Chem. Res. Toxicol. 1996, 9 (8), 1305–1312. 10.1021/tx9600964. [DOI] [PubMed] [Google Scholar]
  53. Cody V.; Luft J. R. Conformational Analysis of Flavonoids: Crystal and Molecular Structures of Morin Hydrate and Myricetin (1:2) Triphenylphosphine Oxide Complex. J. Mol. Struct. 1994, 317 (1–2), 89–97. 10.1016/0022-2860(93)07867-V. [DOI] [Google Scholar]
  54. Cody V.; Middleton E.; Harborne J. B. Plant Flavonoids in Biology and Medicine. Biochemical, Pharmacological, and Structure-Activity Relationships. Proceedings of a Symposium. Buffalo, New York, July 22–26, 1985. Prog. Clin. Biol. Res. 1986, 213, 1–592. [PubMed] [Google Scholar]
  55. Van Acker S. A. B. E.; Van Den Berg D.; Tromp M. N. J. L.; Griffioen D. H.; Van Bennekom W. P.; Van Der Vijgh W. J. F.; Bast A. Structural Aspects of Antioxidant Activity of Flavonoids. Free Radical Biol. Med. 1996, 20 (3), 331–342. 10.1016/0891-5849(95)02047-0. [DOI] [PubMed] [Google Scholar]
  56. Koç E.; Üngördü A.; Candan F. Antioxidant Activities of Alyssum Virgatum Plant and Its Main Components. Struct. Chem. 2022, 33 (1), 267–279. 10.1007/s11224-021-01856-1. [DOI] [Google Scholar]
  57. Zerrouki M.; Benkaci-Ali F. DFT Study of the Mechanisms of Nonenzymatic DNA Repair by Phytophenolic Antioxidants. J. Mol. Model. 2018, 24 (4), 78. 10.1007/s00894-018-3599-6. [DOI] [PubMed] [Google Scholar]
  58. Hu-Jun X.; Wang-Shu M.; Fu-Rong L.; Jie-Hui X.; Qun-Fang L.; Wen-Jun F. Radical Scavenging Activity of Myricetin. Acta Phys. Chim. Sin. 2013, 29 (07), 1421–1432. 10.3866/pku.whxb201304222. [DOI] [Google Scholar]
  59. Lien E. J.; Ren S.; Bui H. H.; Wang R. Quantitative Structure-Activity Relationship Analysis of Phenolic Antioxidants. Free Radical Biol. Med. 1999, 26 (3–4), 285–294. 10.1016/S0891-5849(98)00190-7. [DOI] [PubMed] [Google Scholar]
  60. Sadasivam K.; Kumaresan R. Antioxidant Behavior of Mearnsetin and Myricetin Flavonoid Compounds — A DFT Study. Spectrochim. Acta, Part A 2011, 79 (1), 282–293. 10.1016/j.saa.2011.02.042. [DOI] [PubMed] [Google Scholar]
  61. Sadasivam K.; Kumaresan R. A Comparative DFT Study on the Antioxidant Activity of Apigenin and Scutellarein Flavonoid Compounds. Mol. Phys. 2011, 109 (6), 839–852. 10.1080/00268976.2011.556576. [DOI] [Google Scholar]
  62. Lengyel J.; Rimarčík J.; Vagánek A.; Klein E. On the Radical Scavenging Activity of Isoflavones: Thermodynamics of O-H Bond Cleavage. Phys. Chem. Chem. Phys. 2013, 15 (26), 10895–10903. 10.1039/c3cp00095h. [DOI] [PubMed] [Google Scholar]
  63. Sroka Z.; Zbikowska B.; Hladyszowski J. The Antiradical Activity of Some Selected Flavones and Flavonols. Experimental and Quantum Mechanical Study. J. Mol. Model. 2015, 21 (12), 307. 10.1007/s00894-015-2848-1. [DOI] [PubMed] [Google Scholar]
  64. Rice-Evans C. A.; Miller N. J.; Paganga G. Structure-Antioxidant Activity Relationships of Flavonoids and Phenolic Acids. Free Radical Biol. Med. 1996, 20 (7), 933–956. 10.1016/0891-5849(95)02227-9. [DOI] [PubMed] [Google Scholar]
  65. Milicevic A. The Relationship between Antioxidant Activity, First Electrochemical Oxidation Potential, and Spin Population of Flavonoid Radicals. Arh. Za Hig. Rada I Toksikol. Ind. Hyg. Toxicol. 2019, 70 (2), 134–139. 10.2478/aiht-2019-70-3290. [DOI] [PubMed] [Google Scholar]
  66. Cai S. L.; Kong Y. Y.; Xiao D.; Chen Y.; Wang Q. A. Primary Aminomethyl Derivatives of Kaempferol: Hydrogen Bond-Assisted Synthesis, Anticancer Activity and Spectral Properties. Org. Biomol. Chem. 2018, 16 (11), 1921–1931. 10.1039/C7OB02927F. [DOI] [PubMed] [Google Scholar]
