Unveiling the Health Related Biological Activities of Sarcocornia Quinqueflora, Atriplex Nummularia and Apium Prostratum

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Unveiling the Health Related Biological Activities of Sarcocornia quinqueflora, Atriplex nummularia and Apium prostratum

A thesis in fulfilment of the requirements for the degree of

MASTER OF PHILOSOPHY

Phitchakorn Norchai

School of Chemical Engineering

Faculty of Engineering

April 2019 Thesis/Dissertation Sheet

Surname/Family Name : Norchai Given Name/s : Phitchakorn Abbreviation for degree as give in the University calendar : Master of Philosophy Faculty : Engineering School : Chemical Engineering Unveiling the Health Related Biological Activities of Thesis Title : Sarcocornia quinqueflora, Atriplex nummularia and Apium prostratum

Abstract Australian native plants have a long history of being used for nutritional and medicinal purposes; however, the scientific foundation of such uses is not well established. In this thesis, three native Australian plants, namely samphire (Sarcocornia quinqueflora), saltbush (Atriplex nummularia) and sea parsley (Apium prostratum), were investigated for their phenolic compositions, antioxidant capacities and inhibitory activities on four enzymes: α-glucosidase, α-amylase, pancreatic lipase and hyaluronidase, for the first time. These enzymes are closely related to disorders such diabetes, overweight, obesity and inflammation. Furthermore, the phenolic compounds in sea parsley were identified and quantified by a combination of high- performance liquid chromatography (HPLC) and liquid chromatography-high resolution mass spectrometry (LC-HDMS) analyses. Phenolic compounds in the plants were extracted with methanol 80% (v/v) and purified with XAD-7 Amberlite® resin. The three plants contained relatively high levels of phenolic compounds and antioxidant capacities as well as enzyme- inhibition activities that are comparable with other Australian native plants. Of the three plants, sea parsley had the highest total phenolic content, exhibited the largest ABTS and DPPH free-radical scavenging capacities, and was the most potent inhibitor of α-glucosidase, α-amylase and pancreatic lipase. Samphire had the highest ferric reducing antioxidant power (FRAP) and was the most potent inhibitor of hyaluronidase while saltbush had the lowest phenolic content, displayed the lowest antioxidant capacity and was the least potent inhibitor of all the enzymes. Purification of the extracts resulted in a significant concentration of the phenolic compounds (1.43-2.67 times) with corresponding increases in the bioactivities. Seven phenolic compounds were identified in sea parsley, with the main ones being apiin (48.2%), apigenin (24.8%), caffeic acid (6%) and ferulic acid (2%), while the minor compounds were ρ-coumaric acid, luteolin and catechin which were present at trace levels (<1%). Significantly, catechin was identified for the first time in the Apium genus. The high levels of health- related bioactivities and the presence of several phenolic compounds known to have disease-preventing effects indicate that the consumption of these native Australian plants could bring significant health benefits to the consumer.

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I hereby grant to the University of New South Wales or its agents the right to archive and to make available my thesis or dissertation in whole or in part in the University libraries in all forms of media, now or here after known, subject to the provisions of the Copyright Act 1968. I retain all property rights, such as patent rights. I also retain the right to use in future works (such as articles or books) all or part of this thesis or dissertation.

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iii

ACKNOWLEDGEMENT

I would like to express my thanks to my supervisor Associate Professor, Jian Zhao, for providing me with the opportunity to conduct research in the Food Science and Technology group at the School of Chemical Engineering, UNSW Sydney, and for his guidance, gigantic knowledge and patience. With the spirit of a ‘true supervisor’ he devoted a great deal of his time to help me. His valuable advice and enthusiastic supervision throughout the course of this study are much appreciated. To Dr. Izabela Konczak, I would like to send my thanks for her inspiration and support. Thank you very much to both of them for their supervision and invaluable advice.

I am grateful to Mr. Camillo Taraborreli for his continued technical assistance and support in the laboratory throughout my Master's degree.

Sincere thanks also to Mr. Lewis Alder from the Bioanalytical Mass Spectrometry Facility for providing expert advice on Liquid Chromatography-High Resolution Mass Spectrometry.

My sincere gratitude goes to my parents for their continuing encouragement and beliefs in me. I also would like to especially thank my sister for her unconditional love and prayer throughout my study.

