Substituent Effects on in Vitro Antioxidizing Properties, Stability, and

Review pubs.acs.org/JAFC

Substituent Effects on in Vitro Antioxidizing Properties, Stability, and Solubility in Flavonoids Merichel Plaza,*,† Tania Pozzo,‡ Jiayin Liu,† Kazi Zubaida Gulshan Ara,‡ Charlotta Turner,† and Eva Nordberg Karlsson‡ †

Department of Chemistry, Centre for Analysis and Synthesis, and ‡Department of Chemistry, Biotechnology, Lund University, P.O. Box 124, SE-221 00 Lund, Sweden ABSTRACT: Antioxidants are widely used by humans, both as dietary supplements and as additives to different types of products. The desired properties of an antioxidant often include a balance between the antioxidizing capacity, stability, and solubility. This review focuses on flavonoids, which are naturally occurring antioxidants, and different common substituent groups on flavonoids and how these affect the properties of the molecules in vitro. Hydroxyl groups on flavonoids are both important for the antioxidizing capacity and key points for further modification resulting in O-methylation, -glycosylation, -sulfation, or -acylation. The effects of O-glycosylation and acylation are discussed as these types of substitutions have been most explored in vitro concerning antioxidizing properties as well as stability and solubility. Possibilities to control the properties by enzymatic acylation and glycosylation are also reviewed, showing that depending on the choice of enzyme and substrate, regioselective results can be obtained, introducing possibilities for more targeted production of antioxidants with predesigned properties. KEYWORDS: antioxidant, solubility, stability, enzymes, flavonoid, acylation, glycosylation, methylation, sulfation

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INTRODUCTION Antioxidants are molecules that diminish oxidative stress and prevent or delay oxidation by scavenging free radicals. On the market, antioxidants are divided into two major groups: functional food ingredients and antioxidants for preservation.1 Use of antioxidants derived from natural resources is for both purposes gaining more and more attention. Polyphenols are secondary metabolites, widespread among plant species, and are the most common antioxidants in the human diet.2−6 In addition, due to their presence in various types of biomass, polyphenols have potential as additives to industrially produced products.7 Polyphenolic antioxidants are composed of at least one aromatic ring with one or more hydroxyl groups as well as other substituents.8 The major type of polyphenols is the flavonoids, composed of a benzene ring (A), condensed with a six-membered pyran ring (C) carrying a phenyl ring (B) in the 2- or 3-position (Figure 1). Flavonoids have high antioxidizing capacity, which has in vitro been shown to be higher than those of vitamins E and C.9,10 Their capacity to act like antioxidants was recognized already in the 1930s,11

when they were called vitamin P (a term that, however, nowadays has been abandoned). Although flavonoids have high antioxidizing capacity in vitro, they may be less efficient in vivo, and more knowledge on the rate and extent of their absorption, metabolism, and tissue or cell distribution, is needed to elucidate their role in disease prevention.12 Reactive oxygen species (ROS), namely, superoxide anion (O2•−), hydroxyl radical (OH•), hydrogen peroxide (H2O2), and hypochlorous acid (HOCl), attack biological macromolecules (e.g., DNA and proteins) under conditions of oxidative stress. ROS are generated as unwanted byproducts of regular oxygen metabolism by all aerobic organisms and can in vivo give rise to a number of chronic degenerative diseases (e.g., arthritis, cancer, cardiovascular diseases, diabetes, inflammatory diseases, ischemia-reperfusion injury, and neurodegenerative diseases).13−17 Flavonoids act as inhibitors of enzymes involved in the generation of ROS (e.g., xanthine oxidase, protein kinases, enzymes activated by calmodulin, cyclooxygenase, lipoxygenase, and NADPH oxidase) and also chelate prooxidant metal ions (e.g., copper and iron), thus preventing ROS formation while their free radical scavenging capability is kept.18,19 A proposed way to combat health risks imposed by ROS is to adopt diets involving consumption of foods rich in antioxidants (precluding progression of chronic diseases, diminishing mortality rates caused by the same),6 which, for example, has been identified as beneficial under conditions of hypertension.20 Metabolism of dietary flavonoids results in the

Figure 1. Classification of flavonoids according to IUPAC.26 Three different structural backbones with ring labeling and atom numbering of the C-ring are shown.

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© 2014 American Chemical Society

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dx.doi.org/10.1021/jf405570u | J. Agric. Food Chem. 2014, 62, 3321−3333

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Table 1. Structures of the Major Dietary Groups of Flavonoids and a Few Examples

formation flavonoid metabolites that may or may not have similar effects in vivo as their precursors. Hence, it makes sense to investigate related compounds (with different substituent patterns) in vitro to shed light on the influence of structural changes introduced during metabolism.21 Antioxidants are also used as additives to different compounds to prolong their life span.22,23 Sectors of industry with interest in antioxidants include the food industry, but also plastics and rubber, gas and fuel, lubricants, adhesives, and cosmetics. Increased interest in obtaining products from renewable resources in these sectors (e.g., oils that are more oxidation prone when derived from renewables) has led to a growing market.24 For industrial use it is important that the antioxidant can be dissolved together with the target compound and also that its action will proceed over a significant time span, keeping sufficient stability. The antioxidizing power of flavonoids is along with other physicochemical characteristics (i.e., stability and solubility) important for function and depends on the substituents that decorate the flavonoid backbone. The focus of this paper is to review the influence of some major substituents occurring naturally, or as a consequence of in vitro modifications, in different subclasses of flavonoids (especially focusing on hydroxyl groups and their modification by glycosylation and acylation). This is followed by a discussion of current knowledge of the influence of these substituent groups in relation to their application, including methodologies to

measure antioxidizing properties, solubility, and stability. Finally, current attempts to modify substituent patterns in vitro by biotechnological methods are reviewed.