  67. Sarkar A.; Middya T. R.; Jana A. D. A QSAR Study of Radical Scavenging Antioxidant Activity of a Series of Flavonoids Using DFT Based Quantum Chemical Descriptors - The Importance of Group Frontier Electron Density. J. Mol. Model. 2012, 18 (6), 2621–2631. 10.1007/s00894-011-1274-2. [DOI] [PubMed] [Google Scholar]
  68. Adejoro I. A.; Akintemi E.; Adeboye O. O.; Ibeji C. Quantum Mechanical Studies of the Structure-Activity Relationship and Electronic Vibration of Some Dietary Flavonoids. Asian J. Appl. Sci. 2014, 7 (3), 117–128. 10.3923/ajaps.2014.117.128. [DOI] [Google Scholar]
  69. Marković Z. S.; J.M D.; Doličanin Ć. A PM6 Study on the Reactivity of OH Groups in Fisetin: Structural and Electronic Features of Fisetins Radicals. J. Serb. Soc. Comput. Mech. 2009, 3 (2), 43–55. [Google Scholar]
  70. Kiyooka S.; Kaneno D.; Fujiyama R. Parr’s Index to Describe Both Electrophilicity and Nucleophilicity. Tetrahedron Lett. 2013, 54 (4), 339–342. 10.1016/j.tetlet.2012.11.039. [DOI] [Google Scholar]
  71. Ayers P. W.; Parr R. G. Variational Principles for Describing Chemical Reactions: The Fukui Function and Chemical Hardness Revisited. J. Am. Chem. Soc. 2000, 122 (9), 2010–2018. 10.1021/ja9924039. [DOI] [PubMed] [Google Scholar]
  72. Flippin L. A.; Gallagher D. W.; Jalali-Araghi K. A Convenient Method for the Reduction of Ozonides to Alcohols with Borane-Dimethyl Sulfide Complex. J. Org. Chem. 1989, 54 (6), 1430–1432. 10.1021/jo00267a035. [DOI] [Google Scholar]
  73. Parr R. G.; Szentpály L. v.; Liu S. Electrophilicity Index. J. Am. Chem. Soc. 1999, 121 (9), 1922–1924. 10.1021/ja983494x. [DOI] [Google Scholar]
  74. Chattaraj P. K.; Roy D. R. Aromaticity in Polyacene Analogues of Inorganic Ring Compounds. J. Phys. Chem. A 2007, 111 (21), 4684–4696. 10.1021/jp071030s. [DOI] [PubMed] [Google Scholar]
  75. Padmanabhan J.; Parthasarathi R.; Subramanian V.; Chattaraj P. K. Electrophilicity-Based Charge Transfer Descriptor. J. Phys. Chem. A 2007, 111 (7), 1358–1361. 10.1021/jp0649549. [DOI] [PubMed] [Google Scholar]
  76. Obi-Egbedi N. O.; Obot I. B.; El-Khaiary M. I.; Umoren S. A.; Ebenso E. E. Computational Simulation and Statistical Analysis on the Relationship between Corrosion Inhibition Efficiency and Molecular Structure of Some Phenanthroline Derivatives on Mild Steel Surface. Int. J. Electrochem. Sci. 2011, 6 (11), 5649–5675. 10.1016/S1452-3981(23)18435-5. [DOI] [Google Scholar]
  77. El-Mansy M. A. M.; Suvitha A. R.; Maharani N. Y.; Vedi A. Boosted Electronic, NLO and Absorption Characteristics for Quercetin and Taxifolin: Comparative Experimental and DFT Studies. Biointerface Res. Appl. Chem. 2022, 12 (6), 7886–7902. [Google Scholar]
  78. Mir S. A.; Nayak B. In Silico Analysis of Binding Stability of Quercetin with CmpA and In Vitro Growth Inhibition Study of Cyanobacterial Species Using Azadirachta Indica Extracts. Chem. Africa 2022, 5 (3), 691–701. 10.1007/s42250-022-00335-2. [DOI] [Google Scholar]
  79. Sulaiman G. M. In Vitro Study of Molecular Structure and Cytotoxicity Effect of Luteolin in the Human Colon Carcinoma Cells. Eur. Food Res. Technol. 2015, 241 (1), 83–90. 10.1007/s00217-015-2436-8. [DOI] [Google Scholar]
  80. Teles Fujishima M.; Silva N.; Ramos R. D.; Batista Ferreira E.; Santos K.; Silva C.; Silva J.; Campos Rosa J.; Santos C. An Antioxidant Potential, Quantum-Chemical and Molecular Docking Study of the Major Chemical Constituents Present in the Leaves of Curatella Americana Linn. Pharmaceuticals 2018, 11 (3), 72. 10.3390/ph11030072. [DOI] [PMC free article] [PubMed] [Google Scholar]
