Application of Liquid Chromatography in the Analysis of Flavonoid Metabolism in Plant

Abstract

Plants have evolved the capacity to create a wide range of chemicals during the process of their existence. In contrast to specialized metabolites that accumulate in a small number of plant species, flavonoids are broadly distributed across the plant kingdom. Therefore, a detailed analysis of flavonoid metabolism in genomics and metabolomics is an ideal way to investigate how plants have developed their unique metabolic pathways during the process of evolution. Among the analysis methods used for flavonoids, the coupling of liquid chromatography (LC) with ultraviolet (UV) and/or electrospray ionization (ESI) mass spectrometric detection has been demonstrated as a powerful tool for the identification and quantification of phenolics in plant extracts. This chapter mainly introduces of chemistry and metabolism of flavonoids and the application of liquid chromatography in the analysis of plant flavonoids.

1. Introduction

For decades, it has been commonly accepted that plant compounds have a wide range of biological activities. They are secondary metabolites that have considerable pharmacological characteristics and play an important role in improving human health, and flavonoids are one of the substances that have been isolated. Flavonoids, which are responsible for the color and perfume of flowers, have long been known to be synthesized in specific locations, and there are presently around 6000 flavonoids that contribute to the colorful pigments of fruits, herbs, vegetables, and medicinal plants [1]. Flavonoids are hydroxylated phenolic substances known to be formed by plants in response to microbial infection and are a broad set of polyphenolic chemicals with a benzo-pyrone structure synthesized through the phenylpropanoid pathway [2, 3, 4]. Fruits, vegetables, cereals, bark, roots, stems, flowers, tea, and wine all contain it. The chemical properties of flavonoids are determined by their structural class, degree of hydroxylation, various substitutions and conjugations, and degree of polymerization [5].

Flavonoids and other phenylpropanoids are generated from phenylalanine in plants, including the subgroups of flavanones (e.g., flavanone, hesperetin, and naringenin), flavones (e.g., flavone, apigenin, and luteolin), isoflavones (e.g., daidzein, genistein, glycitein). The quantity of oxidation and pattern of substitution of the C ring change within flavonoid classes, whereas individual compounds within a class differ in the pattern of substitution of the A and B rings [6]. Flavonoids are regarded to have health-promoting effects as dietary components due to their strong antioxidant activity in both in vivo and in vitro systems [7, 8]. Flavonoids’ functional hydroxyl groups influence their antioxidant activities by scavenging free radicals and/or chelating metal ions [9, 10]. In addition to antioxidant capabilities, flavonoids have been shown to have antiviral, antibacterial, anti-inflammatory, vasodilatory, anticancer, and anti-ischemic activities. The metabolism of flavonoids is performed by one or more membrane-associated multienzyme complexes rather than free-floating “soluble” enzymes [11]. The primary enzymes involved in flavonoid metabolism are chalcone synthase (a key enzyme in the phenylpropanoid pathway).

Although the separation, identification, and quantification of constituents in complex plant extracts and most likely will be a challenging task, today a multiplicity of different separation techniques, specific stationary phases, and detectors are available, helping to achieve the desired selectivity, sensitivity, and speed for nearly any separation problem. The most prominent and popular technique in this area of research is liquid chromatography [12]. Chromatographic techniques contributed significantly to the area of natural products, especially regarding identification, separation, and characterization of bioactive compounds from plant sources [13]. Flavonoid metabolism is a strong supporter in disease treatment and prevention with chemicals and is an indispensable ingredient in many nutritional, pharmaceutical, and cosmetic applications. The extensive research of flavonoid metabolism in the genome and metabolism is a great technique to investigate how plants’ unique metabolic pathways originated during evolution. The coupling of liquid chromatography (LC) with ultraviolet (UV) and/or electrospray ionization (ESI) mass spectrometric detection is a potent instrument for the identification of phenolics in plant extracts. This chapter focuses on the chemistry and metabolism of flavonoids, as well as the use of liquid chromatography in the study of plant flavonoids.

2. Overview of chemistry and metabolism of flavonoid

Flavonoids are phytonutrients that belong to the polyphenol class. Polyphenols have been employed in Chinese and Ayurvedic medicine for centuries. A novel chemical was extracted from oranges in 1930. It was thought to be a member of a novel class of vitamins at the time and was labeled as vitamin P. Later, it was discovered that this chemical was a flavonoid (rutin), and over 4000 other flavonoids have been found [14].

