Using Esterase and Laccase Enzymes to Derivatize Bioactive

Plant Phenolics for Altered Chemistry

by

Mohammed Sherif

A thesis submitted in conformity with the requirements

for the degree of Doctor of Philosophy

Department of Cell and Systems Biology

University of Toronto

If someone said

© Copyright by Mohammed Sherif 2015

ii

Using Esterase and Laccase Enzymes to Derivatize Bioactive Plant

Phenolics for Altered Chemistry

Mohammed Sherif

Doctor of Philosophy

Department of Cell and Systems Biology

University of Toronto

2015

ABSTRACT

Plant phenolics have notable antioxidant activity and there is potential to improve their

action by chemical modification. Two enzyme classes carry out reactions that can act on the

hydroxyl moiety of phenolics. Esterase enzymes can be used in non-aqueous solvents to esterify

a long chain acyl group onto the phenolic compound. Laccase enzymes can be used to form

phenoxy radicals that can then couple to form larger molecular weight oligomers. Both

enzymatic modifications may produce a new antioxidant with altered chemistry.

One archaeal esterase (AF1753) from Archaeoglobus fulgidus and one bacterial esterase

(PP3645) from Pseudomonas putida were assayed for activity in organic solvents. Both

enzymes catalyzed hydrolysis of phenyl acetate and vinyl acetate in 98:2 (v/v) (t-amyl

alcohol):buffer; with continued activity up to 96 h of reaction. However, the enzymes were not

able to catalyze transesterification of 4’-hydroxyacetophenone with vinyl acetate in 9:1 (v/v)

cyclohexane:(t-amyl alcohol), which was not explained by enzyme inactivation during

lyophilization. Still, alanine scanning mutagenesis revealed that R37A substitution improved

activity of AF1753 on long-chain p-nitrophenyl (pNP) esters.

A multicopper oxidase (SCO6712) from Streptomyces coelicolor displayed activity on a

variety of phenolics including caffeic acid, ferulic acid, resveratrol, quercetin, morin,

iii

kaempferol and myricetin. Among the products formed by action on flavonols were dimers of

quercetin, morin, and myricetin. Quercetin and myricetin dimers showed longer retention time

on reversed phase chromatography. All three dimers could be detected by 5 min of reaction but

depleted by 3 h and 24 h. The TRAP and FRAP antioxidant activity of the whole reaction

mixture of modified quercetin, morin, and myricetin decreased, as starting phenolic was

depleted over 24 h. Accordingly, mass spectrometry was used to shed light on the molecular

structure of the dimers produced from quercetin and myricetin. In both cases, mass

spectrometric analyses ruled out dimer formation through the A ring of each monomer. For

myricetin, the most likely linkage structure was determined to be between either two B rings or

a B ring with a C ring. These predicted linkage positions are in agreement to those observed for

quercetin dimers previously extracted from natural plant sources.

iv

Acknowledgments

I would like to thank my supervisor Professor Emma Master for giving me the

opportunity to work on this project and providing guidance. I will also thank the members of my

Supervisory Committee, Professor Brad Saville and Professor Dinesh Christendat, for their

advice throughout the project. I thank collaborators who gave assistance on this project

including members of the group Structural Proteomics in Toronto (SPiT) in Toronto and

members from Natural Resources Canada (NRCan) in Sault Ste. Marie. BioZone administration

was helpful in making sure things ran smoothly. Colleagues in the Master lab helped with a lot

of theoretical and technical aspects of the project. I thank my family for their support over the

course of my PhD.

v

Table of Contents

LIST OF TABLES ................................................................................................................... VIII

LIST OF FIGURES .................................................................................................................... IX

LIST OF ABBREVIATIONS .................................................................................................... XI

CHAPTER 1. OVERVIEW .......................................................................................................... 1

CHAPTER 2. LIERATURE SURVEY........................................................................................ 3 2.1. Plant phenolic compounds ................................................................................................... 3

2.1.1. Types and distribution .............................................................................................. 3

2.1.2. Biosynthesis and role in plants ................................................................................. 7

2.1.3. Health benefits of phenolics ................................................................................... 12

2.1.4. Examples of antioxidant activity ............................................................................ 12

2.1.5. Phenolic antioxidant activity for food preservation .............................................. 14 2.1.6. Structure-functional correlations among phenolics with antioxidant properties 15 2.1.7. In vitro measurement of antioxidant activity ......................................................... 17

2.1.8. Solubility considerations for antioxidant activity .................................................. 18 2.2. Derivatization of plant phenolics ....................................................................................... 20

2.2.1. Enzymatic strategies used in plant phenolic derivatization .................................. 20 2.2.2. Increasing hydrophilicity of phenolic compounds ................................................ 21 2.2.3. Increasing lipophilicity of phenolic compounds ................................................... 23

2.3. Esterases/lipases .................................................................................................................. 26 2.3.2. Structural features .................................................................................................. 27

2.3.3. Catalytic mechanism ............................................................................................... 29 2.3.4. Transesterification reactions .................................................................................. 30

2.3.5. Applied Use ............................................................................................................. 33 2.4. Laccases ............................................................................................................................... 34

2.4.2. Structural features .................................................................................................. 35

2.4.3. Catalytic mechanism ............................................................................................... 37 2.4.4. Effect of Redox potential ........................................................................................ 41

2.4.5. Applied use .............................................................................................................. 42 2.5. Research Hypotheses and Objectives................................................................................ 45

CHAPTER 3. CHARACTERIZATION OF SOLVENT-TOLERANT

CARBOXYLESTERASES WITH ARYLESTERASE ACTIVITY ....................................... 46 3.1. Introduction ........................................................................................................................ 47

3.2. Materials and methods ....................................................................................................... 49 3.2.1. Gene cloning and protein purification ................................................................... 49

3.2.2. Hydrolytic activity of esterases AF1753 and PP3645 in t-amyl alcohol/water

(98:2, v/v) ........................................................................................................................... 49 3.2.3. Transesterification activity of esterases AF1753 and PP3645 in t-amyl

alcohol/cyclohexane (1:9, v/v) .......................................................................................... 50 3.2.4. Protein structure modeling and site-directed mutagenesis of esterase AF1753 .. 51

vi

3.2.5. Activity of wild type and mutant AF1753 esterases on varying chain-length pNP

esters .................................................................................................................................. 51 3.3. Results and discussion ........................................................................................................ 52

3.3.1. Hydrolytic activity of esterases AF1753 and PP3645 in t-amyl alcohol/water

(98:2, v/v) ........................................................................................................................... 52 3.3.2. Transesterification activity of esterases AF1753 and PP3645 in t-amyl

alcohol/cyclohexane (1:9, v/v) .......................................................................................... 55 3.3.3. Activity of wild type and mutant AF1753 esterases on varying chain-length pNP

esters .................................................................................................................................. 61 3.4. Conclusions .......................................................................................................................... 66

CHAPTER 4. BIOCHEMICAL STUDIES OF THE MULTICOPPER OXIDASE

(SMALL LACCASE) FROM STREPTOMYCES COELICOLOR USING BIOACTIVE

PHYTOCHEMICALS AND SITE-DIRECTED MUTAGENESIS ........................................ 67 4.1. Introduction ........................................................................................................................ 68

4.2. Materials and methods ....................................................................................................... 69 4.2.1. Gene cloning and protein purification ................................................................... 69

4.2.2. Site-directed mutagenesis ....................................................................................... 70 4.2.3. Copper content analysis of wild-type and mutant SCO6712 laccases .................. 70 4.2.4. Substrate profile of wild-type SCO6712 and the Ser292Ala mutant laccases ...... 70

4.2.5. Kinetics of wild-type and Ser292Ala laccases on select substrates ....................... 72 4.2.6. Docking of 2,6-dimethoxyphenol substrate to wild type and Ser292Ala SCO6712

laccase ............................................................................................................................... 72 4.3. Results and discussion ........................................................................................................ 72

4.3.1. Effect of microaerobic cultivation on copper content and activity of SCO6712

laccase ............................................................................................................................... 72

4.3.2. Substrate profile of wild-type SCO6712 laccase .................................................... 73

4.3.3. Kinetics of wild-type SCO6712 laccase on select substrates ................................. 77 4.3.4. Site-directed mutagenesis ....................................................................................... 79

4.4. Conclusions .......................................................................................................................... 82

CHAPTER 5. CHARACTERIZATION OF PRODUCT FORMATION FROM

ENZYMATICALLY OXIDIZED PLANT PHENOLICS AND ASSAY OF

ANTIOXIDANT ACTIVITY ..................................................................................................... 83 5.1. Introduction ........................................................................................................................ 84 5.2. Materials and methods ....................................................................................................... 85

5.2.1. HPLC-MS analysis of flavonol products after SCO6712 laccase treatment ........ 85 5.2.2. HPLC-MS analysis of flavonol dimer presence over laccase reaction time......... 85

5.2.3. Total radical-trapping antioxidant parameter (TRAP) assay of whole laccase

reaction mixture ................................................................................................................ 85 5.2.4. Ferric reducing antioxidant power (FRAP) assay of whole laccase reaction

mixture .............................................................................................................................. 86

5.2.5. HPLC-MS/MS analysis of quercetin dimer and myricetin dimer......................... 86 5.3. Results and discussion ........................................................................................................ 87