I wish to express my profound appreciation and thanks to Dr. Chatchaporn Uraipong for her advice, ongoing encouragement, and beliefs in me.

Finally, thanks also to all of my friends, especially Julia Ratna Wijaya. Their supportive and friendly manner has created a stress-free working environment.

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Phitchakorn Norchai

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TABLE OF CONTENTS

TABLE OF CONTENTS…………………………………………………………...... 1

LIST OF TABLES……………………………………………………………………..5

LIST OF FIGURES…………………………………………………………………....7

ABBREVIATIONS………………………………………………………………...…..8

ABSTRACT…………………………………………………………………………...10

CHAPTER 1 INTRODUCTION…………………………………………………….11

CHAPTER 2 LITERATURE REVIEW…………………………………………….16

2.1. Introduction and scope……………………………………………….………….16 2.2. Phytochemicals……………………………………………….………….……….16 2.3. Phenolic compounds……………………………………...….………….….……19 2.3.1. Phenolic acids…………………………………………………………..….…19 2.3.2. Flavonoids…………………………………………………………………….20 2.3.3. Tannins………………………………………………………………………..22 2.4. Health beneficial properties of phenolic compounds…………………………..23 2.4.1. Antioxidant activity……………………………………………………...…....23 2.4.1. The mechanism of antioxidant activity of phenolic compounds……...... 23 2.4.2. Antimicrobial activity……………….……………………………………...... 26 2.4.3. Anti-cancer activity…………………………………………………..……….27 2.4.4. Inhibitory activity on digestion enzymes……………………………………..28 2.4.4.1. α-amylase and α-glucosidase inhibitory activities……………….……29 2.4.4.2. Lipase inhibition activity……………………………………………....30 2.4.5. Anti-inflammatory activity…………………………………………...……….31 2.5. Phenolic compounds in Australian native plants and their health properties…...32 2.5.1. Phenolic compounds identified in Australian native plants……………..……32 2.5.2. Health related biological properties of Australian native plants……………...36 2.5.3. Antioxidant activities of Australia native plants……………………………...36 2.5.4. Inhibitory properties of Australian native plants on digestive enzymes……...40 2.5.5. Anti-inflammatory properties in Australian native plants…………………….43 2.5.6. Antimicrobial properties of Australian native plants…………………………45 1

2.5.7. Other health related properties of Australian native plants…………………...46 2.6. Plants investigated in this study…………………………………………………47 2.6.1. Samphire (Sarcocornia quinqueflora)………………………………………...47 2.6.2. Saltbush (Atriplex nummularia)………………………………………………50 2.6.3. Sea parsley (Apium prostratum)………………………………………………51 2.7. Conclusion………………………………………………………………………...54

CHAPTER 3 MATERIALS AND METHODS……………………………………..55

3.1. Plant materials……………………………………………………………………55 3.2. Chemicals and reagents……………………………………….………………….55 3.3. Extraction of phenolic compounds………………………………………………56 3.3.1. Preparation of crude phenolic extracts………………………………………..56 3.3.2. Preparation of purified extracts……………………………………………….57 3.4. Proximate analysis………………………………………………………………..58 3.4.1 Moisture content……………………………………………………………….58 3.4.2. Ash content……………………………………………………………………58 3.4.3. Lipid analysis………………………………………………………………….59 3.4.4. Protein content……………………………...…………………………………59 3.4.5. Carbohydrate content………………………………………………………….59 3.5. Assay of total phenolic content and antioxidant capacity……………………...60 3.5.1. Preparation of plant extract solutions…………………………………………60 3.5.2. Total phenolic content………………………………………………………...60 3.5.3. Ferric Reducing Antioxidant Power (FRAP) assay……………………...……60 3.5.4. DPPH radical scavenging capacity assay……………………………………..61 3.5.5. ABTS radical scavenging capacity assay……………………………………..61 3.6. Assays of enzyme inhibitory activities…………………………………………..62 3.6.1. Preparation of plant extract solutions…………………………………………62 3.6.2. Hyaluronidase inhibitory activity assay………………………………………62 3.6.3. Pancreatic lipase inhibitory assay………………….………………………….63 3.6.4. α-Amylase inhibitory activity assay……………………………………..……64 3.6.5. α-Glucosidase inhibitory activity assay………….……………………………65 3.7. Identification of and quantification of phenolic compounds…………………..66 3.7.1. High Performance Liquid Chromatography-Photodiode Array Detector (HPLC-DAD) analysis……………………………………………………...….66 2