2.1 Basic chemistry

2.1.1 General characteristics of the C₁₅ unit

Flavonoids exist in the form of aglycones, glycosides, and methylated derivatives. Flavonoids have a diphenyl propane skeleton with 15 carbon atoms in their main nucleus: two six-membered rings coupled with a three-carbon unit that may or may not be part of a third ring. Two benzene rings (A and B in Figure 1) are primarily connected together by a third heterocyclic oxygen-containing pyrene ring (C) [15].

Figure 1.

General structure of flavonoid.

Isoflavones are flavonoids in which the B ring is connected in position 3 of the C ring. Those with the B ring linked in position 4, are referred to as neoflavonoids whereas those with the B ring linked in position 2 can be further classified into many subgroups based on the structural characteristics of the C ring. Flavones, flavonols, flavanones, flavanonols, flavanols or catechins, anthocyanins, and chalcones are the subclasses [1].

2.1.2 Hydroxylation patterns of A-, B-, and C-rings

Positions 3, 5, 7, 2, 3′, 4′, and 5′ are frequently hydroxylated in flavonoids. The most prevalent A-ring hydroxylation pattern is 5,7-hydroxylation; however, a 7-hydroxy ring (also known as a 5-deoxy ring) is seen in isoflavonoid subgroups and several proanthocyanidins. On rare occasions, a 5,7,8 or 5.6.7-hydroxylation pattern is discovered. The B-ring is often 4′-, 3,4′-, or 3′,4,5′-hydroxylation. Rare flavonoids do not have B-ring oxygenation. A 2′-hydroxylation pattern is present in isoflavonoids. In isoflavonoids, the C ring is commonly hydroxylated at the carbon 3 position and occasionally at the carbon 6a position. This six-membered ring can have a carbonyl group, a hydroxyl group, a double bond between positions 2 and 3, or it can be totally unsubstituted, as in unsubstituted flavans. The isoflavonoid pterocarpans have extra rings as a consequence of 2′-hydroxylation of the original B-ring or cyclization of the added prenyl groups (Figure 2) [16].

Figure 2.

Structure backbone of the main flavonoid group.

2.2 Basic substitution

2.2.1 Hydroxylation

There are just a few flavonoid structures with no hydroxyl groups in the A-ring or one hydroxyl group in position 6 [17]. Such unusual structures appear to occur most frequently in the Primulaceae, Rutaceae, and Thymelaeaceae groups. However, the mechanisms of their biochemical synthesis remains unclear. The great majority of flavonoids have a basic 5,7-hydroxylation pattern of the A-ring, which is formed from malonyl-CoA during chalcone synthesis.

The C6-C3 precursor employed by chalcone synthase determines the hydroxylation pattern of the B-ring first. The physiological standard precursor is typically p-coumaroyl-CoA (4-hydroxycinnamoyl-CoA). The resultant basic C15 chalcone intermediate, naringenin chalcone, has a hydroxyl group in position 4′, which is seen in typical flavonoid structures, and all derived flavonoid structures have the hydroxyl group in position 4′. Thus, cinnamate 4-hydroxylase, a cytochrome P450-dependent monooxygenase that catalyzes the production of p-coumaric acid [18, 19], performs a pre-flavonoid step by introducing the hydroxyl group in position 4′.

The majority of flavonoid families have a hydroxyl group in C-ring position 3. (Figure 1). The well-studied flavanone 3-hydroxylase, a 2-oxoglutarate, Fe(II), and ascorbate-dependent dioxygenase [20], introduces the hydroxyl group at the flavanone level. The soluble dioxygenase catalyzes the 3-hydroxylation of the flavanone C-ring to create 3-hydroxyflavanone (flavanol). The same dioxygenase has also been linked to the catalysis of flavone synthases in several plants, with a 2-hydroxylation of the flavanone C-ring proposed as an intermediary step [16].

2.2.2 Methylation

Methylated flavonoids are a form of natural flavonoid derivative with possibly many health advantages, including enhanced bioavailability when compared to flavonoid precursors [21]. According to studies, methylating these flavonoids might boost their promise as pharmacological agents, leading to innovative uses [22]. Flavonoids have been shown to have a wide range of bioactivities, including anticancer, immunomodulation, and antioxidant activities, which can be enhanced to some extent by methylation [21]. Methylation of flavonoids through their free hydroxyl groups or C atoms significantly boosts their metabolic stability and improves membrane transport, resulting in easier absorption and significantly enhanced oral bioavailability [22]. Although cinnamic acid derivatives are methylated at the phenylpropanoid level in certain circumstances, and feruloyl-CoA serves as a poor substrate for chalcone synthase in others, methylation mainly happens at the C15 level. Poulton presents an overview of plant transmethylation and demethylation processes [23]. Grisebach documented the substrate specificities of several O-methyltransferases from parsley and soybean cell cultures, as well as shoots of Chrysosplenium americanum [24], while Heller and Forkmann added further test characteristics [25]. The methyl donor in all of these reactions is S-adenosyl-L-methionine. Ibrahim’s group has now identified five different O-methyltransferases in the flavonol pathway from C. americanum [26].