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OVERVIEW OF NATURAL FLAVONOIDS AND THEIR SUBSTITUENTS

Flavonoids are classified according to chemical structure, and their functionality depends on this structure including the relative orientation of different substituent groups on the molecule.25 As flavonoids have attracted the attention of researchers in various areas (ranging from chemistry to biology, pharmaceutical sciences, and medicine), they have been named by diverse criteria. The numbers of subclasses vary, depending on the classification system used. On the basis of the backbone structure the IUPAC nomenclature recommends classification of the flavonoids into flavones, isoflavan, and neoflavone (Figure 1).26 However, in the literature flavonoids have been divided into many groups, which are not all correct; thus, to avoid the confusion about nomenclature IUPAC is running a project called “Recommendations on Nomenclature of Flavonoids” (www.iupac.org). The classification that we find most sensible is the one shown in Table 1, which is built on the major dietary components, named as flavones, flavonols, flavanones, isoflavones, flavan-3-ols, and anthocyanidins. Other minor groups of flavonoids include chalcones, 3322



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Table 2. Overview of Generally Occurring Flavonoid Glycosides and Acyls in Nature subclass of flavonoids

position

flavones

5, 6, 7, 2′, 4′

flavonols

3, 7, 5, 4′

flavanones

5, 6, 7, 4′

flavanonols isoflavons flavan-3-ols (catechin) anthocyanidins

3, 7 7 7 2, 3, 4, 5, 6, 7, 3′, 5′

monosaccharide

disaccharide

glucose, galactose, xylose, rhamnose, arabinose, mannose glucose, rhamnose, xylose

gentiobiose, rutinose, cellobiose, diglucose, dirhamnose rutinose,

glucose, rhamnose, galactose, fucose, arabinose glucose, rhamnose, arabinose glucose glucose

rhamnose-glucose rhamnose-galactose not reported not reported

acylated sugar, isobutyric acid malonic acid

glucose, galactose, rhamnose, xylose sophorose, rutinose, sambubiose, robinobiose, gentiobiose

dihydrochalcones, dihydroflavonols, flavan-3,4-diols, coumarins, and aurones.27 The exact structure of flavonoids varies broadly within the different classes as a result of substitutions, and in plants the various substituent groups include, for example, OH groups, Oand C-methyl groups; methylenedioxy groups; O- and Cprenylation; furano, pyrano, and aromatic substitution; esterification, sulfation, and chlorination.28−30 There are also many flavonoids available in nature in oligomeric forms, and one example is the tannins widely found in tea.31 Glycosyl groups (furanose and pyranose groups) are common in flavonoids and are connected via either O- or Cglycosylation. In the C-glycosides (found, e.g., in rice and cereals) the glycosyl group is linked to an aglycone carbon, usually at the C6- or C8-position.32 More commonly, substitutions in flavonoids occur via reactions with the hydroxyl groups of the aglycone backbone, and O-glycosides are examples of this. The wide structural variation among flavonoid glycosides is thus influenced by several factors including linkage, number and nature of the sugars, and the hydroxylation pattern of the aglycone, which in turn determines the position of O-glycosylation. In nature, C-linked glycosylation is mostly found in the flavone group. In general, dicots or higher families have a tendency to accumulate flavone O-glycosides, whereas phylogenetically more primitive families such as ferns or gymnosperms produce more flavonol O-glycosides.30,33 The Oglycosylated flavonoids are due to bioavailability frequently selected for analysis of potential effects on antioxidizing capacity, involving, for example, enzymatic modification trials in vitro (discussed below). A list stating the type and position of the O-linked glycosides and acyl groups of various types of flavonoids is given in Table 2. The substituents are affecting not only the antioxidizing properties but also the stability and solubility of the compounds. In the coming section we start by focusing attention on the antioxidizing capacity, including methods to obtain this on naturally occurring and differently substituted flavonoids.

acyl group 2-methylbutyric acid, 3-hydroxy-3methylglutaric acid, acylated sugar isovaleric, vinylpropionic acid, acylated sugar acylated sugar, p-coumaroyl