5.3.1. HPLC-MS analysis of flavonol products after SCO6712 laccase treatment ........ 87 5.3.2. Antioxidant assay of whole laccase reaction mixture ......................................... 109 5.3.3. HPLC-MS/MS analysis of quercetin dimer and myricetin dimer....................... 120

vii

5.4. Conclusions ........................................................................................................................ 124

CHAPTER 6. DISCUSSION .................................................................................................... 125

CHAPTER 7. FUTURE RESEARCH ..................................................................................... 137

7.1. Further characterization of esterases for transesterification potential, continuing work

of Chapter 3 .............................................................................................................................. 137 7.2. Further assessment of biochemical potential of laccase SCO6712, continuing work of

Chapter 4 .................................................................................................................................. 138 7.3. Further examination of bioactivity of flavonol dimers, continuing work of Chapter 5

................................................................................................................................................... 139

REFERENCES .......................................................................................................................... 141

APPENDIX 1. SUPPLEMENTAL INFORMATION FOR CHAPTER 3 ......................... 159

APPENDIX 2. SUPPLEMENTAL INFORMATION FOR CHAPTER 4 ......................... 162

APPENDIX 3. SUPPLEMENTAL INFORMATION FOR CHAPTER 5 ......................... 163

APPENDIX 4. TRAP ANTIOXIDANT ASSAY USING LINOLEIC ACID IN PLACE OF

DCFH ........................................................................................................................................ 171

APPENDIX 5. QUANTIFYING SOLUBILITY OF QUERCETIN MONOMER FOR

COMPARISON TO QUERCETIN DIMER ......................................................................... 174

viii

LIST OF TABLES

Table 2.1. Different classes of plant phenolic compounds (Balasundram et al., 2006). .............. 3 Table 2.2. Major food sources of plant phenolic compounds (Manach et al., 2004). .................. 6

Table 2.3. Tree/shrub sources of plant phenolic compounds. ....................................................... 6 Table 2.4. Popular antioxidant activity assays. ........................................................................... 17 Table 4.1. Kinetic parameters of wild type SCO6712 and the Ser292Ala variant enzyme. ....... 78 Table 5.1. HPLC-MS data for quercetin, morin, myricetin, and their dimers produced after

enzymatic reaction. ...................................................................................................................... 88

Table 5.2. HPLC-MS data for representative intermediate molecular weight products present in

late time point enzymatic reactions of quercetin, morin, and myricetin. .................................... 99 Table 5.3. Top 10 products from late time point enzymatic reactions of quercetin, morin, and

myricetin. ................................................................................................................................... 100

Table 5.4. Top 10 products from 24 h reaction of quercetin without and with laccase enzyme.

................................................................................................................................................... 105

Table A1.1. Sequences of primers used for construction of esterase AF1753 point mutants. . 159

Table A2.1. Sequences of primers used for construction of laccase SCO6712 point mutants. 162 Table A3.1. HPLC-MS data for kaempferol and product produced after enzymatic reaction. 163

Table A3.2. Fragment ions observed for quercetin dimer #1 after HPLC-MS/MS. ................. 163 Table A3.3. Fragment ions observed for myricetin dimer #2 after HPLC-MS/MS. ................ 164 Table A3.4. Fragment ions observed for non-enzymatically produced quercetin dimer after

HPLC-MS/MS. .......................................................................................................................... 166 Table A3.5. Fragment ions observed for non-enzymatically produced myricetin dimer after

HPLC-MS/MS. .......................................................................................................................... 167

ix

LIST OF FIGURES

Figure 2.1. Structures of some members of A) hydroxybenzoic acids, B) hydroxycinnamic

acids, and C) stilbenes; and D) basic skeleton of all flavonoids, and specifically flavones and

flavonols. ....................................................................................................................................... 5 Figure 2.2. Key steps in biosynthesis pathways for production of hydroxybenzoic acids and

hydroxycinnamic acids. ............................................................................................................... 10 Figure 2.3. Key steps in biosynthesis pathway for production of stilbenes. .............................. 11 Figure 2.4. Key steps in the biosynthesis pathway for production of flavones and flavonols. .. 11

Figure 2.5. Reactions involved in scavenging of peroxyl radicals by a phenolic antioxidant. ... 13 Figure 2.6. Structures of some classes of flavonoids. ................................................................. 16 Figure 2.7. Possible routes to adding long-chain alkyl groups to phenolic compounds to

increase lipophilicity of the phenolic. .......................................................................................... 24

Figure 2.8. Naturally occurring dimers (n=1), trimers (n=2), and tetramers (n=3) made of

successive epicatechin molecules linked to catechin................................................................... 25

Figure 2.9. Reaction catalyzed by esterases and lipases. ............................................................ 27 Figure 2.10. Prototypical α/β hydrolase fold structure. .............................................................. 28

Figure 2.11. Catalytic mechanism of esterases and lipases. ....................................................... 31 Figure 2.12. Reaction catalyzed by laccases............................................................................... 35 Figure 2.13. Structure of laccase TvL from Trametes versicolor. A) One of the cupredoxin

domains of TvL (the protein has three such domains). ............................................................... 37 Figure 2.14. Proposed catalytic mechanisms of oxidation by laccase enzymes. ........................ 40

Figure 2.15. Structures of closely related phenols. ..................................................................... 41 Figure 3.1. Known sites of action of feruloyl esterases and arylesterases.................................. 48 Figure 3.2. Hydrolysis reactions of A) vinyl acetate and B) phenyl acetate carried out in t-amyl

alcohol/water (98:2, v/v) using esterases AF1753 and PP3645. ................................................. 54

Figure 3.3. Transesterification reaction of vinyl acetate and 4’-hydroxyacetophenone carried out

in different solvent mixtures of t-amyl alcohol/co-solvent (1:9, v/v) using commercial lipase PS

(from Amano). ............................................................................................................................. 57

Figure 3.4. Transesterification reaction of vinyl acetate and 4’-hydroxyacetophenone carried out

in t-amyl alcohol/cyclohexane (1:9, v/v) using recombinant esterase AF1753 and PP3645. ..... 58

Figure 3.5. Hydrolysis reaction of ester substrate in transesterification reaction-mix. .............. 59 Figure 3.6. Activity of wild type and mutants of AF1753 on pNP substrates. ........................... 63

Figure 3.7. Images of predicted protein structure of AF1753. ................................................... 65 Figure 4.1. Substrate selectivity of SCO6712. ........................................................................... 76 Figure 4.2. Chemical structures of natural bioactive phenolic substrates. ................................. 77 Figure 4.3. Ribbon image of 2,6-dimethoxyphenol (2,6-DMP) docked in silico to binding

pocket of SCO6712 A) wild type enzyme and B) Ser292Ala mutant. ........................................ 81

Figure 5.1. HPLC-MS chromatograms and m/z spectra for 20 min reaction samples with laccase

enzyme for A) quercetin, B) morin, and C) myricetin. ............................................................... 91

Figure 5.2. HPLC-MS chromatogram and m/z spectrum for 20 min reaction sample with laccase

enzyme for kaempferol. ............................................................................................................... 91 Figure 5.3. HPLC-MS chromatograms for 5 min reaction sample with laccase enzyme for (A)

quercetin, (B) morin, and (C) myricetin. ..................................................................................... 94 Figure 5.4. Mass spectrum for myricetin dimer of m/z 635 (from peak D3 in chromatogram of

Fig. 5.3. C); refer to Table 5.1) for 5 min reaction sample with laccase enzyme. ..................... 94

x

Figure 5.5. HPLC-MS analysis of flavonol dimers from A) quercetin (dimer #1), B) morin, C)

myricetin (dimer #2), and D) myricetin (dimer #3). .................................................................... 98 Figure 5.6. Proposed reaction scheme for production of some of the quercetin oxidation

degradation products. ................................................................................................................. 104

Figure 5.7. Structures of the isomers quercetin and morin. ...................................................... 108 Figure 5.8. TRAP antioxidant activity of laccase reaction-mixes from A) quercetin, B) morin,

and C) myricetin reactions. ........................................................................................................ 113 Figure 5.9. TRAP antioxidant activity of laccase reaction-mixes from A) quercetin, B) morin,

and C) myricetin reactions. ........................................................................................................ 117

Figure 5.10. FRAP antioxidant activity of laccase reaction-mixes from A) quercetin, B) morin,

and C) myricetin reactions. ........................................................................................................ 119 Figure 5.11. Fragmentation patterns of flavonols in positive ion mode tandem mass

spectrometry. ............................................................................................................................. 121

Figure 5.12. Proposed fragment structures of quercetin dimer #1 (refer Table 5.1) after tandem

mass spectrometry. .................................................................................................................... 122

Figure 5.13. Proposed fragment structures of myricetin dimer #2 (refer Table 5.1) after tandem

mass spectrometry. .................................................................................................................... 123

Figure 6.1. Two possible mechanisms of quercetin dimer formation. ..................................... 130 Figure 6.2. Mechanism of antioxidant antagonism proposed by Peyrat-Maillard et al. (2003).