2.2.3 Glycosylation

The stability of flavonoids under glycosylation reaction circumstances is an important element to consider. For some flavonoids, direct glycosylation might result in the degradation of the changed molecules. It should also be noted that glycosylation is more than just attaching the carbohydrate residue to the flavonoid component of the molecule; it also involves the removal of the protecting groups. Some flavonoids may be partially destroyed as a result of this procedure [27]. It is generally agreed that glycosylation is a late or terminal step in flavonoid glycoside biosynthesis, except for acylation and prenylation reactions. Since the glycosylation step converts the flavonoid into a more water-soluble constituent, a step necessary for the retention of some flavonoids in the vacuole, the site of the glycosylation might be expected to act at the tonoplast boundary during transfer via the cisternae of, or vesicles derived from the endoplasmic reticulum [16]. The process of direct glycosylation for some classes of flavonoids can lead to the destruction of the modified compounds. There are two major types of linkages that form either O-glycosides or C-glycosides. Parsley preparations contain both types. There is a strict specificity for the position of the hydroxyl group, generally at the 3, 5, and 7 positions of the C- and A-rings. Both 3′ and 4’ B-ring glycosides are known. Recently, rarer 2′ and 5′ glycosides of highly methylated flavonol glucosides have been identified in Ibrahim’s laboratory [26].

2.2.4 Acylation

Many flavonoid families include acylated sugars. They exhibit a variety of physicochemical characteristics and biological activity; however, they have limited solubility and stability. To make use of these features, various publications have indicated that enzymatic acylation of these molecules with fatty and aromatic acids by protease and lipase under varied working conditions is a potential strategy. However, it is critical to strike a balance between increasing stability and solubility while maintaining biological activity. In fact, the acylation site (regioselectivity) can significantly alter these features [28]. The acyl groups are often aromatic acids like hydroxycinnamic acids or aliphatic acids like malonic acid. They appear to be position-specific for glucoside. Malonyl glucosides, which are catalyzed by malonyl transferases, are found in isoflavonoids, flavones, flavonols, and potentially anthocyanins. O-Malonyltransferases were isolated from parsley, which included malonated flavones and flavonols. Aromatic acylation, particularly of anthocyanins, has been observed in Silene and Matthiola sp. In both cases, the acyl groups transferred were either 4-coumaroyl or caffeoyl. Acylation has been observed to promote flavonoid absorption into parsley vacuoles; alterations in the molecular symmetry of the malonylglucosides are thought to be responsible for flavonoid vacuolar entrapment inside the vacuole [29].

2.2.5 Prenylation

Prenylflavonoids are useful natural compounds found in a wide range of plants. They frequently have diverse biological features, such as phytoestrogenic, antibacterial, antitumor, and antidiabetic qualities [28]. Prenyl groups are frequently found in phytoalexins and stress-induced isoflavonoids. They are occasionally cyclized. Pterocarpans having a 2′-oxy function and a phenyl group linked to the B-ring were the most active insect feeding deterrents [30]. Elicitor-challenged bean cell cultures include a prenyltransferase in a microsomal fraction that adds a prenyl group at position 10 on the “B-ring” (also known as the D-ring) of 3,9-dihydroxypterocarpan to create phaseollidin, which is then cyclized to phaseollin. Dimethylallylpyrophosphate was the prenyl donor. The identical preparations were capable of introducing prenyl groups into medicarpin and coumestrol, but the products were not recognized. Previously, two distinct flavonoid-specific prenyltransferases that need Mn²⁺ for full activity were discovered in soybean cotyledons and cell suspension cultures [31].

2.2.6 Sulfonation

Nature mostly employs sulfation of endogenous and external substances to minimize possible harm. Sulfonated flavonoids have recently been found in substantial numbers. The majority of them are sulfate esters of common flavones and flavonols. Flavone sulfates are mostly composed of apigenin, luteolin, or its 6- and 8-hydroxy derivatives. Flavonol glycosides were sulfated through the sugar or a separate hydroxyl group. They are found in both dicots and monocots, primarily in herbaceous species or advanced morphological groupings. They are exclusively seen in ferns on rare occasions and have not been identified in bryophytes or gymnosperms [32].