p-coumaric acid, caffeic acid, malonic acid, ferulic acid, sinapic acid

Antioxidizing (or reducing) properties can be displayed as either antioxidant activity or capacity. Antioxidant activity is molecule specific and can be described as a “rate constant of a reaction between a specific antioxidant and a specific oxidant”, whereas antioxidant capacity measures “the amount (in moles) of a given free radical scavenged by a defined sample”.35 Measurements of antioxidant capacity can hence, depending on sample composition, yield scavenging ability either of an individual antioxidant or of the combined action of a mixture of antioxidants. Flavonoids are known to be efficient scavengers of free radicals, such as OH•, O2•, and LOO• (lipid peroxide radicals).9 Their potential as antioxidants is predicted by their reducing properties as hydrogen- or electron-donating agents, and evaluation is made using a variety of antioxidant assays, with different strengths and limitations. It is also difficult to compare data from investigations that use different methods, and thus it is recommendable to use a combination of assays. For this purpose, it is advisable to pick methods that are validated, standardized, and widely reported. A number of such methods are summarized in Table 3, along with a brief description of the analysis principle for the respective method. These methods, for example, include oxygen radical absorbance capacity (ORAC), Trolox equivalent antioxidant capacity (TEAC), 2,2-diphenyl-1-picrylhydrazyl (DPPH), ferric reducing antioxidant power (FRAP), cyclic voltammetry total reducing capacity (CV), Folin−Ciocalteu reducing capacity (FC), cupric reducing antioxidant capacity (CUPRAC), and total radicaltrapping antioxidant parameter (TRAP). Antioxidant assays can mainly be divided into one of the two groups: hydrogen atom transfer (HAT)-based assays52 or electron transfer (ET)-based assays (see Table 3). HAT-based assays measure the capacity of an antioxidant to quench free radicals by hydrogen atom donation, and the majority involve a competitive reaction scheme. In these assays, thermal decomposition of azo-compounds generates peroxyl radicals, resulting in competition between the antioxidant and the substrate.40 ET-based assays measure the ability of a potential antioxidant to transfer one electron to reduce any compound (which could be metals, carbonyls, and radicals). In principle, ET-based assays measure the capacity of an antioxidant to reduce an oxidant probe that changes color upon reduction40 according to the following scheme:

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MEASURING THE ANTIOXIDIZING PROPERTIES The capacity of an antioxidant has, according to Biedrzycka and Amarowics,34 been defined by Rice-Evans et al.9 and is determined by (i) its reactivity as a hydrogen- or electrondonating agent (depending on its reduction potential), (ii) the fate of the resulting radical (controlled by the capacity to stabilize and delocalize the unpaired electron), (iii) the reactivity with other antioxidants, and (iv) the transition metal-chelating potential.

probe (oxidant) + antioxidant → reduced probe + oxidized antioxidant 3323


ORAC

oxygen electrode

lag time expressed as Trolox equivalents

fluorometric

net AUCk expressed as Trolox equivalents

36−38

technique

quantification

references

radical

j

TEAC

36, 40, 41

Trolox equivalents

spectrophotometric

ABTS•+ radical cationj is reduced by antioxidants, causing absorbance decrease at 414 or 734 nm

ABTS

•+

c d

42

EC50l, Trolox equivalents

spectrophotometric

DPPH• radicald reduce causing absorbance decrease at 520 nm

DPPH radical

DPPH •

d

36, 43−48

Trolox equivalents

spectrophoto-metric

bis(neocuproine) copper(II) cation is reduced by antioxidants, causing absorbance decrease at 450 nm

36, 49

ferrous ions equivalents

spectrophotometric

ferric 2,4,6-tripyridyl-s-triazine complex is reduced by antioxidants causing absorbance increased at 595 nm

ferric 2,4,6-tripyridyl-s-triazine complex

FRAPf

assays by electron-transfer reaction

bis(neocuproine) copper(II) cation

CUPRAC

e

36, 50

oxidation potential (E1/2), intensity of the anodic current (Ia), area under the anodic wave (S)

electrochemical cell

intensity of anodic current is increased due to oxidation of antioxidant compounds at the surface of the electrode

CVg

36, 51

gallic acid equivalents

spectrophotometric

tungstate−molybdate complexes are reduced by antioxidants, causing absorbance increase at 750 nm

tungstate−molybdate complexes

FCh

ORAC, oxygen radical absorbance capacity. bTRAP, total radical-trappping antioxidant parameter. cTEAC, Trolox equivalent antioxidant capacity. dDPPH, 2,2-diphenyl-1-picrylhydrazyl. eCUPRAC, cupric reducing antioxidant capacity. fFRAP, ferric reducing antioxidant power. gCV, cyclic voltammetry total reducing capacity. hFC, Folin−Ciocalteu reducing capacity. iAAPH, 2,2′-azobis(2amidinopropane) dihydrochloride. jABTS•+, 2,2′-azinobis(3-ethylbenzothiazoline-6-sulfonic acid) radical cation. kAUC, area under the curve that represents the analytical property monitored along time. l EC50, efficient concentration.

36, 39

oxygen consumption

fluorescence decay along time due to oxidation of probe is inhibited by antioxidants

principle of measurement

a

AAPH (peroxyl radical generator)

i

TRAP

AAPH (peroxyl radical generator)