  81. Payan-Gomez S. A.; Flores-Holguin N.; Perez-Hernandez A.; Pinon-Miramontes M.; Glossman-Mitnik D. Computational Molecular Characterization of the Flavonoid Morin and Its Pt(II), Pd(II) and Zn(II) Complexes. J. Mol. Model. 2011, 17 (5), 979–985. 10.1007/s00894-010-0789-2. [DOI] [PubMed] [Google Scholar]
  82. Coates J.Interpretation of Infrared Spectra, A Practical Approach. In Encyclopedia of Analytical Chemistry; Meyers R. A., Ed.; John Wiley & Sons, Ltd, 2004, pp 1–23. [Google Scholar]
  83. Merrick J. P.; Moran D.; Radom L. An Evaluation of Harmonic Vibrational Frequency Scale Factors. J. Phys. Chem. A 2007, 111 (45), 11683–11700. 10.1021/jp073974n. [DOI] [PubMed] [Google Scholar]
  84. Nakamoto K.Infrared and Raman Spectra of Inorganic and Coordination Compounds; 4.; John Wiley & Sons, Ltd: Milwaukee, 1986. [Google Scholar]
  85. Mohan J.Organic Spectroscopy: Principles and Applications; Narosa Publishing House: India, 2000. [Google Scholar]
  86. Varsányi G.Vibrational Spectra of Benzene Derivatives; Elsevier, 1969. [Google Scholar]
  87. Stuart B.Infrared Spectroscopy: Fundamentals and Applications; Wiley, 2004. [Google Scholar]
  88. Ramoji A.; Yenagi J.; Tonannavar J.; Jadhav V. B.; Kulkarni M. V. Vibrational Assignments and Electronic Structure Calculations for 3-Acetylcoumarin. Spectrochim. Acta, Part A 2007, 68 (3), 504–509. 10.1016/j.saa.2006.12.019. [DOI] [PubMed] [Google Scholar]
  89. Lipinski C. A.; Lombardo F.; Dominy B. W.; Feeney P. J. Experimental and Computational Approaches to Estimate Solubility and Permeability in Drug Discovery and Development Settings. Adv. Drug Delivery Rev. 2012, 64, 4–17. 10.1016/j.addr.2012.09.019. [DOI] [PubMed] [Google Scholar]
  90. Lipinski C. A.; Lombardo F.; Dominy B. W.; Feeney P. J. Experimental and Computational Approaches to Estimate Solubility and Permeability in Drug Discovery and Development Settings1PII of Original Article: S0169–409X(96)00423–1. The Article Was Originally Published in Advanced Drug Delivery Reviews 23 (1997) 3. Adv. Drug Delivery Rev. 2001, 46 (1–3), 3–26. 10.1016/S0169-409X(00)00129-0. [DOI] [PubMed] [Google Scholar]
  91. Daina A.; Michielin O.; Zoete V. SwissADME: A Free Web Tool to Evaluate Pharmacokinetics, Drug-Likeness and Medicinal Chemistry Friendliness of Small Molecules. Sci. Rep. 2017, 7 (1), 42717. 10.1038/srep42717. [DOI] [PMC free article] [PubMed] [Google Scholar]
  92. Bhowmik D.; Nandi R.; Jagadeesan R.; Kumar N.; Prakash A.; Kumar D. Identification of Potential Inhibitors against SARS-CoV-2 by Targeting Proteins Responsible for Envelope Formation and Virion Assembly Using Docking Based Virtual Screening, and Pharmacokinetics Approaches. Infect. Genet. Evol. 2020, 84, 104451. 10.1016/j.meegid.2020.104451. [DOI] [PMC free article] [PubMed] [Google Scholar]
  93. Sinha M.; Jagadeesan R.; Kumar N.; Saha S.; Kothandan G.; Kumar D. In-Silico Studies on Myo Inositol-1-Phosphate Synthase of Leishmania Donovani in Search of Anti-Leishmaniasis. J. Biomol. Struct. Dyn. 2020, 40, 3371–3384. 10.1080/07391102.2020.1847194. [DOI] [PubMed] [Google Scholar]
  94. Sur V. P.; Sen M. K.; Komrskova K. In Silico Identification and Validation of Organic Triazole Based Ligands as Potential Inhibitory Drug Compounds of Sars-Cov-2 Main Protease. Molecules 2021, 26 (20), 6199. 10.3390/molecules26206199. [DOI] [PMC free article] [PubMed] [Google Scholar]