2.3 Stereochemistry

Flavanones have a unique structural property known as chirality that separates them from all other groups of flavonoids (Figure 3). The chemical structure of all flavanones is based on a C6–C3–C6 configuration consisting of two aromatic rings connected by a three-carbon bond [33]. Almost all flavanones have one chiral carbon atom in position 2 (Figure 3). Except for a subgroup of flavanones known as 3-hydroxyflavanones or dihydroflavonols, which have two chiral carbon atoms in positions 2 and 3 (Figure 4). In the C7 position of ring A, certain flavanones include an extra d-configured mono or disaccharide sugar. These flavanone-7-O-glycosides occur as diastereoisomers or epimers with opposing configurations at just one of two or more tetrahedral stereogenic centers in the corresponding chemical entities [34].

Figure 3.

Spatial disposition of the enantiomers of chiral flavanones.

Figure 4.

Chemical structure of the chiral 3-hydroxyflavanones or dihydroflavonols.

Most natural flavonoids now only have one stereoisomer at C-2. The RS nomenclature identifies the R or S that changes at carbon 2 without any change in stereochemistry, depending on the choice of the change adjacent groups should lead to confusion for flavonoid metabolism. It is not sufficient to utilize (+) - or 2,3-cis or -trans alone to define the four potential isoforms of dihydroquercetin or catechin; consequently, consideration should be given to alternate terminology. For mirror pictures, the ent-prefix is utilized. (+)-Catechin (2,3-trans isomer) with 2R, 3S absolute stereochemistry is simply known as catechin, whilst its mirror counterpart (−)-catechin (2,3-trans) with 2R, 3R stereochemistry is simply known as ent-catechin. Similarly, the (−)-epicatechin (2,3-cis) isomer (2R, 3R) and its mirror image (2S,3S) are known as epicatechin and ent-epicatechin [35].

There are other structures designated for hydroxylation patterns and inter-liquid bonding. To minimize ambiguity in the RS system of the configuration of the interflavanoid bond at C-4, Porter and Hemingway provided sugar chemistry terminology, particularly when defining proanthocyanidin isomers. The words are also used to characterize the stereochemistry of the added hydroxyl group at the C-3 position, which results in the 2,3-cis (a-OH) and more prevalent 2,3-trans (B-OH) forms. However, such language does not accurately describe the metabolic route [16].

2.4 Overall pathways metabolism of flavonoid

Flavonoids, which include chalcones, flavones, flavonols, anthocyanins, and proanthocyanidins, are abundant in plants and have been extensively researched using biochemical and molecular biology approaches. Until recently, liverworts and mosses were thought to be the earliest flavonoid-producing plants. Genes encoding enzymes in the phenylpropanoid biosynthetic pathway, including the first two enzymes for flavonoid biosynthesis (chalcone synthase and chalcone isomerase), have not been found in the algal genera Chlamydomonas, Micromonas, Ostreococcus, and Klebsormidium, whereas genes encoding enzymes in the shikimate pathway have been found in algae, liverwort [36].

The overall route to main flavonoid groups via 5,7-hydroxy A-rings. The key intermediates in the production of flavonoids are 4-coumaroyl-coA and 3-malonyl-coA. Synthesis of 4-coumaroyl-CoA and malonyl-CoA naringenin chalcone is synthesized by chalcone synthase, an enzyme involved in the phenylpropanoid pathway. Naringenin chalcone has the ability to spontaneously cyclize to naringenin. Furthermore, naringenin chalcone synthesizes a variety of chemicals such as chalcones, aurones, and biflavonoids (it can synthesize from flavanones). Naringenin has three routes for drug synthesis. The first is the direct synthesis of isoflavonoids, followed by the addition of 3’ OH to produce eriodictyol, followed by the addition of 5’OH to produce 5’OH-erio flavones. Eriodictyol may be combined with 3-OH to get DHQ (dihydroquercetin), then with 5’OH to form DHM (dihydromyricetin) to form flavonols and flavan-3,4-diols (it can synthesize flavan-3-ols, proanthocyanidins, and anthocyanidins). Finally, adding 3-OH to naringenin results in DHK (dihydrokaempferol) (Figure 5) [16].

Figure 5.

Overall pathway to major flavonoid groups with 5,7-hydroxyl a rings (DHK = dihydrokaempferol; DHQ = dihydroquercetin; DHM = dihydromyricetin).