i

b

probe

method

a

assays by hydrogen atom transfer reactions

Table 3. In Vitro Antioxidant Capacity Assays

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pyrogallol group (hydroxyl groups at the 3′-, 4′-, and 5′positions on the B-ring) enhances antioxidant capacity compared to flavonoids with single hydroxyl groups in ring B59 (Figure 3B). Electron delocalization from ring B is promoted by a C-2, C-3 double bond combined with the 4oxo function in the pyran ring (C-ring),58 which also contributes to the antioxidant capacity (Figure 3C). In line with this, measurements using the TEAC assay (Table 3) show that quercetin presents enhanced antioxidant capacity compared to catechin60 (see Table 1 for structures). The 3- and 5OH groups (in the A- and C-rings, respectively) neighboring the 4-oxo function of the C-ring (Figure 3D) also promote high antioxidant capacity.61 The presence of a free 3-OH group (Figure 3E) seems important as further modification (e.g., 3glycosylation) leads to reduction of antioxidizing capacity.55,62 In the A-ring, pairwise hydroxyl substitutions involving position 5 or 8 promote high antioxidant capacity. This is true for both 5,8- and 7,8-hydroxylation (Figure 3F), whereas corresponding groups at the 5,7-positions had little influence. A single 7hydroxyl also had little effect, whereas a single 5-hydroxyl group (next to the 4-oxo function) increased antioxidant capacity.63,64 Following these criteria, a structure−antioxidant capacity relationship of flavonoids (quercetin > quercetin-3-O-rutinoside > kaempferol > luteolin) was proposed by Brown et al.65 Quercetin (Table 1) has all requirements to exercise effective antioxidant function, mainly due to the presence of both a catechol group in the B-ring and a 3-hydroxyl group in the Cring. Several studies66,67 have also reported that flavanones with hydrogenated C-2 and C-3 (Table 1) are in principle inactive, indicating the importance of the double bond between C-2 and C-3 (Figure 3C), whereas luteolin (with the C-2, C-3 double bond and a 3′,4′-dihydroxyl group) has relatively high capacity.66 The antioxidizing capacity of the anthocyanin molecule (with a structurally different C-ring, Table 1) is, however, high and in the same range as that of quercetin. The high capacity of the anthocyanin molecule has been attributed to its electron delocalization ability combined with its ability to form resonance structures upon changes in pH. The completely conjugated structure of anthocyanin allows electron delocalization, resulting in very stable radicals, which from this perspective is favorable.59,68 Moreover, the positively charged oxygen atom in the C-ring makes it a better hydrogen-donating antioxidant compared to oligomeric proanthocyanidins and other flavonoids.69 Clearly, the antioxidant capacity of flavonoids is related to a combination of these chemical and structural elements. Generally, the higher degree of hydroxyl groups present in the flavonoid ring, the stronger the free radical scavenging will be. It is also apparent that hydroxyl groups of flavonoids are frequently sites of further modification, modulating antioxidant capacity by glycosylation, methylation, or, more rarely, acylation or sulfation. Glycosylation Decreases Antioxidant Capacity. Most flavonoids are glycosylated in nature, and the position and nature of the sugar substituents are species specific.70 Glycosylated flavonoids generally show decreased antioxidant capacity compared to the corresponding aglycones,71 but glycosylation also modulates parameters, for example, solubility and stability, as discussed below. Zielinska et al.72 determined antioxidant properties of quercetin and its glucosides from onion by the CV assay (Table 3) and showed that highest antioxidant capacity was obtained using the aglycone. The order of the antioxidant capacity was quercetin > quercetin-3-

After completion (no more change in color), the degree of color change is proportional to the concentration of antioxidant.40

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ANTIOXIDANT CAPACITIES OF DIFFERENTLY SUBSTITUTED FLAVONOIDS Structures of A-, B-, and C-Rings and Their Hydroxylation. The phenolic groups of flavonoids (B-ring, Figures 1 and 2) supply readily available H-atoms ensuring that the

Figure 2. Structures of flavonols and their oxidized forms. R = OH, quercetin; R = H, kaempferol.

radicals produced can be delocalized over the flavonoid structure.53 During the radical scavenging reaction, the flavonoid molecule donates a hydrogen atom to the reacting radical,54 as shown in Figure 2. The flavonoid free radical formed in the reaction is more stable than the ROS that donates the electron due to the delocalization of the electron in the benzene ring (A-ring, Figures 1 and 2). In vitro experiments have shown that substituents on the molecule affect the scavenging capability. Important for high antioxidizing capacity are the number and arrangement of hydroxyl groups on the aromatic backbone (A-, B-, and Crings)55−57 (Figure 3). Hydroxyls on the B-ring are important for the hydrogen-donating capacity. An o-dihydroxy group (catechol group) gives higher stability to the radical structure and takes part in electron delocalization58 (Figure 3A). A

Figure 3. Basic structural features that characterize antioxidant functionality in polyphenols. Areas circled in red show features that are critical for efficient antioxidant function. Letters A, B, and C denote the ring structures of the flavonoid moiety. 3325