  95. Saxena P.; Zangerl-Plessl E.-M.; Linder T.; Windisch A.; Hohaus A.; Timin E.; Hering S.; Stary-Weinzinger A. New Potential Binding Determinant for HERG Channel Inhibitors. Sci. Rep. 2016, 6 (1), 24182. 10.1038/srep24182. [DOI] [PMC free article] [PubMed] [Google Scholar]
  96. Mosquera-Yuqui F.; Lopez-Guerra N.; Moncayo-Palacio E. A. Targeting the 3CLpro and RdRp of SARS-CoV-2 with Phytochemicals from Medicinal Plants of the Andean Region: Molecular Docking and Molecular Dynamics Simulations. J. Biomol. Struct. Dyn. 2020, 40, 2010–2023. 10.1080/07391102.2020.1835716. [DOI] [PMC free article] [PubMed] [Google Scholar]
  97. Menéndez C. A.; Accordino S. R.; Gerbino D. C.; Appignanesi G. A. Hydrogen Bond Dynamic Propensity Studies for Protein Binding and Drug Design. PLoS One 2016, 11 (10), 1–12. [DOI] [PMC free article] [PubMed] [Google Scholar]
  98. Ghallab D. S.; Shawky E.; Metwally A. M.; Celik I.; Ibrahim R. S.; Mohyeldin M. M. Integrated: In Silico - In Vitro Strategy for the Discovery of Potential Xanthine Oxidase Inhibitors from Egyptian Propolis and Their Synergistic Effect with Allopurinol and Febuxostat. RSC Adv. 2022, 12 (5), 2843–2872. 10.1039/D1RA08011C. [DOI] [PMC free article] [PubMed] [Google Scholar]
  99. Santi M. D.; Paulino Zunini M.; Vera B.; Bouzidi C.; Dumontet V.; Abin-Carriquiry A.; Grougnet R.; Ortega M. G. Xanthine Oxidase Inhibitory Activity of Natural and Hemisynthetic Flavonoids from Gardenia Oudiepe (Rubiaceae) in Vitro and Molecular Docking Studies. Eur. J. Med. Chem. 2018, 143, 577–582. 10.1016/j.ejmech.2017.11.071. [DOI] [PubMed] [Google Scholar]
  100. Cao H.; Pauff J. M.; Hille R. X-Ray Crystal Structure of a Xanthine Oxidase Complex with the Flavonoid Inhibitor Quercetin. J. Nat. Prod. 2014, 77 (7), 1693–1699. 10.1021/np500320g. [DOI] [PubMed] [Google Scholar]
  101. Pauff J. M.; Cao H.; Hille R. Substrate Orientation and Catalysis at the Molybdenum Site in Xanthine Oxidase: Crystal Structures in Complex with Xanthine and Lumazine. J. Biol. Chem. 2009, 284 (13), 8760–8767. 10.1074/jbc.M804517200. [DOI] [PMC free article] [PubMed] [Google Scholar]
  102. Lin S.; Zhang G.; Liao Y.; Pan J.; Gong D. Dietary Flavonoids as Xanthine Oxidase Inhibitors: Structure-Affinity and Structure-Activity Relationships. J. Agric. Food Chem. 2015, 63 (35), 7784–7794. 10.1021/acs.jafc.5b03386. [DOI] [PubMed] [Google Scholar]
  103. Hadi S.; Nastiti K. Karamunting Screening as an Antifungal Using The in Silico Method Targeting N-Myristoyltransferase. Sci. Midwifery 2023, 10 (6), 4481–4487. [Google Scholar]
  104. Wang J.; He H.; Zhou Z.; Bai L.; She X.; He L.; He Y.; Tan D. Chemical Constituents And Bioactivities Of Blumea Balsamifera (Sembung): A Systematic Review. Food Sci. Technol. 2023, 43, e132322 10.1590/fst.132322. [DOI] [Google Scholar]
  105. Pratama I. P. A. A. C.; Putra I. M. H.; Pujasari L. W. S.; Dewi K. D. M.; Laksmiani N. P. L. The Potency Of Blumeatin And Luteolin As Caspase-1 İnhibitor By Molecular Docking. Pharm. Rep. 2022, 2 (1), 22. 10.51511/pr.22. [DOI] [Google Scholar]
  106. Xia K.; Qi W. J.; Wu X. Q.; Song Y. Y.; Zhu J. J.; Ai Y.; Cui Z.; Zhang Z. P.; Tang S. A.; Gui Y. T.; Yuan Y.; Wang L.; Zhong H. Synthesis, Structure Revision, and Anti-inflammatory Activity Investigation of Putative Blumeatin. ACS Omega 2023, 8, 14240–14246. 10.1021/acsomega.3c01247. [DOI] [PMC free article] [PubMed] [Google Scholar]

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