3.7.2. Quantification of phenolic compounds by HPLC-PDA………………….…...66 3.7.3. Liquid Chromatography-High Resolution Mass Spectrometry analysis……...67 3.8. Statistical Analysis………………………………………………………………..67

CHAPTER 4 PHENOLIC COMPOSITION AND ANTIOXIDANT ACTIVITIES OF SALTBUSH, SAMPHIRE AND SEA PARSLEY……………………………...68

4.1. Introduction……………………………………………………………………....68 4.2. Results and discussion……………………………………………………………69 4.2.1. Proximate composition of saltbush, samphire and sea parsley……………….69 4.2.2. Yield of phenolic extracts from saltbush, samphire and sea parsley……….....69 4.2.3. General discussion on the extraction and purification of saltbush, samphire and sea parsley……………………………………………………….70 4.2.4. Phenolic composition of Australian native saltbush, samphire and sea parsley ………………………………………………………………………………….72 4.2.5. Antioxidant capacity of saltbush, samphire and sea parsley………………….73 4.2.5.1. ABTS free radical scavenging capacity……………………………….73 4.2.5.2. DPPH free radical scavenging capacity……………………………….74 4.2.5.3. Ferric reducing antioxidant power (FRAP) assay……………………..75 4.3. Correlation analysis……………………………………………………………...76 4.4. Conclusion………………………………………………………………………...78

CHAPTER 5 INHIBITORY EFFECTS OF SALTBUSH, SAMPHIRE AND SEA PARSLEY ON HEALTH RELATED ENZYMES………………………………....79

5.1. Introduction………………………………………………………………………79 5.2. Results and discussion……………………………………………………………80 5.2.1. Inhibition against α-amylase and α-glucosidase………………………………80 5.2.1.1. Inhibition against α-amylase…………………………………………...80 5.2.1.2. Inhibition against α-glucosidase……………………………………… 82 5.2.2. Inhibition against pancreatic lipase…………………………………………...86 5.2.3 Inhibition against hyaluronidase……………………………………………….89 5.3. Conclusion………………………………………………………………………92

CHAPTER 6 IDENTIFICATION AND QUANTIFICATION OF PHENOLIC COMPOUNDS IN SEA PARSLEY (APIUM PROSTRAUM)…………………….. 93

6.1. Introduction………………………………………………………………………93

6.2. Results and discussion……………………………………………………………94 6.2.1. Determination of phenolic compounds in sea parsley (Apium prostratum) by high-performance liquid chromatography-photodiode array detector……………….94 6.2.2. Identification and confirmation of phenolic compounds in sea parsley (Apium prostratum) by liquid chromatography-high resolution mass spectrometry (LC- HRMS)………………………………………………………………………………97 6.2.3. Quantification of phenolic compounds by high-performance liquid chromatography-photodiode array detector analysis…………………………….....103 6.3. Conclusion……………………………………………………………………….110