................................................................................................................................................... 133

Figure 6.3. Mechanism of antioxidant antagonism that involves reaction of quercetin radicals

with laccase-generated quercetin products. ............................................................................... 135

Figure A1.1. Protein purification of AF1753 mutants. ............................................................. 161

Figure A3.2. Remaining phenolic monomer in laccase reactions, as measured by UV-Vis

spectrophotometry and by intensity of HPLC-MS peak............................................................ 170

Figure A5.1. The maximum amount of quercetin that can be dissolved in A) 50 mM sodium

phosphate buffer (pH 7.4) with 0.1 M NaCl and B) 1-octanol. ................................................. 177

xi

LIST OF ABBREVIATIONS

2,3-DHB - 2,3-dihydroxybenzoic acid

2,6-DMP - 2,6-dimethoxyphenol

AAPH - 2,2′-azobis(2-methylpropionamidine)

ABTS - 2,2′-azino-bis(3-ethylbenzothiazoline-6-sulfonic acid)

Ala - alanine

AMVN - 2,2’-azobis (2,4-dimethylvaleronitrile)

Arg - arginine

DCFH - 2′,7′-dichlorofluorescin

DCFH-DA - 2′,7′-dichlorofluorescin diacetate

DHA docosahexaenoic acid

DMSO - dimethylsulfoxide

DPPH - 2,2-diphenyl-1-picrylhydrazyl

EDTA - ethylenediaminetetraacetic acid

EPA - eicosapentaenoic acid

ET - electron transfer

FRAP - ferric reducing antioxidant power

Glu - glutamate

HAT - hydrogen atom transfer

HPLC-MS - high performance liquid chromatography-mass spectrometry

LDL - low density lipoprotein

L-DOPA - 3,4-dihydroxy-L-phenylalanine

MCO - multicopper oxidase

N-HPI - N-hydroxyphthalimide

NMR - nuclear magnetic resonance

pNP - p-nitrophenol

ROS - reactive oxygen species

Ser - serine

TPTZ - 2,4,6-tris(2-pyridyl)-s-triazine

TRAP - total radical-trapping antioxidant parameter

1

CHAPTER 1. OVERVIEW

Plant materials are a rich source of bioactive phenolic compounds. Phenolics are a

common constituent of the human diet via fruits, vegetables, and beverages (Balasundram et al.,

2006) but they can also be derived from forest sources such as with the monolignols, stilbenes,

lignans, and certain types of flavonoids (Stevanovic et al., 2009). Phenolic compounds have

well known antioxidant activities that can find use in prevention of diseases affecting human

health, and in food preservation against oxidative decay (Balasundram et al., 2006).

There is potential to increase the protective effect of these compounds towards lipid

targets by increasing their hydrophobicity through the addition of alkyl groups. Hydrophobicity

of the phenolics might also be increased by increasing their molecular weight through

oligomerization. Such modifications may increase the miscibility of the bioactive compound in

emulsified food oils, imparting a preservative effect (Frankel et al., 1994); or in low density

lipoproteins (LDL), thereby reducing the frequency of health problems such as atherosclerosis

(Vafiadi et al., 2008).

The overall objective of this research project was to alter the chemistry of phenolic

compounds by enzymatic modification (for example to increase hydrophobicity of the

phenolics), while maintaining antioxidant activity of the phenolic. Towards this goal two

enzyme types were investigated for their potential to modify phenolics. The first enzyme class,

esterases, were used to try to esterify alkyl chains onto phenolic compounds and the second

enzyme class, laccase (a type of multicopper oxidase), was used to oxidatively dimerize

phenolic compounds via radical coupling reactions. The esterase reaction must be performed in

an organic solvent to avoid hydrolytic breakdown of the desired ester product. By contrast,

laccase reactions can be carried out in aqueous conditions but because of radical delocalization

it is difficult to know in advance which products will be formed from the wide array of potential

products. The feasibility of esterase-mediated and laccase-mediated modification of bioactive

phenolics was investigated, following the literature survey (Chapter 2), in subsequent chapters.

First, the chosen esterases (one bacterial (Pseudomonas putida) and one archaeal

(Archaeoglobus fulgidus)) were assessed for their activity in predominantly organic solvent

media. The enzymes were hydrolytically active in the organic solvent-water mixture but did not

catalyze transesterification reactions. Therefore, attention was focused on the second enzymatic

approach, i.e. laccase oxidation followed by oxidative coupling of phenolics to produce

2

increased molecular weight products as a means of modifying phenolic chemistry. In this case,

initial work focused on evaluating determinants of activity of a bacterial laccase from

Streptomyces coelicolor on a range of phenolic compounds from diverse classes, including

hydroxycinnamic acids, stilbenes, flavonols, and flavones. As laccase activity was observable

on the well-known antioxidant flavonols (among other compounds), the products from these

reactions were analyzed. Presence of dimer products from the flavonols quercetin, morin, and

myricetin was examined using HPLC-MS. Tandem mass spectrometry was used to gain initial

information about the structure of the dimers of quercetin and myricetin. The change in

antioxidant activity resulting from laccase action was assayed using the total radical-trapping

antioxidant parameter (TRAP) assay and the ferric reducing antioxidant power (FRAP) assay on

whole laccase reaction mixtures.

Summary of scholarly contributions

Peer reviewed publications:

Sherif, M., Waung, D., Korbeci, B., Mavisakalyan, V., Flick, R., Brown, G., Abou-Zaid,

M., Yakunin, A. F., & Master, E. R. (2013). Biochemical studies of the multicopper oxidase

(small laccase) from Streptomyces coelicolor using bioactive phytochemicals and site-directed

mutagenesis. Microbial Biotechnology, 6(5), 588-597.

Manuscripts in preparation:

Sherif, M., Wang, L., Tchigvintsev, A., Brown, G., Mavisakalyan, V., Tillier, E. R. M,

Savchenko, A. V, Master, E. R, & Yakunin, A. F. Solvent-tolerant and thermophilic

carboxylesterase with arylesterase activity from Archaeoglobus fulgidus.

Sherif, M., Qazi, S., Abou-Zaid, M., & Master, E. R. Identification of products and

antioxidant activity of reaction mixtures from treatment of four flavonols with a multicopper

oxidase SLAC (small laccase) from Streptomyces coelicolor.

Qazi, S., Sherif, M., Master, E. R, & Abou-Zaid, M. Tandem mass spectrometric and

NMR structural characterization of quercetin dimer produced by multicopper oxidase treatment

of quercetin.

3

CHAPTER 2. LIERATURE SURVEY

2.1. Plant phenolic compounds

2.1.1. Types and distribution

Plant phenolic compounds constitute secondary metabolites and are among the most

prevalent phytochemicals, appearing in both food and non-food sources (reviewed in

Balasundram et al., 2006; Manach et al., 2004). A structural categorization of plant phenolics

leads to the following classes based on the configuration of the carbon skeleton (Table 2.1)

(carbon skeleton in brackets): simple phenolics (C6), hydroxybenzoic acids (C6-C1),

phenylacetic acids (C6-C2), hydroxycinnamic acids (C6-C3), quinones (diverse carbon skeleton),

xanthones (C6-C1-C6), stilbenes (C6-C2-C6), flavonoids (C6-C3-C6), lignans ((C6-C3)2),

biflavonoids ((C6-C3-C6)2), lignins ((C6-C3)n), and tannins (diverse carbon skeleton)

(Balasundram et al., 2006). The flavonoids are further subdivided into the flavones, isoflavones,

flavonols, flavanones, anthocyanidins, and flavanols (Manach et al., 2004). Of the plant

phenolics, the most structurally diverse group are the flavonoids (Balasundram et al., 2006) and

of the flavonoids, the flavonols are the most common in foods (Manach et al., 2004). In many

cases, representatives of these classes of plant phenolics can be found conjugated to monomeric

and oligomeric sugars.

Table 2.1. Different classes of plant phenolic compounds (Balasundram et al., 2006).

Phenolic class Carbon skeleton

Simple phenolics C6

Hydroxybenzoic acids C6-C1

Phenylacetic acids C6-C2

Hydroxycinnamic acids C6-C3

Quinones Diverse

Xanthones C6-C1-C6

Stilbenes C6-C2-C6

Flavonoids C6-C3-C6

Lignans (C6-C3)2

Biflavonoids (C6-C3-C6)2

Lignins (C6-C3)n

Tannins Diverse

4

The higher molecular weight classes mentioned above are oligomeric/polymeric forms

of the lower molecular weight classes. Lignans are dimerized forms, while lignin is a large

complex polymer, of the alcohols of hydroxycinnamic acids. The stilbenes are known to exist in

oligomeric forms of the simplest structural units (C6-C2-C6) of their class (Quideau et al., 2011).

As the name suggests, biflavonoids are dimeric versions of flavonoids. The tannins can be

divided into the condensed tannins (carbon skeleton of (C6-C3-C6)n) and hydrolyzable tannins.

Condensed tannins are oligomeric forms of flavanols while hydrolyzable tannins are composed

of monomeric and polymeric hydroxybenzoic acid units esterified onto sugars (Quideau et al.,

2011). The remainder of this section (Section 2.1.1) and the subsequent section (Section 2.1.2)

will focus on the distribution (dietary and non-dietary), biosynthesis, and role in plants of

hydroxybenzoic acids, hydroxycinnamic acids, stilbenes, and two of the flavonoids (flavones

and flavonols) (Fig. 2.1). These classes of phenolics were chosen because they contain among

the most well-studied and most effective antioxidant compounds.