3. Liquid chromatography in the analysis of flavonoid metabolism

To show the chemical variety of flavonoids, chromatographic methods have been utilized to examine their structures. Previously, the major methods used to analyze flavonoids were paper chromatography, thin layer chromatography, column chromatography, and liquid chromatography (LC) [36]. Efficient screening of plant extracts may be accomplished using biological assays as well as chromatographic techniques such as high-performance liquid chromatography (HPLC) in conjunction with different detection modalities [37]. Because it permits systematic profiling of complex plant samples and especially focuses on their identification and consistent assessment of the found compounds, HPLC is a potent tool for the quick investigation of bioactive ingredients. Modern HPLC separation of flavonoids nearly entirely uses reversed-phase liquid chromatography (RP-LC), with significant exceptions being normal phase liquid chromatography (NP-LC) for oligomeric proanthocyanins [38] and the recent rising use of hydrophilic interaction chromatography (HILIC). Other flavonoid separation methods include mixed-mode ion-exchange-reversed phase separation of anthocyanins [39, 40, 41], size exclusions chromatography (SEC) analysis of flavonol glycosides [42], and theaflavins and proanthocyanidins [43]. However, because of the infrequent usage of the later modes, this chapter will concentrate mostly on RP-LC in line with the extent and predominance of this method in flavonoid literature.

RP-LC has proved its applicability for the separation of flavonoids depending on the nature of the aglycone (including the oxidation state, substitution patterns, and stereochemistry), the type and degree of glycosylation, and the nature and degree of acylation. The vast majority of RP-LC separations are accomplished using C18 octadecyl-silica (ODS) phases, however, C₈ [44], C₁₂ [45], phenyl or phenyl-hexyl [46, 47, 48, 49, 50], pentafluorophenyl (PFP) [51, 52, 53] and polar embedded RP phases [54, 55, 56] as well as polymeric RP-LC phases were still widely used [57]. Aqueous/organic phases including methanol, acetonitrile, and less commonly tetrahydrofuran [58], isopropanol [59] or ethanol [57], and acidic modifiers such as acetic acid, formic acid, ammonium acetate, or trifluoroacetic acid (TFA) [59] are typical mobile phases (phosphoric, citric, or perchloric acids have also been used in combination with UV detection, although these are not suited to hyphenation with MS). Highly acidic mobile phases (>4–10% formic acid, 0.1–0.6% TFA) [60, 61, 62, 63] are utilized for anthocyanins to assure the presence of flavylium cationic species in solution and therefore increase chromatographic efficiency. To detect and/or identify flavonoids, a variety of detectors may and have been used in conjunction with HPLC separation. These include electrochemical detection (ED) [64, 65], fluorescence (FL) [66], UV–Vis, diode array [59, 67], NMR [68, 69], and of course MS detectors [70, 71]. The most prevalent currently are diode array and MS detectors, which will be explored briefly in this and the next sections.

3.1 Liquid chromatography (LC) with ultraviolet (UV) detector

Particularly in early flavonoid research, the conjugated aromatic nature of flavonoids proved to be a significant advantage: absorption at relatively long wavelengths increases the selectivity of qualitative and quantitative spectrophotometric methods, and the distinctive spectra of various classes of flavonoids allow differentiation between them. These qualities are similarly helpful when HPLC separation is combined with UV–Vis detection. Flavonoids exhibit two UV–Vis absorption maxima: Band II (Band II), which is attributed to the A-ring, and Band III (Band III), which is attributed to the B-ring (Band 274 I). Due to the fact that it offers more specialized information and since all flavonoids absorb between 240 and 285 nm, the latter of these is more useful. Due to the absence of conjugation between the A and B rings, flavanols, flavanones, dihydroflavonols, and isoflavones only display Band II absorption (269–279 nm). Anthocyanidins may be easily identified by their Band I absorption between 460 and 550 nm in the visible range, in contrast to flavonols and flavones, which exhibit Band I absorption between 300 and 380 nm [72]. Figure 6 provides typical illustrations of the UV–Vis absorbance spectra of the major groups of flavonoids.

Figure 6.

UV–vis absorbance spectra of the principal classes of flavonoids: (a) Luteolin [73]; (b) quercetin [74].

In research by Mohamed A. Farag et al., an integrated approach utilizing HPLC–UV was used for the large-scale and systematic identification of polyphenols in Medicago truncatula root and cell culture. UV spectra (200–600 nm) were recorded for different flavonoid sub-classes including 26 isoflavones (peaks 1–6, 8, 10, 12, 14, 19–29, and 31–35), 4 flavones (peaks 9, 13, 18, and 31), 2 flavanones (peaks 21 and 30), 2 aurones (peaks 7 and 11), and a chalcone (peak 32) with isoflavone representing the major sub-class. For example, isoflavones typically have a maximum absorbance near 255 nm with a second maximum between 300 and 330 nm (peak 34 in Figure 7), whereas aurones have the first maximum near 250 nm and the second peak around 390 nm (peak 7 in Figure 7) [75].

Figure 7.