CHAPTER 7 CONCLUSIONS AND RECOMMENDATION…………………...111

REFERENCES………………………………………………………………………115

APPENDICES……………………………………………………………………….141

LIST OF TABLES

Table 2.1. Structures and examples of the main classes of dietary flavonoids ...... 21 Table 2.2. Phenolic compounds found in Australian native plants ...... 33 Table 2.3. Antioxidant capacity in Australia native plant ...... 37 Table 2.4. Correlation analysis between the total phenolic and their antioxidant capacities of Australian native plants ...... 40 Table 2.5. Inhibitory properties of Australian native plants on digestive enzymes ...... 42 Table 2.6. Anti-inflammatory properties in Australian native plants ...... 44 Table 2.7. Australian species of Sarcocornia, their distribution, habitat, growth pattern and number of chromosomes ...... 48 Table 2.8. General information, properties and uses of sea parsley ...... 53 Table 4.1. Proximate composition of Australian native saltbush, samphire and sea parsley (w/w, fresh weight1) ...... 69 Table 4.2. Yield of crude and purified extracts from native Australian saltbush, samphire and sea parsley (%), DW1 ...... 70 Table 4.3. Total phenolic content of crude and purified extracts of saltbush, samphire and sea parsley samples in dry weight of the extract (mg GAE/g, Extract) ...... 73 Table 4.4. ABTS free radical scavenging capacities of crude and purified extracts from native Australian saltbush, samphire and sea parsley1 ...... 74 Table 4.5. DPPH free radical scavenging capacities of crude and purified extracts from native Australian saltbush, samphire and sea parsley1 ...... 75 Table 4.6. Ferric reducing antioxidant power of crude and purified extracts from native Australian saltbush, samphire and sea parsley1...... 76 Table 4.7. Pearson correlation coefficient (r) and significance level (p) for relationships between total phenolic content, and antioxidant assays of Australian native plants ...... 77 Table 5.1. α-Amylase inhibitory activity of crude and purified extracts obtained from native Australian saltbush, samphire and sea parsley ...... 82 Table 5.2. α-Glucosidase inhibitory activity of crude and purified extracts obtained from native Australian saltbush, samphire and sea parsley ...... 84 Table 5.3. Pancreatic lipase inhibitory activity of crude and purified extracts obtained from native Australian saltbush, samphire and sea parsley ...... 88 Table 5.4. Hyaluronidase inhibitory activity of crude and purified extracts obtained from native Australian samphire and sea parsley and saltbush ...... 91 5

Table 6.1. Retention times, co-elution results and UV spectra of the phenolic compounds identified in the methanolic extract of sea parsley (Apium prostratum) as compared with reference standards ...... 97 Table 6.2 .Molecular mass, empirical formula and HPLC retention times of the phenolic compounds identified in the methanolic extract of sea parsley (Apium prostratum) ... 101 Table 6.3 .Quantification of phenolic compounds identified in the methanolic extract of sea parsley (Apium prostratum) by HPLC-PDA ...... 103

LIST OF FIGURES

Figure 2.1. Classification of phytochemicals identified in plants, concentrating mainly on phenolic compounds ...... 18 Figure 2.2. Basic chemical structure of flavonoids ...... 20 Figure 2.3. The three stages of autoxidation: initiation, propagation and termination .. 24 Figure 2.4. Morphological characteristics and growth forms of Sarcocornia ...... 47 Figure 2.5. Salt land site and saltbush ...... 50 Figure 2.6. Sea parsley morphology...... 52 Figure 5.1. α-Amylase inhibitory activity of the crude and purified extracts obtained from native Australian saltbush, samphire and sea parsley...... 81 Figure 5.2. α-Glucosidase inhibitory activity of crude (A) and purified (B) extracts obtained from native Australian saltbush, samphire and sea parsley...... 83 Figure 5.3. Pancreatic lipase inhibitory activity of crude and purified extracts obtained from native Australian saltbush, samphire and sea parsley...... 87 Figure 5.4. Hyaluronidase inhibitory activity of crude and purified extracts obtained from native Australian saltbush, samphire and sea parsley ...... 90 Figure 6.1. Processes used to identify phenolic compounds in the purified extract of sea parsley (Apium prostratum)...... 94 Figure 6.2. Initial HPLC chromatogram of purified sea parsley (Apium prostratum) extracts with a run time of 60 min and the peaks recorded at 320 nm...... 96 Figure 6.3. HPLC chromatogram of purified sea parsley (Apium prostratum) extract with the peaks recorded at 320 nm. Provisional identity of the peaks: catechin (1), caffeic acid (2), ρ-coumaric acid (3), apiin (4), ferulic acid (5), luteolin (6) and apigenin (7)...... 96 Figure 6.4. LC-HRMS total ion current (TIC) chromatogram of sea parsley (Apium prostratum) using heated electrospray ionisation with a mass range between m/z 120-950...98 Figure 6.5. LC-HRMS total ion current (TIC) chromatogram of sea parsley (Apium prostratum) using atmospheric pressure chemical ionisation with a mass range between m/z 120-950...... 99 - Figure 6.6. Comparison of molecular ion [M-H] (m/z 563) of apiin (C26H28O14) in sea parsley (Apium prostratum) (A), the mass spectra of apiin standard (B) and the simulated data generated from Xcalibur (C)...... 100 Figure 6.7. Chemical structure of compounds identified in purified sea parsley (Apium prostratum) extract...... 102 7