Hydroxybenzoic acids are not widely found in plant material consumed by people. The

few major sources in the human diet include in certain red fruits, black radish, onion, and tea

(Table 2.2) (reviewed in Manach et al., 2004). One of the more studied of the hydroxybenzoic

acids, gallic acid, can be found in esterified form in the bark of Quercus stenophylla (Nishimura

et al., 1984), in the flowers of Tamarix nilotica (reviewed in Van Sumere, 1989), in maple

species (for example as a glycoside conjugate in leaves of Acer rubrum (Abou-Zaid &

Nozzolillo, 1999) and as a methyl ester in leaves of Acer rubrum, Acer saccharinum, and Acer

saccharum (Abou-Zaid et al., 2009)), (Table 2.3) and various other plant species (for an

extensive list of plants containing gallic acid and other phenolics see Harborne et al., 1990).

More recently, it was isolated from aerial plant parts of Pelargonium reniforme (Latté et al.,

2008). Aside from being obtained from plant material after comparatively gentle solvent

extraction, hydroxybenzoic acids, such as vanillic acid and syringic acid, can also be obtained

upon hydrolytic treatment of lignocellulosic materials, due to oxidation and breakdown of the

lignin polymer (reviewed in Garrote et al., 2004).

5

Figure 2.1. Structures of some members of A) hydroxybenzoic acids, B) hydroxycinnamic

acids, and C) stilbenes; and D) basic skeleton of all flavonoids, and specifically flavones and

flavonols.

OH

OH

OHO

OH

OH

OH

OHO

OH

OHO

O OH

OH

O OH

OH

OH H3CO

O OH

OH

OCH3

O OH

OH

OCH3

OH

OCH3

OHO

H3CO

OH

OH

OH

OH

OCH3

OCH3

OH

OH

O

O

O

O

O

OH

gallic acid protocatechuic acid salicylic acid

A)

B)

syringic acid

p-coumaric acid caffeic acid ferulic acid sinapic acid

C)

resveratrol pterostilbene pinosylvin

D)

flavonoids flavones flavonols

6

Table 2.2. Major food sources of plant phenolic compounds (Manach et al., 2004).

Phenolic class Major food sources

Hydroxybenzoic acids red fruits

black radish

onion

tea

Hydroxycinnamic acids blueberry

kiwi

plum

cherry

apple

Stilbenes grape

Flavones parsley

celery

Flavonols onion

leek

broccoli

blueberry

Table 2.3. Tree/shrub sources of plant phenolic compounds.

Compound Tree species Reference

Hydroxybenzoic acids Quercus stenophylla

Tamarix nilotica

Pelargonium reniforme

Acer spp.

Nishimura et al. (1984)

Van Sumere (1989)

Latté et al. (2008)

Abou-Zaid et al. (2009)

Hydroxycinnamic acids Tsuga heterophylla

Catalpa ovata

Harborne (1990)

Stilbenes Veratrum formosanum

Picea abies

Pinus sibirica

Stevanovic et al. (2009)

Flavones and flavonols Eucalyptus spp.

Crataegus sp.

Pinus spp.

Stevanovic et al. (2009)

Abou-Zaid & Nozolillo (1991)

Hydroxycinnamic acids are mostly found in conjugated forms and the four most

common compounds are conjugated forms of p-coumaric, caffeic, ferulic, and sinapic acids

(Manach et al., 2004). As opposed to the hydroxybenzoic acids, the hydroxycinnamic acids can

be found in a variety of food sources, and highest amounts have been found in blueberries,

kiwis, plums, cherries, and apples (Table 2.2) (Manach et al., 2004). Similar to the

hydroxybenzoic acids, the hydroxycinnamic acids can also be obtained upon hydrolytic

treatment of lignocellulosic materials (Garrote et al., 2004). In addition to making up the lignin

7

polymer (in the alcohol form), the hydroxycinnamic acid ferulic acid (and its dimers) can be

found esterified onto hemicelluloses (Manach et al., 2004).

Stilbenes are not abundant in the human diet except from grapes and their juices (Table

2.2) (Manach et al., 2004). The most widely studied stilbene is resveratrol. Resveratrol and its

glucoside have been found in Veratrum formosanum and in the bark of Picea abies,

respectively, and stilbenes can notably be found from the knotwood extracts of pines (Table

2.3) (reviewed in Stevanovic et al., 2009).

Flavonoids are the most structurally diverse phenolic compound in plants. Among the

most well-known of the flavonoids is the flavonol quercetin because of its very strong

antioxidant activity. Flavones are chiefly found in parsley and celery in the human diet (Table

2.2) (Manach et al., 2004). On the other hand, flavonols are more widely prevalent and can be

found in onions, leeks, broccoli, blueberries, and other food sources (Table 2.2) (Manach et al.,

2004). Additionally, flavonols can be found in leaves of forest trees including birches and

eucalyptus (Stevanovic et al., 2009). Another source of flavonols (and flavones) is in trees of the

family Crataegus, with 13 flavonols and 20 flavones previously identified (Stevanovic et al.,

2009). Furthermore, flavonol glycosides were observed from needles of pine trees such as Pinus

banksiana (Table 2.3) (Abou-Zaid & Nozolillo, 1991).

2.1.2. Biosynthesis and role in plants

The biosynthesis of plant phenolic compounds can be traced back to the shikimate

pathway and the polyketide pathways with the polyketide pathway providing precursors for

production of simple phenolics, while the shikimate pathway provides precursors for the other

phenolic types (Harborne 1989). Starting from shikimate, phenylalanine is produced by the

shikimate pathway. Phenylalanine is then deaminated to produce cinnamic acid, which is then

hydroxylated to produce p-coumaric acid. As such, cinnamic acid is the first precursor to

production of the other plant phenolics (Fig. 2.2) (Harborne, 1989; Dewick, 1995).

Hydroxybenzoic acids are thought to be produced from cinnamic acids by removal of an acetate

unit (Fig. 2.2) (Gross, 1992). However, based on tracer experiments with radiolabelled carbon,

it has also been proposed that gallic acid biosynthesis could proceed via direct dehydrogenation

of shikimic acid without going through a pathway involving cinnamic acid production (Gross,

1992). The hydroxycinnamic acids are formed via aromatic substitution of cinnamic acid by

8

undergoing sequential hydroxylations and methylations (Harborne, 1989; Dewick, 1995).

Stilbene biosynthesis occurs by reaction of three malonyl-CoA molecules with p-coumaroyl-

CoA, followed by a decarboxylation accompanied by cyclization (Fig. 2.3) (Dewick, 1995).

Similar to the stilbenes, for the flavonoids in general, their biosynthesis starts by reaction of

three malonyl-CoA molecules with p-coumaroyl-CoA followed by a cyclization to produce the

flavanones, resulting in the basic carbon skeleton of all flavonoids (Fig. 2.4) (Heller &

Forkmann, 1994; Dewick, 1995). Flavones derive from flavanones by formation of a double

bond between C-2 and C-3 while flavonols are formed by first hydroxylating position 3 of

flavanones followed by formation of a double bond between C-2 and C-3 (Heller & Forkmann,

1994; Dao et al., 2011).

Plant phenolic compounds have been postulated to have a diverse array of functions,

some representative examples being: the role of benzoic acids in photosynthesis (Van Sumere,

1989); the role of ferulic acid in regulating germination of barley seeds (Van Sumere, 1989); the

role of stilbenes as antimicrobial compounds (Gorham, 1989); the role of flavones and/or

flavonols in 1) protection from UV light, insects, and microorganisms, 2) hormonal control, 3)

enzyme inhibition, and 4) attracting pollinators (Markham, 1989).

9

O O-

OH

OH

O-

O

OH

O OH

OH OH

OH

OHO

OH

PO4

CH2 COO-

OH

OHO4P

O

OH

OH

OH

O

OO

-

OH

OHO

O O-

OH

OH

O-

O

O4P

OH

O

O-

O

O4P

CH2

O

O-

OH

O

O-

O

CH2

O

O-

O

OO

-

OH

O-

ONH2

OO

-

OH

O-

ONH2

O O-

OH

OHO4P

OH

O

O

OH

phenylalanine

shikimate

cinnamic acid gallic acid p-coumaric acid

+

PEP

E4P

DAHPS

DAHP

DHQS

3-dehydroquinate

DHQD

3-dehydroshikimate

SDH SK

shikimate 3-phosphate

EPSPS

EPSP

chorismate

CS CM

prephenate

PAT

arogenate

ADT

PAL C4H

10

Figure 2.2. Key steps in biosynthesis pathways for production of hydroxybenzoic acids and

hydroxycinnamic acids. The shikimate pathway produces chorismate which goes on to produce

phenylalanine. Phenylalanine is metabolized to yield hydroxycinnamic acids and

hydroxybenzoic acids. Hydroxybenzoic acids can also be produced from shikimate pathway

intermediates without going through phenylalanine production. Abbreviations for intermediates:

PEP, phosphoenol pyruvate; E4P, erythrose-4-phosphate; DAHP, 3-deoxy-D-arabino-

heptulosonate-7-phosphate; EPSP, 5-enolpyruvylshikimate-3-phosphate. Abbreviations for

enzymes above arrows: DAHPS, 3-deoxy-D-arabino-heptulosonate-7-phosphate synthase;

DHQS, 3-dehydroquinate synthase; DHQD, 3-dehydroquinate dehydratase; SDH, shikimate

dehydrogenase; SK, shikimate kinase; EPSPS, 5-enolpyruvylshikimate-3-phosphate synthase;

CS, chorismate synthase; CM, chorismate mutase; PAT, prephenate aminotransferase; ADT,

arogenate dehydratase; PAL, phenylalanine ammonia-lyase; C4H, cinnamic acid 4-hydroxylase.