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O-glucoside > quercetin-4′-O-glucoside > quercetin-3,4′-Odiglucoside. These results are in agreement with a report by Rice-Evans et al.55 indicating that the antioxidant properties of quercetin glucosides were most affected by glycosylation of the hydroxyl group in the catechol group of the B-ring. Plumb et al.73 have also reported that antioxidant capacities of flavonol glycosides from tea decreased as the number of glycosidic moieties increased. Besides the total number of glycosidic moieties, the type of glycosylation (O- or C-) and the position and structure of the sugar play important roles. CGlycosylation in the A-ring was in this case shown to decrease antioxidant capacity,74 and this effect may be caused by the properties of the sugar molecule. Flavonoid glycosidic moieties occur most frequently as O-glycosides at the 3- or 7-position. The 7-glycoside (in ring A) resulted in a more pronounced decrease of capacity than 3-glycosylation in the C-ring.75 Obviously, the presence of substituents also affects the bioavailability (uptake) of flavonoids by changing the solubility in hydrophilic solvents. Acylation Is Mostly Investigated after in Vitro Modification. A way to improve the hydrophobic nature of flavonoids is, for example, to connect fatty acids to the hydroxyls by esterification. Acylated flavonoids may occur in natural sources (Table 2), but these modifications are not as common as glycosylation and do normally not encompass long hydrophobic molecules. In this field, in vitro acylation of flavonoids by fatty acids has been made and the effect of these substitutions on antioxidant capacity monitored (see also enzymatic modification, below). Hydroxyl groups on the sugar of a flavonoid glycoside (and not the backbone) are common targets for acylation and generally lead to reduced antioxidant capacity, although the results appear to be dependent on the model assay chosen to measure the resulting antioxidizing capacity. Acylation of quercetin-3-rutinoside via glucose with lauric acid (n-dodecanoic acid), more so than palmitic acid (hexadecanoic acid), negatively influenced overall metal chelation when compared to unesterified quercetin-3-O-rutinoside.76 Acylation of the glucoside in quercetin-3-O-glucoside with fatty acids esters by Salem et al.,77 on the other hand, resulted in the enhanced scavenging capacity of the quercetin3-O-glucoside-ester against ABTS radicals but decreased it against DPPH and the superoxide radical. Furthermore, quercetin-3-O-glucoside esters exhibited an antioxidant capacity that depended on the acyl chain length, where the antioxidant capacity decreased with increasing acyl chain length. Other Substituents. Methylation and sulfation of the free hydroxyl groups are examples of other substitutions, and these follow the trends seen for glycosylation and acylation in generally reducing antioxidant capacity. Rimbach et al.78 studied the effect of sulfation and found that genistein-4′sulfate was a less efficient antioxidant than genistein and genistein-4′,7-disulfate even less efficient. This indicates that sulfation masks important hydroxyl groups of the isoflavone molecule and decreases the antioxidant capacity of the resulting compound. O-Methylated quercetin is also an example of a less potent peroxyl radical scavenger than unsubstituted quercetin, and several studies have reported that steric effects perturb the planarity of O-methylation, which may induce the suppression of antioxidant capacity.75,79 Investigations, mainly using assays based on DPPH or ABTS (Table 3), have shown that the Bring is especially sensitive to the position of methoxyl groups. Reduction of the antioxidant capacity was seen for 2′-O-

methyl/4′,6′-hydroxy substitution in the B ring, whereas 2′,6′hydroxy/4′-O-methyl substitution resulted in a lower effect.80 Zima et al.81 also showed that O-dihydroxy substitution of the flavanone B ring resulted in higher capacity than methoxylated compounds or compounds with para-located hydroxyl groups. Accordingly, reduction of antioxidant capacity by methylation depends on a combination of free hydroxyl groups and the position of methoxyl groups in the flavonoid. Antioxidizing Effects Also Depend on the Chosen Model System. It is apparent that the antioxidant capacity of flavonoids has been studied in many different model systems to gain insights on structure−antioxidant capacity relationships and that the choice of assay system (depending on its mechanism) may affect the result obtained. In a recent study performed by Ishimoto et al.,52 antioxidative properties of five typical flavonoids were assessed (kaempferol, quercetin, myricetin, isorhamnetin, and quercetin-3-glucuronide, Table 1) by the ORAC assay (Table 3). Despite previous findings that an increase in the number of hydroxyl groups in the B ring of a flavonol is generally correlated with an increase in antioxidative capacity, it was found that the ORAC level for kaempferol (with one hydroxyl group in the B-ring) was 1.5-fold higher than that of quercetin and myricetin (with two and three hydroxyl groups in their respective B-rings). It was also shown that the ORAC score of quercetin was comparable to the scores of both its methylated metabolite (isorhamnetin) and its 3-O-glucuronide.52 In the same study, the ORAC score of (−)-epigallocatechin gallate was 1.5 times lower than that of its methylated metabolite, (−)-epigallocatechin 3-O-(3″-O-methyl)-gallate. On the other hand, (+)-catechin and (−)-epicatechin both showed remarkable antioxidative capacity (ORAC levels of 9.0 and 10.0 mol Trolox equiv/mol) compared to all 3′- and 4′-Omethyl-(+)-catechins and 3′- and 4′-O-methyl-(−)-epicatechins (ORAC values in the range of 5.7−6.5 mol Trolox equiv/mol). This shows that the scavenging ability depends not only on the structure of the antioxidant (discussed above) but also on the target molecule chosen to monitor oxidation. In conclusion, antioxidant capacities are dependent on many specific structural features, of which one is the number and/or position of free hydroxyl groups of flavonoids. It is, however, also important to note that the choice of model system for measuring antioxidant capacity influences the results. Antioxidant assays differ from each other in terms of reaction mechanisms, oxidant and target/probed species, reaction conditions, and the form in which results are expressed.36 Therefore, it is prudent to use more than one type of antioxidant assay and also consider the relevance of the assay in relation to putative use of the assayed molecule.

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STABILITY AND SOLUBILITY ARE OTHER IMPORTANT PROPERTIES MODIFIED BY FLAVONOID SUBSTITUENTS Most studies involving flavonoids have focused on the antioxidizing capacity. There are, however, other properties of importance, especially for use and selection of antioxidants for applied purposes. Such properties include the stability and solubility of the molecules. With increased industrial interest for use of antioxidants, the choice of antioxidant for a certain application will be dependent on both the possibility to dissolve the antioxidant and the possibility to prolong the shelf life of the antioxidant or of product compounds to which the antioxidant is added. 3326