ABBREVIATIONS

[M-H]- Molecular ions 4-MUO 4-methylumbelliferyl oleate ABTS 2, 2’-azino-bis (3-ethylbenzothiazoline-6-sulphonic acid) diammonium salt AC Absorbance of control ACB Absorbance of control blank ACE Angiotensin-converting enzyme ANOVA Analysis of variance APCI Atmospheric pressure chemical ionization AS Absorbance of sample ASB Absorbance of sample blank CAT Catalase COX-1 Cyclooxygenase-1 COX-2 Cyclooxygenase-2 DMAB 4-dimethylaminobenzaldehyde DMSO Dimethyl sulfoxide DPPH Di (phenyl)-(2,4,6-trinitrophenyl) iminoazanium DW Dry weight ESI Electrospray FC Fluorescence of control FCB Fluorescence of control blank FRAP Ferric reducing antioxidant power FS Fluorescence of sample FSB Fluorescence of sample blank FW Fresh weight GE Gallic acid equivalent GSH Glutathione GSH-Px Glutathione peroxidase HESI Heated Electrospray Ionisation HHDP Hexahydroxydiphenoyl HPLC High Performance Liquid Chromatography HPLC-DAD High Performance Liquid Chromatography-Diode Array Detector

HPLC-PDA High Performance Liquid Chromatography-Photodiode Array Detector IC50 Half maximal inhibitory concentration iNOS Nitric oxide synthases LC-HRMS Liquid Chromatography-High Resolution Mass Spectrometry LC-MS Liquid Chromatography-Mass Spectrometry LDL Low-density lipoprotein LOD Limit of detection LPF Lipofuscin m/z Mass-to-charge ratio MAPK Mitogen-activated protein kinase MCwb Moisture content on a wet basis MDA Maleic dialdehyde MIC Minimum inhibitory concentration PGE2 Prostaglandin E2 P-NPG p-nitrophenyl-β-ᴅ-glucopyranoside PTFE polytetrafluoroethylene RNS Reactive nitrogen species ROS Reactive oxygen species SD Standard deviation SOD Superoxide dismutase TE Trolox equivalent TEAC Trolox equivalent antioxidant capacity TIC Total ion current TPTZ 2,4,6-tripyridin-2-yl-1,3,5-triazine v/v Volume per volume w/w Weight per weight Wf Weight of final sample Wi Weight of initial sample

ABSTRACT

Australian native plants have a long history of being used for nutritional and medicinal purposes; however, the scientific foundation of such uses is not well established. In this thesis, three native Australian plants, namely samphire (Sarcocornia quinqueflora), saltbush (Atriplex nummularia) and sea parsley (Apium prostratum), were investigated for their phenolic compositions, antioxidant capacities and inhibitory activities on four enzymes: α-glucosidase, α-amylase, pancreatic lipase and hyaluronidase, for the first time. These enzymes are closely related to disorders such diabetes, overweight, obesity and inflammation. Furthermore, the phenolic compounds in sea parsley were identified and quantified by a combination of high-performance liquid chromatography (HPLC) and liquid chromatography-high resolution mass spectrometry (LC-HDMS) analyses. Phenolic compounds in the plants were extracted with methanol 80% (v/v) and purified with XAD-7 Amberlite® resin. The three plants contained relatively high levels of phenolic compounds and antioxidant capacities as well as enzyme-inhibition activities that are comparable with other Australian native plants. Of the three plants, sea parsley had the highest total phenolic content, exhibited the largest ABTS and DPPH free-radical scavenging capacities, and was the most potent inhibitor of α-glucosidase, α-amylase and pancreatic lipase. Samphire had the highest ferric reducing antioxidant power (FRAP) and was the most potent inhibitor of hyaluronidase while saltbush had the lowest phenolic content, displayed the lowest antioxidant capacity and was the least potent inhibitor of all the enzymes. Purification of the extracts resulted in a significant concentration of the phenolic compounds (1.43-2.67 times) with corresponding increases in the bioactivities. Seven phenolic compounds were identified in sea parsley, with the main ones being apiin (48.2%), apigenin (24.8%), caffeic acid (6%) and ferulic acid (2%), while the minor compounds were ρ-coumaric acid, luteolin and catechin which were present at trace levels (<1%). Significantly, catechin was identified for the first time in the Apium genus. The high levels of health-related bioactivities and the presence of several phenolic compounds known to have disease-preventing effects indicate that the consumption of these native Australian plants could bring significant health benefits to the consumer.