11

Figure 2.3. Key steps in biosynthesis pathway for production of stilbenes. p-coumaroyl-CoA

comes from the phenylpropanoid biosynthetic pathway. STS indicates stilbene synthase enzyme.

Figure 2.4. Key steps in the biosynthesis pathway for production of flavones and flavonols. p-

coumaroyl-CoA comes from the phenylpropanoid biosynthetic pathway. Enzyme abbreviations:

CHS, chalcone synthase; CHI, chalcone isomerase; FNS, flavone synthase; F3H, flavanone 3-

hydroxylase; FLS, flavonol synthase.

O SCoA

OH

OH

OSCoA

O

O

OH

OO

AoCSOC

OH

OHOH

malonyl-CoA

stilbene (e.g.

resveratrol

CO2

+ 3x

p-coumaroyl-

CoA

STS STS

O SCoA

OH

OH

OSCoA

O

OH

O

OH

OHOH

OH

O

O

OHOH

OH

O

O

OHOH

OH

O

O

OHOH

OH

OH

O

O

OHOH

OH

p-coumaroyl-

CoA

malonyl-CoA

flavanone (e.g.

naringenin)

flavonol (e.g.

kaempferol) flavone (e.g.

apigenin)

+ 3x CHS CHI

FNS

F3H

FLS

dihydrokaempferol

12

2.1.3. Health benefits of phenolics

Plant-derived phenolic compounds are implicated in a wide array of health benefits. This

includes benefits against cardiovascular disease, neurodegenerative disease, cancer, and diabetes

(reviewed in Scalbert et al., 2005). However, in vitro findings can not always be translated into

similar in vivo effects and tests from different labs do not always give the same results (Scalbert

et al., 2005). Atherosclerosis has been observed to be inhibited by consumption of food

phenolics and, based on animal studies, it is thought that the phenolics mediate their effects by

reducing oxidation and uptake of low density lipoprotein (LDL) by macrophages (Kaplan et al.,

2001; Miura et al., 2001). However, human studies have shown mixed results, with some

studies showing that consumption of tea protects against ex vivo oxidation of LDL (Ishikawa et

al., 1997) while other studies showed no such benefit of tea consumption (Van het Hof et al.,

1997). Animal models have also shown anticarcinogenic effects of food phenolics but the doses

used in such experiments are usually much larger than typical consumption levels in the human

diet, making it difficult to correlate epidemiological results to these animal models (Scalbert et

al., 2005). The importance of dosage is further highlighted by the fact that low doses (less than

10 µM) of epigallocatechin gallate was found to be neuroprotective in cell culture models using

the neurotoxin 6-hydroxydopamine, while higher doses of epigallocatechin gallate had cytotoxic

effects (Levites et al., 2002). Use of plants by indigenous peoples for treatment of diabetes has

been documented and animal model studies have also shown antidiabetic effects of plant

extracts containing phenolics (Scalbert et al., 2005; Giordani et al., 2015). It is thought that one

of the mechanisms by which the plant compounds reduce diabetes is through inhibition of α-

glucosidase enzymes, which normally break down carbohydrates so that the sugars can be

absorbed in the gut (Giordani et al., 2015).

2.1.4. Examples of antioxidant activity

Plant phenolics are well-known for their antioxidant activity, which may, in some cases,

partially mediate their other health effects (reviewed in Scalbert et al., 2005). The antioxidant

activity of plant phenolics is due to reaction with free radicals, but may also involve inhibition

of enzymes and chelation of metal ions (Huang et al., 2005). In the case of reacting with free

13

radicals, the phenolic antioxidant sacrificially becomes oxidized to a relatively stable radical.

Free radical scavenging may occur by the phenolic transferring a hydrogen atom (hydrogen

atom transfer (HAT)) or by the phenolic transferring an electron (electron transfer (ET))

followed by reversibly transferring a proton (Fig. 2.5) (Wright et al., 2001). The bond

dissociation enthalpy of the hydroxyl groups on phenolics will influence hydrogen atom transfer

whereas the ionization potential is important for determining electron transfer (Wright et al.,

2001). Also, in buffer solutions, the phenolic compound can exist in different protonation states

depending on the pH. Under more basic conditions the phenolic hydroxyls can be deprotonated,

and in this case the phenolic will scavenge radicals by electron transfer that is preceded by

proton loss (Fig. 2.5) (Wang & Zhang, 2005).

Hydrogen atom transfer

AOH + ROO. AO. + ROOH (1)

Electron transfer followed by reversible proton transfer

AOH + ROO. AOH+ + ROO- (2)

AOH+ + H2O ⇌ AO. + H3O+ (3)

H3O+ + ROO- ⇌ H2O + ROOH (4)

Deprotonated phenolic transferring an electron

AOH AO- (5)

AO- + ROO. AO. + ROO- (6)

Figure 2.5. Reactions involved in scavenging of peroxyl radicals by a phenolic antioxidant.

AOH and ROO. represent a phenolic antioxidant and a peroxyl radical, respectively.

The phenolic compound may act as an antioxidant by inhibiting enzymes that produce

reactive oxygen species. For example, xanthine oxidase is an enzyme that can produce the

reactive oxygen species superoxide from hypoxanthine. However, the flavonols quercetin,

kaempferol, and myricetin can inhibit xanthine oxidase activity as seen by inhibition of the

ability of the enzyme to convert xanthine to uric acid (Selloum et al., 2001). Moreover,

phenolics may chelate transition metal ions to prevent the transition metal from producing

14

reactive oxygen species. For example, ferrous iron (Fe2+) is able to generate hydroxyl radicals

from hydrogen peroxide and the hydroxyl radical can then damage DNA. However,

epigallocatechin-3-gallate (and also other phenolics) can prevent DNA damage induced by Fe2+

(Perron et al., 2008). Since the impact of epigallocatechin-3-gallate is reduced upon addition of

ethylenediaminetetraacetic acid (EDTA), the researchers attributed the inhibition of DNA

damage to the formation of phenolic/Fe2+ chelates.

2.1.5. Phenolic antioxidant activity for food preservation

Oils and fats in foods are susceptible to oxidative decay, which can lead to rancidity or

“off-flavours” arising primarily from aldehyde products (Kaur & Perkins, 1991). Aside from

affecting flavour, as noted above (Section 2.1.3) lipid oxidation products might also cause

cardiovascular health problems (see also Addis & Warner (1991) for more on dietary lipid

oxidation products). With more and more people living in cities, more food items undergo a

long transit from raw material to the end consumer. Furthermore, there is increasing trend

towards processed foods containing multiple ingredients, some of which are sensitive to

oxidative decay. In this regard, the omega-3 polyunsaturated fatty acids eicosapentaenoic acid

(EPA) and docosahexaenoic acid (DHA) have been proposed to have health benefits (reviewed

in Mori, 2014) and in 2004 the US Food and Drug Administration allowed for qualified health

claims of reduced risk of coronary heart disease for foods containing EPA and DHA (US Food

and Drug Administration, 2004). Polyunsaturation makes these fatty acids particularly

susceptible to oxidation reactions (Kaur & Perkins, 1991). Antioxidants are a logical choice as

additives to prevent spoilage of foods containing oxidizable lipids. Among the antioxidants used

most widely in industry are the phenolic compounds butylated hydroxyanisole (BHA), butylated

hydroxytoluene, (BHT), butylated hydroxyquinone (TBHQ), and esters of gallic acid (Loliger,

1991). However, some of these (specifically BHA and BHT) have shown toxic effects in some

animal studies, although in these cases dosages were greater than would be expected to be

ingested by humans (European Food Safety Authority (EFSA), 2012). While the synthetic

antioixdants BHT and BHA are still allowed by regulatory agencies, there is a continual search

for new phenolic (and non-phenolic) antioxidants, particularly from natural sources such as

plant food powders (for example carrot, tomato, broccoli, and beetroot) (Neacsu et al., 2015),

15

mint leaf and citrus peel extracts (Viji et al., 2015), honey (Tahir et al., 2015), and licorice

extract (Zhang et al., 2014), to name a few recent works.

2.1.6. Structure-functional correlations among phenolics with antioxidant properties

General relationships between molecular structure of phenolic compounds and

antioxidant activity have been previously identified. The number and location of hydroxyl

groups, along with the presence of double bonds that increase the degree of conjugation, all

seem to have a role in determining antioxidant activity (reviewed in Balasundram et al., 2006).