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Substituents (such as the O-glycosyl groups connected to the hydroxyl groups in the flavonoids) not only regulate the antioxidizing capacity but also influence both the stability and solubility of the molecules. Although less attention has been given to these factors, they are important for successful use. Glycosylation frequently occurs in natural flavonoids and, in addition, both glycosylation and acylation have been done in vitro, to tune naturally occurring antioxidants to more desired properties for applied use. The effect of glycosylation and acylation on both stability and solubility is thus reviewed according to our current knowledge in the field. Effect of Glycosylation on Stability. Relatively little information is available on the effect of glycosylation on the stability of flavonoid molecules. A few examples are however found where glycosylation has been shown to contribute to stability. For example, Buchner et al. measured the changes in concentration of rutin (also called quercetin-3-O-rutinoside or sophorin) and quercetin, in aqueous solution at 100 °C with air perfusion, and the results showed that rutin had higher stability compared to quercetin.82 In another study of metal-catalyzed oxidative degradation, rutin and quercetin were dissolved in phosphate buffer containing Fe2+ and Cu2+. It was also found that rutin had a slower degradation rate by monitoring the concentration changes during the degradation process.83 Other oxidation studies (e.g., oxidation by peroxidase) of rutin and quercetin also reached the same conclusion.84 In the study by Buchner et al.,82 both quercetin and rutin were degraded in aqueous solutions at pH 5 and 8. Again, and at both conditions, the degradation of quercetin seemed to be much faster compared to that of rutin. Rutin is glycosylated at a single point involving a disaccharide linked to the 3-OH on the quercetin C-ring. The stability of flavonoids is, however, also influenced by the number of glycosylated hydroxyl groups. As an example, studies have shown that boiling in water at 100 °C caused a greater decrease in quercetin-4′-glucoside content than in the content of quercetin-3,4′-diglucoside,85,86 making the diglucoside more stable than the monoglucoside. The degradation was successive, and, for example, quercetin diglucosides almost completely broke down to aglycone via the monoglucose derivative by hydrolytic enzymes.87−89 Acylation Can Further Promote Stability. Although the mechanism is not known, the introduction of acyl groups has also been shown to enhance the thermostability of flavonoids. In Ishihara and Nakajima’s study, in vitro monoacylation of quercetin-3-glucoside via a glucoside hydroxyl with nine different aromatic carboxylic acids all improved the thermostability and light-resistivity of the resulting flavonoids.90 It was suggested that the reason for the improved stability by acylation might be that the intra- and intermolecular interactions, between the flavonoid skeleton and the aromatic ring in the acyl moiety, would prevent the degradation of the molecule.90 A similar observation was reported by Fossen et al.91 that complex anthocyanins such as petanin, which contains one aromatic acyl group, showed higher color intensity and stability than cyanidin-3-glucoside at pH 4.0−8.1. In conclusion, the addition of the above substituents increased the stability of flavonoids. The additional sugar or acyl moiety could protect the aglycone from degradation to some extent, although so far there are no studies available to confirm the mechanism behind this. Effect of Glycosylation on Solubility. Solubility is another important parameter that will determine whether it is

possible to utilize an antioxidant as an additive. Solubility in oils can be desired to prevent oxidation,92 and in addition solubility in water has previously been shown to improve the uptake of flavonoids supplied in the diet.93 There is, however, very limited data on direct measurement of solubility of differently substituted flavonoids, like, for example, comparisons of glycosylated and nonglycosylated compounds. Moreover, some solubility data were obtained by comparing extraction94 or chromatography data,95 which makes it difficult to compare the solubility between different studies. Octanol−water distribution constant (log Kow) values are related to water solubility, and such values can be used as index values for comparing the hydrophilicity of the flavonoids.96,97 Frequently, available comparisons, however, report water solubility at a given temperature for different flavonoid compounds. The presence of a sugar moiety usually increases the solubility of the flavonoids in water solutions. For example, quercetin has a low solubility in water at 25 °C (about 0.92 g/ L).98 Glycosylated quercetin99 as well as other glycosylated flavonols94 was reported to have considerably enhanced water solubility compared to the corresponding aglycone, although it is difficult to find exact numbers on the improvement from the literature. In the study of Chen et al.94 it is reported that the higher solubility in water of glycosylated flavonols led to improved extraction yields. There was, however, no quantitative measurement of the solubility of the flavonols in the above studies. The position of the sugar moiety and the number of the bound sugars also play important roles. For example, the solubility of quercetin-3,4′-diglucoside, quercetin-3-glucoside, and quercetin-4′-glucoside in water solution decreases in the same order.96,97 On the other hand, glycosylation will decrease the lipophilicity of the flavonoid aglycones. The review by Chebil et al., for example, shows that the solubility of rutin (quercetin-3-rutinoside) and isoquercitrin (quercetin-3-glucoside) was decreased in acetone and acetonitrile as compared to the solubility of the aglycone quercetin.100 Effect of Acylation on Solubility. If an increased lipophilicity is desired, acylation, particularly with long-chain fatty acids or vinyl esters from C4 to C18 in length, increases the solubility of flavonoids in organic solvent, that is, reflecting the lipophilicity of the flavonoids.76,101 Such reactions have been done in vitro predominantly as enzymatic biotranformations. Another example is the use of aromatic acids such as hydroxycinnamic acids in acylation reactions, which have been shown to decrease the solubility of anthocyanins in cell cultures, which are hydrophilic environments.95 However, acylation with small inorganic acids such as sulfuric acid instead increases the solubility of quercetin in water solution.95 Therefore, it is reasonable to assume that acylation with hydrophilic acids will increase the hydrophilicity of the targeted flavonoids and vice versa. All in all, although there are not so many studies comparing the absolute solubility data, this is very important in future flavonoid research. Existing literature data support the conclusion that the solubility of flavonoids is affected by the hydrophilicity of the additional moiety either through glycosylation or acylation. Adding hydrophilic groups will increase the hydrophilicity of the flavonoid; for example, glycosylation will increase the solubility of flavonoids in aqueous solution, whereas acylation will increase or decrease the solubility of flavonoids in aqueous solution by adding hydrophilic or hydrophobic acyl groups. Therefore, by choosing an appropriate method, it is possible to modify the solubility of 3327