CHAPTER 1 Introduction

Due to its vast land mass and unique climate, Australia holds a huge and diverse range of plant species, approximately 24,000, with many of them being native to Australia. Around 5000 of these species are edible and another 5000 have been considered as having medicinal functions (Cooper, 2004). Australian native plants have a long history of being consumed as foods and for medicinal purposes by the indigenous population (Roberts et al., 1990, Tang et al., 2016). Over hundreds of years, the indigenous people have built an extensive knowledge on native plants regarding their food value and medicinal uses for treating various illnesses (Barr et al., 1988). However, with the establishment of modern medical services since European settlement, the medicinal role of native plants has gradually diminished. Even the food uses of native plants have also been largely forgotten by the general public. This situation has changed since the 1980s with the discovery of some native plants with unique richness in certain nutrients, e.g., the extremely high vitamin C content of Kakadu plum, which has stimulated the interest of the public and scientific community in native plants (Brand et al., 1982). The unique and exotic sensory properties of many native foods have also attracted the attention of adventurous chefs and consumers (Hodgson and Wahlqvist, 1993). Nowadays, Australian native fruits, vegetables, herbs and spices are becoming increasingly popular with the public with more and more varieties becoming available in restaurants, specialty shops and supermarkets (Phelps, 1997, Sultanbawa, 2016). In 2013, it was estimated that the gross value of the Australian native food production was about AUD15 - 25 million, and is expected to continue the strong growth in the coming years (Clarke, 2013).

Concurrent to the growing popularity of Australia native foods, modern industrialized societies like Australia are facing a number of major health challenges. Among them is the prevalence of the so called “metabolic syndrome”, which is a cluster of adverse medical conditions including overweight and obesity, hypertension, hyperglycemia and high serum triglycerides (Dandona et al., 2005). Metabolic syndrome is associated with a heightened risk of heart disease, stroke and type 2 diabetes, which are among the leading causes of death in many countries (Brand-Miller and Holt, 1998). Unhealthy life style plays a key role in the development of metabolic syndrome, with poor diet such as over

11 consumption of energy intense foods being a major part of it (Bahadoran et al., 2015, Marlatt et al., 2016). In recent decades, a large number of studies have demonstrated that the consumption of plant-based foods, such as fruits, vegetables and whole grains, can help prevent the development of metabolic syndrome, as well as many other serious diseases such as cancer (Sakulnarmrat and Konczak, 2012). The health benefits of these plant-based foods are generally attributed to the phytochemicals present in them. Phytochemicals, which are all the compounds essentially found in plants, can be separated into four main groups: phenolic compounds, carotenoids essential oils and others which include alkaloids, nitrogen-containing and organosulfur compounds (Bellik, 2012). These compounds are widely found in plant derived foods, especially fruits, vegetables, herbs, spices, pulses and whole grains. Over the last few decades, a huge number of studies have explored the health-related biological activities of phytochemicals, especially phenolic compounds, which is the large group of phytochemicals. These studies have generated an increasing body of evidence demonstrating that these compounds can have preventive and therapeutic effects on several major health conditions and diseases including cardiovascular disease, inflammation, diabetes mellitus, cancer, aging and neurological disorders (Surah, 2003, Liu, 2003, Kroon & Williamson, 2005, Huang et al., 2009, Jacob et al., 2012, Khoddami et al., 2013, Xu et al., 2017).