HPLC–UV at 260 nm (a), 1 and 2 represent UV spectra of peak 7 (hispidol 40-O-glucoside) and peak 34 (afrormosin), respectively [75].

Kim-Ngan Huynh Nguyen et al.’s study quantified seven major compounds, including phenolic acids (chlorogenic acid, caffeic acid, and p-coumaric acid) and flavonoids (rutin, quercitrin, quercetin, and kaempferol) in three aerial parts of Physalis angulata, that is, leaves, calyces, and fruits. Chromatographic separation was carried out on a Kromasil C18 column (150 mm × 4.6 mm i.d., 5 μm) with a gradient elution of 0.1% formic acid in acetonitrile, 0.2% ammonium acetate/0.1% formic acid in water and methanol at a flow rate of 1.0 mL/min; detection was at 250 and 300 nm. The applications of liquid chromatography (LC) with ultraviolet (UV) for the analysis of flavonoid metabolism of plants that show in Table 1 (Figure 8) [81].

Plant	Flavonoid	Detection wavelength	Stationary phases (column dimensions)	Mobile phase	Ref.
Medicago truncatula	isoflavone afrormosin, irilone	260-321 nm, 260-325 nm	C18, 5 μm, 4.6 × 250 mm column	Water (0.1% acetic acid) and acetonitrile using gradient mode	[75]
Brassica rapa L. Ssp. chinensis L. (Hanelt.)	Quercetin, kaempferol, isorhamnetin	204 nm, 254 nm, 352 nm	Alltima HP C18 column	0.5% orthophosphoric acid (v/v) in 30% methanol (v/v) and 0.5% orthophosphoric acid (v/v) in methanol	[76]
Lupinus albus, Lupinus angustifolius	Genistein (isoflavone), 2′-hydroxygenistein glycosides	259 nm	RP C-18 silica gel	A (95% acetonitrile, 4.5% H2O, 0.5% acetic acid; v/v/v)	[77]
Brazilian Vernonieae (Asteraceae)	flavones, flavonols, flavone C-glycosides, flavonol O-glycosides	200–600 nm	Kinetex 1.7 mm XB-C18	Water and acetonitrile, both with formic acid 0.1% (v/v) using gradient mode	[78]
Sutherlandia frutescens (L.)	Four flavonoids	260 nm	C18 column	Water (0.1% acetic acid) and acetonitrile (0.1% acetic; acid) using gradient mode	[79]
Orange juice	Flavanones, flavones, flavonols	280 nm, 265 nm, 265 nm	C18 standard-bore column	Aqueous formic acid (pH 2.4)/and acetonitrile (80:20, v/v)	[80]

Table 1.

Applications of liquid chromatography (LC) with ultraviolet (UV) for the analysis of flavonoid metabolism.

Figure 8.

HPLC chromatogram of the mixed standards solution (a), P. angulata leaves (b). P1–P13: Phenolic acids, including P4: Chlorogenic acid, P6: Caffeic acid, and P8: p-coumaric acid. F1–F9: Flavonoids, including F4: Rutin and F9: Quercetin [81].

3.2 Liquid chromatography (LC) with mass spectrometry (MS)

In contrast to 30 years ago, routine separation and preliminary identification of complex mixtures of flavonoids ranging over many orders of magnitude in concentration are now achievable because of the combination of chromatographic resolution offered by HPLC and structural data offered by MS. Furthermore, during the past 10 years, significant advancements in LC technology have been realized. UHPLC (ultra-high pressure liquid chromatography), alternative stationary phases including monoliths and superficially porous phases, high-temperature HPLC [82, 83], and multidimensional HPLC [84, 85, 86, 87] are a few noteworthy advancements.

Cheminformatics methods combined with LC-MS/MS provide a potent tool for high-throughput surveys of flavonoid variety [88, 89]. Utilizing straightforward solvent combinations and LC columns, glycosylated, acylated, and prenylated flavonoid molecules and their aglycones may be separated. For MS/MS analysis, the isolated molecules are ionized next. In order to analyze flavonoids, LC-tandem mass spectrometry (LC-MS/MS) has emerged as the method of choice. Algae had previously been thought to have no flavonoids. But using an LC-MS/MS technique, flavonoids were identified as intermediates and end products, demonstrating the occurrence of flavonoid production in microalgae [90]. It implies that the undiscovered flavonoids in every plant species can be discovered using cutting-edge metabolomics technology.