Hydroxycinnamic acids generally show higher antioxidant activity than corresponding

hydroxybenzoic acids, which may be due to the double bond in the propanoid group of the

hydroxycinnamic acid, providing increased delocalization of the unpaired electron of the

radicalized phenolic (Natella et al., 1999). Increased electron delocalization will stabilize the

phenolic radical thereby making it easier to form, meaning the original phenolic is more

reactive. A study was carried out with the flavonoids to investigate structural features important

for antioxidant activity towards the radical of 2,2'-azinobis(3-ethylbenzothiazoline-6-

sulphonate) (ABTS˙+ radical) in aqueous solution (Rice-Evans et al., 1995). Compounds with an

ortho dihydroxy substitution in the B ring (quercetin and cyanidin) had higher antioxidant

activity than comparable compounds (kaempferol and pelargonidin, respectively) with only a

single hydroxyl group in the B ring (Fig. 2.6). Replacing hydroxyls with O-glycosides (as in 3-

OH in quercetin and 7-OH in naringenin to 3-O-glycoside in rutin and 7-O-glycoside in

naringin, respectively) resulted in decreased antioxidant activity. The presence of the double

bond between carbon two and three in the C ring (present in quercetin but lacking in the

otherwise identical taxifolin (Fig. 2.6)) resulted in higher antioxidant activity. Another study

identified similar relationships between structure and antioxidant activity of flavonoids (Van

Acker et al., 1996). In this case, antioxidant activity was measured for lipid peroxidation and

electrochemical oxidation potentials were also measured. The researchers observed an overall

qualitative correlation between antioxidant activity and oxidation potentials. They identified that

compounds with an ortho dihydroxy substitution in the B ring had the highest activity and that

for such compounds the rest of the molecule was relatively less important in affecting activity.

Among such compounds, quercetin and myricetin showed highest activity. This suggested that,

in combination with the ortho dihydroxy, the double bond between C2 and C3 along with the 3-

16

OH results in a very strong antioxidant, possibly due to the extensive conjugation that such a

compound has (Van Acker et al., 1996). Replacing the 3-OH of quercetin with 3-O-rutinose in

rutin resulted in slightly lower activity for rutin. The importance of the 3-OH became more

prominent when comparing the reduction in activity of a rutin derivative that has OEtOH at 7,

3’, and 4’ positions (and hence lacks the ortho dihydroxy in ring B) to a quercetin derivative that

has OEtOH at 7, 3’, and 4’ positions (Fig. 2.6), reinforcing the idea that in compounds with an

ortho dihydroxy in ring B the rest of the molecular structure is relatively less important for

determining antioxidant activity (Van Acker et al., 1996).

Figure 2.6. Structures of some classes of flavonoids. The ring nomenclature and carbon

numbering system are shown on the flavonol skeleton.

R2

O

O

OHOH

OH

R1

R3

R2

O+

OHOH

OH

R1

R3

R2

O

O

OHOH

OH

R3

R1

R2

O

O

OHOH

R1R3

A

B

R1=R3=H, R2=OH; kaempferol

R1=R2=OH, R3=H; quercetin

R1=R2=R3=OH; myricetin

R1=R3=H, R2=OH; pelargonidin

R1=R2=OH, R3=H; cyanidin

flavonol anthocyanidin dihydroflavonol

R1=R2=OH, R3=H; taxifolin

2 3

4

5

6 7

8

2’

3’

flavanone

4’

R1=R3=H, R2=OH; naringenin

5’

6’

C

17

2.1.7. In vitro measurement of antioxidant activity

There are a variety of assays that have been developed to give some measure of

antioxidant activity. These assays measure hydrogen atom transfer (HAT), electron transfer

(ET), or a combination of the two. Among the more popular hydrogen atom transfer assays are:

oxygen radical absorbance capacity (ORAC), total radical-trapping antioxidant parameter

(TRAP), total oxidant scavenging capacity (TOSC), and crocin bleaching assay (Table 2.4)

(Prior et al., 2005). Among the more popular electron transfer assays is the ferric reducing

antioxidant power (FRAP) assay (Prior et al., 2005). Other popular assays that assess both

hydrogen atom donation and electron transfer include trolox equivalent antioxidant capacity

(TEAC) and 2,2-diphenyl-1-picrylhydrazyl (DPPH) (Table 2.4) (Prior et al., 2005).

Table 2.4. Popular antioxidant activity assays.

Assay name Mechanisma Reagents Biological

relevance

Quantification of activity

ORAC

TRAP

TOSC

crocin

bleaching

FRAP

TEAC

DPPH

HAT

ET

HAT and ET

oxidizer (can

vary), probe

(can vary)

FeIII-TPTZ

ABTS

DPPH

Yes

No

lag time until probe

oxidation and/or decrease in

slope depicting rate of probe

oxidation

Reduction of oxidant at

chosen time

a HAT and ET mean hydrogen atom transfer and electron transfer, respectively.

The concept behind the ORAC, TRAP, TOSC, and crocin bleaching assays is essentially

the same, with the main differences being the reagents that are used and the methods of

detection of reaction progress. In all cases, there is a radical generator as a source of in situ

radicals, a target that acts as a probe for oxidation, and the antioxidant that inhibits oxidation of

the probe by itself sacrificially becoming oxidized (Huang et al., 2005). Different versions of

each method have been developed that use different radical generators and probes. Antioxidant

activity can be quantified as the lag time before oxidation of the probe is initiated and/or the

decrease in the rate of oxidation of the probe. In one version of the ORAC and TRAP assays,

18

both make use of 2,2′-azobis(2-methylpropionamidine) (AAPH) as a temperature-sensitive

peroxyl radical generator and 2’,7’-dichlorofluorescein (DCFH) as the probe (Prior et al., 2005;

Huang et al., 2005). As an alternative to AAPH and DCFH, 2,2’-azobis (2,4-

dimethylvaleronitrile) (AMVN) can be used as a peroxyl radical generator in conjunction with a

lipid soluble probe such as 4,4-difluoro-3,5-bis(4-phenyl-1,3-butadienyl)-4-bora-3a,4a-diaza-s-

indacene (BODIPY 665/676) (Huang et al., 2005). The TOSC assay has been used with a

peroxyl radical generator but also with hydroxyl radicals generated from iron/ascorbate and

peroxynitrite radicals generated from 3-morpholinosydnonimine N-ethylcarbamide (SIN-

1)/diethylenetriaminepentaacetic acid (DTPA) allowing characterizations of different radicals

(Regoli & Winston, 1999). The crocin bleaching assay uses crocin as the oxidizable probe.

Crocin was introduced as a substitute to β-carotene because the former only undergoes radical

oxidation while the latter can undergo light and heat induced oxidation (Prior et al., 2005).

The FRAP, TEAC, and DPPH assays are similar to each other in that there is no probe as

in the case of the purely HAT-based assays. Rather, the reaction between an oxidant and the

antioxidant is measured directly, by measuring the change in oxidant concentration at a chosen

time. The oxidants for the FRAP, TEAC, and DPPH are a complex of FeIII-2,4,6-tris(2-pyridyl)-

s-triazine (TPTZ), 2,2’-azinobis(3-ethylbenzothiazoline-6-sulfonic acid) (ABTS) radical cation,

and DPPH radical, respectively. These assays are considered less biologically relevant than the

purely HAT-based assays because they use oxidants that are not oxygen-based.

2.1.8. Solubility considerations for antioxidant activity

In addition to presence of functional groups like hydroxyls, the solubility of the phenolic

also plays a role in antioxidant efficacy in different solution conditions (reviewed in Shahidi &

Zhong, 2011). It has been previously observed that, in general, non-polar antioxidants are better

than their polar counterparts in protecting a lipid compound that is emulsified in an aqueous

solution, whereas the polar antioxidant is found to be more effective in purely lipid systems

(Porter et al., 1989; Frankel et al., 1994; Cuvelier et al., 2000). This phenomenon is proposed to

occur due to preferential partitioning of the phenolic antioxidant at the interface where oxidation

of the lipid is initiated (Frankel et al., 1994). In an emulsion of lipid in water, the more non-

polar antioxidants would partition to the water-lipid interface and scavenge free radicals before

these radicals propagate into the interior of the lipid micelle. As support for this mechanism, the

19

surfactant effectiveness of a series of acylated hydroxytyrosols (with varying acyl chain-length)

correlated with their antioxidant activity in an oil-in-water emulsion, suggesting that the more

effective antioxidants are those that act as surfactants and partition to the interface of the oil-in-

water emulsion (Lucas et al., 2010). Notably, the antioxidant activity of the octanoate ester of

hydroxytyrosol was greater than both lower chain-length and higher chain-length esters, with a

parallel bell-shaped pattern observed for surfactant effectiveness (i.e. the octanoate ester had

greater surfactant effectiveness than both lower chain-length and higher chain-length

hydroxytyrosol esters) (Lucas et al., 2010). Therefore, this previous work also highlighted the

fact that antioxidant polarity and antioxidant activity do not follow an entirely linear relationship

(Lucas et al., 2010; Shahidi & Zhong, 2011). In the case of pure lipids, two different interfaces

have been posited as being relevant. It was originally suggested that oxidation was initiated at

the air-lipid interface and that polar antioxidants were more effective by preferentially

partitioning to this interface while non-polar antioxidants were less effective because they

remained soluble in the bulk lipid (Frankel et al., 1994). It was later suggested that micro-

aqueous environments exist as reverse micelles and that these are the sites where oxidation is

initiated (Chaiyasit et al., 2007). The polar antioxidants would preferentially partition to these

reverse micelles, thereby being more effective than the non-polar antioxidants that are dissolved

in the bulk lipid.