Parallel to this general interest in the health benefits of phytochemicals in plant-based foods, the health-promoting properties of Australian native plants have also received growing attention from researchers. Lassak and McCarthy (2001) investigated a group of native plants with medicinal properties in different parts of Australia and examined the synthesis of some of the active components. Zhao and Agboola (2007) studied the biological properties of over a dozen native fruits, vegetables, herbs and spices and reported that many of them possessed strong antioxidant capacities as well as antimicrobial activities against several foodborne pathogenic and spoilage microorganisms. In the last decades, several research groups in Australia have sought to evaluate the chemopreventive and therapeutic properties of Australian native plants by examining their effects on a number of health disorders that are prevalent in modern societies (Konczak et al., 2008, Konczak et al., 2010a, b, Tan et al., 2011a, b, Sakulnarmrat and Konczak, 2012, Sakulnarmrat et al., 2013, 2014, Vuong et al., 2014, Tang et al., 2016, 2017). These studies have gathered a growing body of evidence about the health benefiting properties of Australian native plants, including antioxidant, antidiabetic, 12 antimicrobial and anticancer properties using in vitro and tissue cell culture methods. Some of these studies have also sought to identify and quantify the phytochemicals in the native plants in order to provide a foundation for their health-promoting functions. However, these studies have only covered a dozen or so Australian native plants. There are many Australian native plants which have shown potential to be used as vegetables or herbs, and many of which are becoming available in supermarkets and restaurants. There is, therefore, a continual need to explore the health beneficial properties of the emerging native plants. In this Master of Philosophy (MPhil) thesis, three native plants, namely samphire, saltbush and sea parsley, were selected for such studies. The hypothesis is that saltbush, samphire and sea parsley contain bioactive compounds which contribute to health related biological activities, such as antioxidant capacity and inhibition of key digestive and physiological enzymes.

Samphire (Sarcocornia quinqueflora) is a perennial plant which generally grows in intertidal areas, as well as along the inland pans and salt lakes (Datson, 2002). It has a long history of being used as an edible plant in Australia and there are several books and TV programs that have promoted its consumption (Cribb and Cribb, 1975, Lear and Turner, 1977). In restaurants, it is considered as a unique and delicious vegetable which is used in salad or cooked by stir frying, simmering or steaming. In recent years, samphire has become available in many Australian supermarkets all year round as an exotic vegetable. Samphire contains about approximately 3.1% of protein, 9.1% of carbohydrate and 0.5% of dietary fibre, and it is rich in several minerals including calcium (Seward, 2019). Samphire has been found to contain fucoidans, a group of sulfated polysaccharides generally found in brown seaweeds, which reportedly have many health benefits including anti-inflammatory (Christenses and Brandt, 2006) and antioxidant effects (Holtkamp et al., 2008).

Saltbush (Atriplex nummularia), also known as old man saltbush, is a plant from the Amaranthaceae family, which includes a number of common vegetables such as spinach and beetroot, Saltbush is widely found in the semi-arid and arid regions of the Australian mainland on heavy clays and saline soils where they can grow up to 3 m high and 2-4 m wide with 1-3 cm long leaves (Viney, 2007). It is tolerant to frost and drought and is well adapted to the low rainfall conditions which are prevalent in the interior regions of Australia (Phelan, 2006). The plant has a high protein content (12-16%) and is rich in 13 several minerals and vitamins such as vitamin E (Milton, 2008). Fresh saltbush or blanched saltbush leaves are used as wrapper for fish, meat or tofu or as a leafy bed in meat or vegetable grills. It is also stir fried with various herbs and spices or used in salads or as a seasoning for pasta, grills and bread, and other uses (Knowles, 2018).

Sea parsley (Apium prostratum), a plant of the Apiaceae family, is a native plant of Australia, which is found abundantly along the southern coastline. It has narrow green leaves similar to the flat leaf of parsley, but it has a flavour which is rather similar to celery (Peter, 2006). It is used like parsley or celery on salads for flavoring in cooking and as a garnish (Herbst, 2001). Although an annual plant, sea parsley has a robust tap root similar to carrot, giving it a semi-perennial capacity (Gamble, 2019). The Apiaceae family consists of a number of common culinary plants such as celery, carrot, coriander and parsley. Studies have shown that these vegetables and herbs exhibit a number of health benefitting properties such as antihypertensive and anti-inflammatory effects and can be used to treat bronchitis, gastrointestinal infections, asthma, and hepatitis (Singh and Handa, 1995).

To date, no studies have investigated the phenolic composition of the three native Australian plants or their health-promoting biological activities. Therefore, the overall aim of this thesis project was to examine the phenolic composition and health-related biological activities of saltbush, samphire and sea parsley. The specific objectives were:

1. To determine the total phenolic content and antioxidant capacity of saltbush, samphire and sea parsley; 2. To investigate the inhibitory activities of the three plants on four metabolically important enzymes: α-glucosidase, α-amylase, pancreatic lipase and hyaluronidase, using in vitro methods; and 3. To identify and quantify the main phenolic components in the plant with the highest bioactive activities using a combination of high-performance liquid chromatography-photodiode array detector (HPLC-PDA) and liquid chromatography- high resolution mass spectrometry (LC-HRMS) analyses.