Review

Table 4. Enzymatic Glycosylation of Flavonoids Using Glycoside Hydrolases source

enzyme

donor

acceptor

product

reference

Aspergillus niger

cellulase

4NP-Fuc

catechin

catechin 4-β-D-fucopyranoside

112

Bacillus stearothermophilus

α-D-glucosidase

maltose

catechin

catechin 7-α-D-glucopyranoside catechin 5-α-D-glucopyranoside

112

Bacillus sp.

α-amylase

dextrin

catechin

catechin 7-α-D-maltoside catechin 5-α-D-maltoside

112

Leuconostoc mesenteroides

glucansucrase

sucrose

luteolin

luteolin-3′-α-D-glucopyranoside luteolin-4′-α-D-glucopyranoside

123


glucansucrase

sucrose

quercetin

quercetin-3′-α-D-glucopyranosides quercetin-4′α-D-glucopyranosides

123


glucansucrase

sucrose

myricetin

myricetin-3′-α-D-glucopyranoside myricetin-4′-α- D-glucopyranoside

123

Pencillium decumbens

cellulase

maltose

quercetin

quercetin-3-rutinoside

94

Humicola insolens

synthase Cel7B_E197S

α-glycosyl fluoride

luteolin

luteolin-5-α-D-glycosyl

119

flavonoid in vitro according to the need of the study, which of course, as described above, will also affect the antioxidant properties of the compounds. Enzymes are typically used to make selective modifications on flavonoids, and below some efforts in this field will be shown.

favor solubility of the product in lipids. Most lipases, however, require glycosylated flavonoid acceptor molecules for function, and they selectively acylate hydroxyl groups on glycosides and do not modify free hydroxyls on the flavonoid aglycone. Efforts in this field using lipases and carboxyl esterases (also including the protease subtilisin) have been carefully reviewed by Chebil and co-workers,97 and it was shown that among the enzymes thus far used, it was only lipase PS from Pseudomonas cepacia (now renamed Burkholderia cepacia)90,100 and carboxyl esterases from Streptomyces rochei and Aspergillus niger100,111 that successfully acylated hydroxyls on a nonglycosylated phenolic antioxidant acceptor, which in both reported cases was catechin. A limited number of donor molecules were tried for these enzymes and were for lipase PS restricted to vinyl acetate (an ester of fossil origin), whereas for the carboxyl esterases a short series of esters were used, including ethyl acetate, ethyl propionate, phenyl propionate, and phenyl butyrate.90,100,111 Most commonly, free or immobilized forms of lipase B from Candida antarctica (CalB) have been used in various synthetic approaches for acylation of glycosylated flavonoids, using a range of donor molecules via both direct esterification and transesterification. The reaction conditions are largely influencing the yield, and immobilized CalB clearly showed the best conversion at water activities close to zero. The observed conversion yields vary largely between different studies, using different acyl donors. Different vinyl esters are relatively commonly used for this purpose100 and, for example, the synthetic reaction in the production of 6-O-((+) catechin 7-Oα-D-glucopyranoside) cinnamate from vinylcinnamate reached a yield of 70%.112 Among acyl donors tested in reactions involving CAL-B to obtain products with increased solubility in lipids, high conversion yields have in some cases been reported using fatty acid donors with carbon chains up to C18. For example, 71% yield has been noted in rutin palmitate synthesis using palmitic acid (C16:0) as donor in reactions using the immobilized form of CAL-B (Novozym435).113

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ENZYMATIC METHODS FOR IN VITRO MODIFICATION OF FLAVONOIDS The use of enzymes and microorganisms has by the European Commission been identified as a key enabling technology in replacing nonrenewable chemicals and materials with renewable alternatives,102−104 and for the purpose of flavonoid modification, enzymes classified as hydrolases (EC 3.x.x.x) are biocatalysts of special interest. Hydrolases can, depending on the reaction conditions, catalyze both hydrolysis and synthesis reactions and thus modify the content of glycosyl groups (using glycoside hydrolases (EC 3.2.1.x)105) or acyl groups (using lipases or carboxyl esterases (EC 3.1.1.x)100) in flavonoids in biotransformation reactions. These biotransformations may affect both the antioxidative power and the physicochemical properties of the target molecule, and the benefits of in vitro modification of flavonoid structures using enzymes include production possibilities in amounts that make bioavailability trials as well as proper investigations of physicochemical properties of designed molecules possible. In addition, novel compounds can be created, thus increasing the natural flavonoid diversity in the laboratory.92,106−108 When it comes to practical use of flavonoids in pharmaceuticals, cosmetics, food, or other industrial applications, the relatively low solubility and stability in lipids or water (dependent on the molecule chosen), can also be improved by designed modifications,109,110 avoiding current limitations. Enzymatic Acylation Frequently Requires Glycosylated Acceptor Flavonoids. The enzymes most commonly used for in vitro modification to date are lipases, which have successfully been used to acylate flavonoids, using different esters (transesterification) and organic acids (direct esterification) as acyl donors, that, depending on the selected donor, can 3328