The condition of emulsification of lipids is common in foods (such as milk and

dressings), and biological systems (such as lipoproteins, whose oxidation is thought to be

associated with cardiovascular disease (Scalbert et al., 2005)). Therefore, one chemical property

of naturally occurring phenolic antioxidants that may be advantageously modified is their

lipophilicity. As such, the lipophilized antioxidants can find application as food preservants; and

may also be used as nutraceuticals that protect against oxidative stress of lipid components in

the human body. At the same time, changes in molecular structure of the phenolic that improve

lipophilicity may also cause changes to the presence of functional groups mentioned above

(hydroxyls and conjugated double bonds) that have been found to be important for antioxidant

activity. Therefore, it may be required to strike a balance between lipophilicity and antioxidant

activity. Two enzyme types that have potential to modify chemistry of natural product phenolics

(for example to increase lipophilicity) are esterases/lipase enzymes and laccase enzymes. As

such, Section 2.3 and Section 2.4 will review esterase/lipase enzymes and laccase enzymes,

20

respectively; but first Section 2.2 will more broadly review approaches to plant phenolic

derivatization.

2.2. Derivatization of plant phenolics

2.2.1. Enzymatic strategies used in plant phenolic derivatization

A variety of enzymes have been used to modify chemistry of plant phenolics. Enzymes

can offer the advantage of stereo- and regioselectivity, which is important in fine-tuning only

selected regions of the molecule. The resultant changes can affect the activity and solubility

properties of the starting compound so that it may be used in novel applications. Lipases can

carry out transesterification reactions using activated esters (such as vinyl esters) to produce

acylated derivatives of phenolics. For example, immobilized lipases were used to acylate the

phenolic hydroxyls of resveratrol (Torres et al., 2010) with vinyl acetate as the acyl donor. In

this case, the authors aimed to selectively acetylate the hydroxyl at position 3 to protect it from

becoming sulfated or glucuronated in the liver, thereby potentially increasing resveratrol’s

bioavailability. The authors found that a lipase from Alcaligenes sp. was able to almost

exclusively acetylate the 3-OH while leaving the other two hydroxyls of resveratrol intact,

whereas other tested lipases showed less selectivity. Similar to the lipases, a select group of

esterases have also been used to acylate phenolic hydroxyls. Topakas et al. (2003) used a

feruloyl esterase to esterify hydroxycinnamic acids in the hopes of improving the lipid solubility

of the compound. Another group of acylating enzymes are the aptly named acyltransferases.

These enzymes typically require an acyl-Coenzyme A (acyl-CoA) substrate as the acyl donor.

For example, a malonyltransferase was used to catalyze the addition of malonyl of malonyl-CoA

to a free hydroxyl of glucose that is covalently linked to an anthocyanin (Suzuki et al., 2002).

The anthocyanins are a type of flavonoid and are notable for the colours they impart to flowers.

Upon malonylation, the anthocyanin pigment was found to be more stable (Suzuki et al., 2002).

Continuing with the theme of transferases, prenyltransferases are another enzyme group

that have been used for modification of bioactive phenolics. Prenylated phenylpropanoids may

have anti-inflammatory and anticancer activity (Paulino et al., 2008; Messerli et al., 2009). A

prenyltransferase from S. spheroides was able to catalyze prenylation of hydroxycinnamic acids,

resveratrol, and some flavonoids (Ozaki et al., 2009). Another group of transferase enzymes that

21

are used in phenolic modification are the methyltransferases. Methylation of flavonoids may be

important for their antifungal and antibacterial properties (Aida et al., 1996; Zhang et al., 2008).

Accordingly, a methyltransferse from tomato was recombinantly expressed in E. coli and

displayed activity on flavonoids including one flavanone, a dihydroflavonol, flavones, and

flavonols (Cho et al., 2012). The enzyme had regiospecificity for the 3’ and 5’ positions of the

flavonoids.

In many cases, natural plant phenolics are found in glycosylated forms. Glycosylation

can impart increased water solubility to the phenolic, among other effects. Some enzymes that

hydrolyze glycosidic bonds also display transglycosylation activity. Such was the case for a

maltogenic amylase from Bacillus stearothermophilus. This enzyme was able to transfer mono-,

di-, and tri-glucose units from maltotriose to the flavonoid naringin (Lee et al., 1999). The major

product, in which maltose had been attached to the already existing glucose of naringin,

displayed not only improved water solubility but also reduced bitterness (Lee et al., 1999).

Finally, oxidase enzymes including laccases and peroxidases can be used to produce

homo- and heterocoupled phenolic products. Makris and Rossiter (2002) used horseradish

peroxidase to oxidize quercetin producing a compound that was later identified as a quercetin

dimer (Gulsen et al., 2007). Ultimately, the authors found the dimer to have reduced activity

(compared to the monomer) in scavenging DPPH radical, hydroxyl free radical, and hydrogen

peroxide (Gulsen et al., 2007). Nicotra et al. (2004) used a laccase from M. thermophyla to

synthesize dimers of resveratrol that might have bioactivities similar to naturally occurring

oligostilbenes. Nugroho Prasetyo et al. (2011) used a laccase from T. hirsuta to couple different

simple phenolics such as catechol onto naringenin. The authors aimed to add hydroxyl groups

that would be conjugated to the isolated C-ring hydroxyl of naringenin, thereby potentially

increasing antioxidant activity (Nugroho Prasetyo et al., 2011). One commom outcome

(intended or unintended) of phenolic derivatization is modification of the solubility of the

phenolic. As this affects the extent to which the phenolic can access different sites in biological

systems, it is a significant motivation of many derivatization processes, and so will be reviewed

in the next two sections (Sections 2.2.2 and 2.2.3).

2.2.2. Increasing hydrophilicity of phenolic compounds

22

Phenolics from the flavonol class have been shown to have protective effects against

ischaemia-reperfusion injury (Williams et al., 2011). For such an application, it is desirable to

increase the water solubility of the antioxidant to administer more of it intravenously into the

blood with fewer injections. One way to increase water solubility is to make a water soluble

prodrug derivative of the phenolic that is converted into the parent compound in vivo. Water

soluble prodrugs of flavonols were shown to have protection against sheep cardiac reperfusion

injury comparable to equimolar quantities of the parent compound (Williams et al., 2011). In

this case, the researchers added phosphate or adipic acid groups to the parent flavonol to make

the prodrug. The prodrug can be converted back to the parent compound by action of

phosphatase or esterase enzymes that are naturally present in tissues and blood. However, this

particular study did not yet examine dosage increases that could potentially be realized with the

water soluble prodrugs and the parent phenolics were administered in solutions containing

DMSO as organic co-solvent. While DMSO is often used in experimental settings, it is

undesirable for clinical applications because it may cause side effects including hemolysis

(Muther et al., 1980; Santos et al., 2003) and its ability to dissolve some plastics used for

intravenous administration (Marshall et al., 1984).

Another approach to increasing the amount of phenolic in aqueous media is to

encapsulate the compound in a carrier that has both hydrophobic core and hydrophilic exterior

(reviewed for quercetin in Cai et al., 2013). In this case, the encapsulated phenolic might also be

protected from undesirable in vivo metabolic modifications on route to the site of action. One

group of encapsulating compounds that can be used is the cyclodextrins. A complex of quercetin

and sulfobutyl ether-7β-cyclodextrin allowed for improved solubilization of quercetin in

aqueous neutral buffer solution (Kale et al., 2006). The orally administered complex also

showed improved tumour growth suppression in mice compared to equivalent doses of the

uncomplexed quercetin (Kale et al., 2006).

A third approach to increase hydrophilicity is to form nanocrystals of the desired

compound. Nanocrystals are highly fine particles of the compound and have a mean particle size

less than 1 µm (typically between 200 nm and 500 nm) (Keck and Muller, 2006). As expected,

the increased surface area of the fine particles leads to increased dissolution rate. Researchers

produced a nanocrystal formulation of the phenolic compound curcumin and this nanocrystal

had a mean diameter of 250 nm compared to 22 µm for crystalline curcumin (Onoue et al.,

23

2010). Compared to crystalline curcumin, the curcumin nanocrystal showed improved

dissolution rate in water and showed improved bioavailability in male Sprague-Dawley rats after

oral administration.

2.2.3. Increasing lipophilicity of phenolic compounds

For some applications, it can be advantageous to increase lipophilicity of antioxidant

phenolic compounds. If a lipid is being targeted for antioxidant protection, then a lipophilic

antioxidant may be more effective than a hydrophilic one. It was previously observed that, in

general, non-polar antioxidants are more effective than their polar counterparts at protecting

lipids dispersed in water (reviewed in Shahidi & Zhong, 2011). In this context, lipophilic

antioxidants can be exploited for preservation of emulsified lipids in foods, such as in

mayonnaise, dressings, and milk.

One of the methods used for increasing lipophilicity of phenolics is to add a long-chain

alkyl group by way of esterification reactions of the phenolic with a long-chain acyl compound

or long-chain alcohol compound (Fig. 2.7). For example, alcohols of varying chain length were

esterified (using sulfuric acid as catalyst) onto caffeic acid to produce lipophilic derivatives

(Aleman et al., 2015). The lipophilic caffeic acid derivatives showed better protection towards

fish oil emulsified in mayonnaise or milk. It should be noted that it was not the longest alkyl

chain ester derivative of caffeic acid that conferred best protection, but rather intermediate chain

length (for mayonnaise) and short chain length (for milk) ester derivatives showed best

protection (Aleman et al., 2015).