With an emphasis on general ionization and fragmentation processes, a brief overview of the underlying knowledge pertinent to the MS detection and MS/MS structural elucidation of flavonoids will be provided in this part. Additionally, specialized research reports provide much more in-depth information on particular classes of flavonoids, including dihydroflavonols [91], isoflavones [92], flavone-di-C-glycosides [93], flavonoid-aglycones [94], flavonoid-O-glycosides [95], and flavonoid glycosides [96]. The discussion that follows will be restricted to the API sources electrospray ionization (ESI), atmospheric pressure chemical ionization (APCI), and atmospheric pressure chemical ionization, which are presently the most pertinent LC-MS ionization sources (APPI). The applications of liquid chromatography (LC) with mass spectrometry (MS) to the analysis of flavonoid metabolism are shown in Table 2.

Plant	Flavonoids	Stationary phase (column dimensions)	Mobile phase(s)	MS analyzer	Ref.
Arabidopsis thaliana	Chrysoeriol (3′ -O-methylluteolin), kaempferol, myricetin, and quercetin	A Superspher 100 RP-18 column	A (95% acetonitrile, 4.5% H2O, 0.5% formic acid, v/v/v) and B (95% H2O, 4.5% acetonitrile, 0.5% formic acid v/v/v)	ESI-QqQ	[97]
Citrus fruits	Naringin, apigenin, eriodictyol, homoeriodictyol, and hesperetin	Waters BEH C18 column, Phenomenex Kinetex C18 column, and Agilent Poroshell 120 EC-C18 column. (2.1 mm × 50 mm, 1.7 μm)	Water and methanol, both with 0.1% formic acid	ESI-QqQ	[98]
Chenopodium hybridum	Rutin, 3-kaempferol	Eclipse XDB-C18 (150 × 4.6 mm, 5 μm	A: 0.1% Acid formic, B: Acetonitrile with 0.1% Acid formic	ESI QqQ-IT	[99]
Orange peel	Kaempferol, neohesperidin, luteolin, homoorientin, tangeretin, diosmetin formononetin quercetin, hesperidin, apigenin, naringenin, naringin, and oleuropein	Mediterranea Sea C18 (150 × 0.46 mm, 3 μm);	A: 0.1% Acid formic, B: Acetonitrile with 0.1% Acid formic	ESI-QqQ	[100]
Vitex negundo var. cannabifolia	Vitexin, hyperoside, luteoloside, liquiritin, and albiflorin	ACQUITY UPLC Cortest C18 (100 × 2.1 mm, 1.6 μm);	A: Acetonitrile, B: 5% MeOH:H2O with 0.1% acid formic	ESI-QqQ	[101]

Table 2.

Applications of liquid chromatography (LC) with mass spectrometry (MS) for the analysis of flavonoid metabolism.

*ESI: electrospray ionization; QqQ: triple quadrupole MS, IT: ion trap MS.

Shoucuang Wang et al. (2017) researched comprehensive profiling of metabolites in citrus fruits. Non-targeted high-performance liquid chromatography with diode array detection and electrospray ionization mass spectrometry (HPLC-DAD-ESI-MS/MS) was used to profile the metabolites in fruit tissues. As a result, 7416 metabolic signals were detected. In addition to those reported metabolites, seven O-glycosylpolymethoxylated flavonoids were newly annotated in the study. To better characterize these flavonoids, the 3′,4′,5,6,7,8-hexamethoxyflavone standard (m70, RT 15.3 min, m/z 403.1389, error − 0.5 ppm) was analyzed first. The precursor ions of the standard compound lost one to four methyl radicals in the MS/MS spectrum to form the base peaks of [M + H - 15]+, [M + H - 30]+, [M + H - 45]+, or [M + H - 60] + (Figure 9A). The characteristic loss of 162 Da was observed in the MS/MS spectra corresponding to the dissociation of a hexose moiety and a series of methyl loss of the diagnostic fragments of 15 and 30 Da (Figure 9B–D) [102].

Figure 9.

Mass spectra and structures of polymethoxylated flavonoids glycosides in citrus. (A) 3, 4, 5, 6, 7,8-hexamethoxyflavone (m070). (B) Dihydroxy-trimethoxyflavone -O-hexoside (m117). (C) Hydroxytetramethoxyflavone-O-hexoside (m119). (D) Monohydroxy-hexamethoxyflavone-O-hexoside (m133). PMFs, DFI, diagnostic fragment ions of polymethoxylated flavonoids [102].

Paola Montoro et al. (2012) researched the metabolic profiles of different extracts (obtained by petals, stamens, and flowers) by LC-ESI-IT MS (liquid chromatography coupled with electrospray mass spectrometry equipped with an ion trap analyzer). MS/MS experiments were diagnostic for the identification of specific fragmentation patterns, that is, sugar loss for flavonoid O-glycosides or the loss of specific esterification units. Interpretation of the ESI-MS/MS experiment obtained by the analysis of Crocus sativus petals extracts allowed us to tentatively identify 31 flavon derivatives in the extracts under investigation, mainly glycosidated and metoxilated derivatives of kaempferol, quercetin, isorhamnetin, and tamaryxetin (Figure 10) [103].