Review

to switch the enzyme mechanism to synthesis only. This methodology is established for oligosaccharide synthesis but has seldom been used for the modification of flavonoids. As the nucleophile of the enzyme is replaced, this approach requires that either fluorinated sugars or external nucleophiles are available in the reaction mixture.125,126 In Yang et al.,119 a highthroughput mass spectrometry-based method was used to assay biocatalysts and substrates for glycosynthesis, demonstrating that finding the right donor and acceptor is crucial for the glycosynthesis reactions. The glycosynthase Cel7B_E197S from Humicola insolens was found to catalyze the reaction, transferring an α-glycosyl fluoride (LacF) into the flavonoid, forming a glycosidic bond with a specificity for position 4′ and 6-linked OH groups on the flavonoid, with a rather high glycosylation yield (75−95%) compared to previous attempts using glycosidases and glycosyltransferases.127 In addition, fair yields (up to 40%) were obtained using a glycosynthase of Thermotoga neapolitana β-glucosidase 3B, using a monoglucosylated quercetin as substrate in reactions obtaining the diglucoside, but required a second mutation in the active site for function.128 In summary, flavonoids are natural antioxidants, and they are getting more attention being molecules from renewable resources that can be utilized as additives in many types of products, ranging from food and pharmaceuticals to cosmetics and eventually larger scale industrial products. To efficiently use and select the antioxidants, a detailed understanding of the interplay between their activity, stability, and solubility is greatly aiding selection of suitable candidates. Substituents connected to their hydroxyl groups definitely have a role in the determination of these properties, and enzymatic routes to modify such substituents and properties are likely to play a role in future modification of natural flavonoids for more selective applications.

From the available literature it can also be seen that the enzymes are to a certain extent regioselective. For example, CAL-B has by different authors been reported to prefer either the primary alcohol (C6-OH)114 or the secondary alcohol at C4-OH115 of the sugar for the acylation reaction. Most papers indicate monoacylation, but for isoquercitrin (quercetin-3glucoside) diacylation on C6-OH and C3-OH has also been shown.100 Glycosides Can Be both Removed and Added by Glycoside Hydrolases. The hydrophilicity of flavonoids can also be varied by changes in the glycosidic group content. Glycoside hydrolases (GH) either catalyze hydrolysis or transglycosylation of glycosidic groups, and are in nature taking part in processes such as (i) degradation and assimilation of exogenous glycosides, (ii) recycling/remodeling of cellular components, or (iii) modification of biological activity of free glycosides.104,116 These activities can of course also be utilized in vitro to deglycosylate flavonoids, for example, using thermostable β-glucosidase,117 as well as to selectively glycosylate flavonoid molecules. The relationship between hydrolytic or transglycosylation activity varies between different enzymes and different GH families and can be modulated if the right conditions are found in the reaction, such as high sugar concentration and controlled water activity. Compared to the use of lipases, rather limited attempts have thus far been made in this field, but interest is increasing, and in addition to the use of natural enzymes, some nucleophile-mutated variants for improved performance have also been constructed and analyzed.118,119 A problem encountered is that glycoside hydrolases require higher water activity (generally aw > 0.6) than lipases,120,121 whereas many flavonoid aglycones have low solubility in water. Successful trials are summarized in Table 4 and include the use of a cellulase from Aspergillus niger and an α-amylase from a Bacillus sp., in reactions with 4-nitrophenyl-α-L-fucoside (4NPFuc) as sugar donor molecules and catechin aglycone as acceptor molecule. Although successful, the yields achieved under the conditions used were not so high, up to 26%.112 Attempts have also been made to optimize the conditions for glycosylation of the flavonoids. Quercetin glycosylation trials at various temperatures, pH values, solvents, and substrates using a Penicillium decumbens cellulase (40−60 °C, pH 6−7, 30−60% ethanol/water v/v) were made with glucose or maltose as the glycosyl donor. In this case, conversions up to 30% were reached.122 Later, the same cellulase was used to assist in extractions of flavonoid compounds in mild solvents such as ethanol/water followed by transglycosylation of flavonoids, converting them into more polar molecules, but the highest transglycosylation yield was only 27% and was achieved using maltose and isorhamnetin.94 To overcome low glycosylation yields, an enzyme with higher natural transglycosylation activity, a glucansucrase from Leuconostoc mesenteroides, was used for glucosylation carried out in aqueous/organic solvents to improve flavonoid solubility. Conversion yields did, however, not improve so much, but varied depending on the flavonoid used, resulting in a conversion to glycosylated forms (Table 4) of luteolin (8%), quercetin (4%), and significantly higher for myricetin (49%).123 In addition, using this enzyme, an α-linked glucoside (quercetin-α-D-glucopyranoside) not occurring in natural isolates was obtained.124 A new approach to glycosylate flavonoids is to use GHs, mutated in the catalytic nucleophile, so-called glycosynthases,

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AUTHOR INFORMATION

Corresponding Author

*(M.P.) E-mail: [email protected], merichelpla@ gmail.com. Phone: +46 765855167. Fax: +46 46 222 82 09. Funding

We acknowledge financial support from the Swedish Research Council Formas for funding the SuReTech research collaboration (229-2009-1527). T.P., K.Z.G.A., and E.N.K. also acknowledge support from the FP7 framework program AMYLOMICS. C.T. acknowledges the Swedish Research Council for funding (2010-333). Notes

The authors declare no competing financial interest.

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