24

Figure 2.7. Possible routes to adding long-chain alkyl groups to phenolic compounds to

increase lipophilicity of the phenolic. In A) the long-chain alkyl group is represented by R1 and

is added to the phenolic as part of an acyl group. In the case of B) the phenolic contains a

carboxylic group on one of its ends. The long-chain alkyl group is represented by R4 and is

added to the phenolic as part of an alkoxy group.

A potential way of increasing lipophilicity is by forming dimer and higher level

oligomers of the phenolic compound. Researchers had previously isolated naturally occurring

dimers, trimers, and tetramers formed of successive epicatechin molecules added to catechin

(Fig. 2.8) (Plumb et al., 1998). The (n-octanol)-water partition coefficient of these compounds

demonstrated increasing preference for the hydrophobic n-octanol phase with increasing degree

of oligomerization after the dimer (Plumb et al., 1998) (the dimer did not have a significant

difference in partition coefficient compared to the monomer). However, in this study, the

researchers saw decreased antioxidant activity towards iron/ascorbate induced oxidation of

phospholipid liposomes with increasing lipophilicity (due to increased oligomerization) of the

antioxidant compound. In a related study, catechin monomer and oligomers up to hexamer were

compared for their antioxidant activity toward L-α-phosphatidylcholine liposome, using

different inducers of oxidation (Lotito et al., 2000). When iron/ascorbate (which would be

present in the aqueous phase) was used to induce oxidation, antioxidant activity decreased with

increasing degree of oligomerization of catechin up to the pentamer, showing similar trends as

seen by Plumb et al. (1998). However, when 2,2’-azobis (2,4-dimethylvaleronitrile) (AMVN,

R1 OR2

O

R1 O

O

R3OH R3

OH

OHO

OH

OR4O

HOR2 + +

+ HOR4 + H2O

A)

B)

⇌

⇌

25

which would be present in the lipid phase) was used to induce oxidation, antioxidant activity

increased with increasing degree of oligomerization of catechin up to the pentamer (Lotito et al.,

2000). These previous examples demonstrate that the more lipophilic phenolic derivative is not

always the better antioxidant for emulsified lipids. Changing the inducer of oxidation can

change the pattern of antioxidant activity, and increasing the molecular weight of the antioxidant

can be beneficial up to a certain point but further increases in molecular weight may become

detrimental.

Figure 2.8. Naturally occurring dimers (n=1), trimers (n=2), and tetramers (n=3) made of

successive epicatechin molecules linked to catechin.

The addition of alkyl groups via esterification has been carried out using acid catalysts.

For example, esters of caffeic acid were synthesized using sulfuric acid as catalyst (Aleman et

al., 2015), while rosmarinic acid esters were produced using the strongly acidic sulfonic resin

Amberlite IR-120H (Panya et al., 2012). The use of acid catalyst represents harsh reaction

conditions. On the other hand, the use of enzymes to synthesize novel lipophilic derivatives of

phenolics is an alternative approach that avoids the use of strong acids. In addition, naturally

occurring oligomeric phenolics are present for a limited subset of the phenolic classes including

flavones, flavanols, and hydroxycinnamic acids. Enzymatic catalysis can allow the synthesis of

additional novel oligomeric compounds not readily available from environmental sources.

OH

OOH

OH

OH

OH

H

H

OH

OOH

OH

H

OH

OH

H

n

epicatechin

catechin

26

Esterases and lipases are enzymes that can catalyze esterification reactions while laccases can

catalyze oxidation that leads to oligomerization.

2.3. Esterases/lipases

Broadly speaking, esterases and lipases are enzymes that catalyze the hydrolysis of ester

bonds (Fig. 2.9). Esterases and lipases belong to the general class of enzymes called ester

hydrolases (EC 3.1) by the Nomenclature Committee of the International Union of Biochemistry

and Molecular Biology (NC-IUBMB) (2010b). The ester hydrolase class is further divided to

result in more classes, among of which are carboxylic-ester hydrolases (i.e. esterases and

lipases) (EC 3.1.1), thioester hydrolases (EC 3.1.2), phosphoric monoester hydrolases (EC

3.1.3), and others. Carboxylic-ester hydrolases (EC 3.1.1) are then divided into

carboxylesterases (EC 3.1.1.1), arylesterases (EC 3.1.1.2), triacylglycerol lipases (EC3.1.1.3),

and others. A less formal but often used distinction for the carboxylic-ester hydrolases is to refer

to them simply as either “esterases” or “lipases”. In this case, esterases (for example EC 3.1.1.1)

are differentiated from lipases (for example EC 3.1.1.3) in the tendency of esterases to prefer

short-chain substrates while lipases show activity on both short-chain and long-chain substrates.

From this point on, the terms esterase and lipase will be used rather than the NC-IUBMB

terminology. In addition to classification based on reaction substrates, these enzymes have also

been classified into families based on amino acid sequence features. For example, the

carbohydrate active enzyme (CAZy) classification system, which focuses on enzymes acting on

carbohydrates, comprises a carbohydrate esterase class that is divided into 16 families (Lombard

et al., 2013). Likewise, in the ESTerases and alpha/beta-Hydrolase Enzymes and Relatives

(ESTHER) classification system, esterases and lipases (along with other hydrolases) have been

classified within 148 families based mainly on sequence features and any available biological

data (Lenfant et al., 2013).

27

Figure 2.9. Reaction catalyzed by esterases and lipases. In the case of hydrolysis, R3 is a

hydrogen atom so that HOR3 is water and the final products are a carboxylic acid and an

alcohol. In the case of transesterification, R3 is an alkyl group so that HOR3 is an alcohol and the

final products are a new ester and an alcohol.

2.3.2. Structural features

Esterases and lipases are characterized by an α/β-hydrolase fold structure, which is

defined as a central β-sheet (as opposed to α/β barrel) of eight β-strands connected and

surrounded by six α-helices (Fig. 2.10) (Ollis et al., 1992). Different hydrolases show variability

around this prototypical structure in terms of the number of β-strands and α-helices. For

example, a lipase from Bacillus subtilis and a lipase from Pseudomonas cepacia are both

composed of six β-strands (Van Pouderoyen et al., 2001; Kim et al., 1997b), while an esterase

from Pseudomonas fluorescens is made of seven β-strands (Kim et al., 1997a).

R1 OR2

O

R1 OR3

O

HOR3 + ⇌ HOR2 +

28

Figure 2.10. Prototypical α/β hydrolase fold structure. A) Image adapted from Ollis et al.

(1992). α helices and β strands represented by cylinders and arrows, respectively. Dark circles in

loop regions after β5, β7, and β8 show positions of catalytic residues serine, aspartate/glutamate,

and histidine, respectively. B) 3-Dimensional structure of P. fluorescens esterase showing α/β

hydrolase fold. Catalytic triad residues serine, aspartate, and histidine are shown as sticks

coloured red, blue, and magenta, respectively. Oxyanion hole residues are shown as sticks

coloured in cyan.

A)

B)

29

The catalytic residues form a triad and are found in the order of serine, aspartate, and

histidine in the primary sequence of the enzyme. In some cases, glutamate is present in place of

aspartate, as in the case of a feruloyl esterase from Pleurotus eryngii (Nieter et al., 2014). The

catalytic nucleophilic serine is typically contained in the consensus sequence Gly-X-Ser-X-Gly.

The catalytic serine is located on a sharp γ-like turn (termed the nucleophilic elbow) of the

enzyme secondary structure, going from β5 to the following alpha helix (αC) in the prototypical

α/β hydrolase structure (Fig. 2.10) (Ollis et al., 1992). An important structural feature of the

enzyme, which helps to stabilize the tetrahedral intermediate of the substrate formed during

catalysis, is known as the oxyanion hole. This oxyanion hole is formed by the main-chain amide

hydrogens of two amino acid residues. One of these residues is right after the nucleophilic serine

in the nucleophilic elbow, while the second residue is located at a loop going from β3 to αA in

the prototypical α/β hydrolase structure (Fig. 2.10) (Ollis et al., 1992).

A structural feature that is unique to lipases as opposed to esterases is the presence of a

lid that covers the enzyme active site. This lid is formed from α-helical segments of the protein

that are mobile to allow access of the substrate to the active site (Brady et al., 1990; Brzozowski

et al., 2000; Grochulski et al., 1994; Kim et al., 1997b). The presence of the lid and its

movement have been proposed as an explanation for the phenomenon of interfacial activation of

lipases (Brzozowski et al., 1991). Interfacial activation is the increase in lipase activity that is

observed when the enzyme is present in a solution where the substrate concentration is high

enough to form a separate phase (Verger, 1997). Upon opening of the lipase lid, hydrophobic

patches on the underside of the lid become exposed and may be stabilized by interaction with

the hydrophobic substrate phase at the solution interface (Brzozowski et al., 1991).

Additionally, the lipase active site becomes exposed for catalysis. A recent experiment showed

the feasibility of altering substrate preference of Rhizopus chinensis lipase to favour short chain

substrates by swapping the lipase