Figure 10.

LC-ESI-MS. TIC and reconstructed ion chromatograms for Crocins qualitative analysis in H2O/EtOH extract of Crocus sativus petals [103].

3.3 High-performance liquid chromatography in chiral flavonoid

Enantiomer separation, resolution, and analysis have traditionally been achieved by the transitory or covalent synthesis of diastereoisomers. Diastereoisomers can be separated on an achiral chromatographic column by differential contact and retention because they have distinct physicochemical characteristics in an achiral environment. On a chemically bonded chiral stationary phase (CSP) with an achiral mobile phase, racemic flavonoid resolution has typically been achieved through chromatographic enantiospecific resolution through transient production of diastereoisomers.

In 1980, one of the earliest publications on flavanone glycoside HPLC separation appeared. Both naringin and narirutin may be acetylated using an equal mixture of pyridine and acetic anhydride and then resolved at low temperatures (between 0 and 5 OC) [104]. Prunus callus (sweet cherries), oranges, and grapefruit’s prunin (naringenin-7-O-glucoside) epimers were initially separated in the middle of the 1980s using benzoylated derivatives [105]. On Cyclobond I columns, the separation of prunin benzoate and naringin benzoate has also been shown. Naringenin derivatization to naringenin tribenzoate and separation on a Chiralcel OD column are also mentioned in the literature. Naringenin’s hydroxyl groups may prevent chiral identification in this stationary phase as the enantiomers could not be resolved [34].

The main advantage of chiral separation methods over achiral methods is a better understanding of the pharmacokinetics of flavanones and the development of effective dosing regimens. In the case of racemic flavanones or stereochemically pure flavanones, this requires knowledge of the in vivo behavior of the enantiomers and epimers. During the drug development process, understanding and comprehending the conformational stability of chiral compounds may have a significant influence on the pharmacological, pharmacokinetic, and pharmacodynamic data. Using stereospecific analytical techniques, racemization and enantiomerization/epimerization may be studied. Rapid interconversion in vivo would eliminate any potential distinctions in the enantiomers’ medicinal or harmful effects, making the synthesis of stereochemically pure enantiomers useless. Chirality must be taken into account from the beginning of the development process for stereochemically pure compounds and racemates [34].

In a study by Gaggeri R et al. (2011), the HPLC enantioselective separation of (R/S)-naringenin (Figure 11), a chiral flavonoid found in several fruits juices and well-known for its beneficial health-related properties, including antioxidant, anti-inflammatory, cancer chemopreventive, immunomodulating and antimicrobial activities, has been performed on both analytical and (semi)-preparative scale using amylose-derived Chiralpak AD chiral stationary phase (CSP). A standard screening protocol for cellulose and amylose-based CSPs was firstly applied to analytical Chiralcel OD-H and Chiralpak AD-H, as well as to Lux Cellulose-1, Lux Cellulose-2, and Lux Amylose-2 in order to identify the best experimental condition for the subsequent scaling-up. Using Chiralpak AD-H and eluting with pure methanol (without acidic or basic additives), relatively short retention times, high enantioselectivity, and good resolution (Rs = 3.48) were observed. Therefore, these experimental conditions were properly scaled up to (semi)-preparative scale using both a prepacked Regispack column and a Chiralpak AD column packed in-house with bulk CSP [106].

Figure 11.

(R/S)-Naringenin.

4. Conclusion

Many beneficial health effects have been attributed to flavonoids, which are popular in the plant. The study of metabolism and bioavailability is very important in defining the pharmacological and toxicological profile of these flavonoid compounds. Due to great structural diversity among flavonoids, these profiles differ greatly from one compound to another, so the most abundant polyphenols in our diet are not necessarily the ones that reach target tissues. Therefore, careful analysis of flavonoids and their metabolites in biological systems is critical. Several hundred papers on the HPLC of flavonoids have been published in the past 20 or so years, yet HPLC methods can detect flavonoids across one, two, or perhaps three subclasses in one run. The improvements in HPLC flavonoid analysis closely resemble and, to a certain extent, build on those in domains like proteomics and metabolomics, which are supported by important breakthroughs.

Acknowledgments

The authors would like to express their hearty gratitude to Can Tho University of Medicine and Pharmacy. The authors also thank all of their colleagues for their excellent assistance.

Conflict of interest

The authors declare no conflict of interest.