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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
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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,
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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.
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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.
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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
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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
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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
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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
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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
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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
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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
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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
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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.
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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
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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).
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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
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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
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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
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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).
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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
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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.
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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
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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
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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
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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),
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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-
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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
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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,
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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
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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,
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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
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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
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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.,
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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).
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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)
⇌
⇌
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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
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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).
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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 +
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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)
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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