The thesis consists of seven chapters. Chapter 1 provides a general introduction to the research topic. Chapter 2 reviews the recent literature on the subject with emphasis on phenolic compounds identified in Australian native plants and their health-related bioactivities. Chapter 3 outlines the research approach and details the experimental procedures used for the whole thesis. Chapter 4 describes the phenolic composition and antioxidant activities of the three Australian native plants. Chapter 5 describes the inhibitory effects of the plants on α-glucosidase, α-amylase, pancreatic lipase and hyaluronidase. Chapter 6 gives a detailed account of the identification and quantification of the phenolic compounds in sea parsley, using HPLC-PDA and LC-HRMS techniques. Finally, Chapter 7 presents a general summary of the main findings of this project and provides some recommendations for future research in the field.

CHAPTER 2 Literature Review

2.1. Introduction and scope

Australia is home to a wide and diverse range of native plants consisting of over 24,000 different species (Cooper, 2004). Around 5000 of these species are edible and another 5000 have been considered as having medicinal functions (Cooper, 2004). Australian native plants have a long history of being consumed as foods and for medicinal purposes by the indigenous populations for thousands of years (Sakulnarmrat et al., 2012). Over the recent decades, there have been numerous studies conducted to investigate the chemical composition and nutritional and health benefitting properties of Australian native plants, which has resulted in the accumulation of a considerable body of scientific knowledge in this area. This review is aimed at summarizing the research findings published in the peer-reviewed scientific literature, focusing on the phenolic compounds that have been identified in Australian native plant foods and the health-related biological functions of these compounds and the plant extracts. The literature review begins with an overview of phytochemicals and phenolic compounds with an emphasis on the types of compounds that are relevant to Australian native plants. The review then provides detail accounts of the phenolic compounds that have been identified from Australian native plants and their biological activities. The review concludes with a description of the three Australian native plant species that will be investigated in this thesis and pointing out the gap of knowledge in the field that this thesis is aimed to address.

2.2. Phytochemicals

Basically, phytochemicals are compounds originated from plants. These compounds are mostly secondary metabolites synthesized by plants with diverse chemical structures and perform important physiological functions in the growth and development of plants, including protecting themselves from the negative effects of the environment such as UV radiation and diseases (Boyer and Liu, 2004, Khoddami et al., 2013). Many phytochemicals are natural antioxidants which can help protecting plants from harmful impacts from the reactive molecules that are generated during photosynthesis (Johnson and Williamson,

2003). From a human health perspective, these compounds are not essential nutrients such as vitamins; however, they contain a diverse range of biologically active constituents that can provide a number of significant health benefits (Surh, 2003, Liu, 2003, Lee and Lee, 2006, Dai and Mumper, 2010).

Phytochemicals can be categorized into four main groups: phenolic compounds, carotenoids, essential oils and alkaloids. Phenolic compounds are molecules which contain hydroxylated aromatic rings where the hydroxyl group is directly attached to the phenyl, substituted phenyl, or additional aryl groups (Bellik et al., 2012). Carotenoids are plant pigments which are responsible for the bright red, yellow and orange hues in most fruits and vegetables. These pigments play an important role in plant health (Khoddami et al., 2013). Essential oils from natural sources encompass a large and diverse group of volatile and aromatic compounds (Marques et al., 2018). Plant alkaloids include nitrogen- containing and organosulfur compounds and others (Bellik et al., 2012). Figure 2.1 presents the classification of phytochemicals identified in plants with particular focus on phenolic compounds (Bellik et al., 2012).

Others (tannins, stilbenes, coumarin)

Gallic, Hydroxybe vanillic nzoic acids acids Phenolic acids Caffeic, Hydroxycin ferulic, -namic acid coumaric acids Phenolic compounds

Quercetin, Flavonols kaempherol , myricetin

Carotenoids Apiginen, Flavones luteolin

Phytochemicals Flavonoids in plants

Hesperetin, Flavonones hesperdin, Terpenoids naringenin Essential oils Catechin, Terpenes Flavanols epicatechin

Anthocyani Cyanidin, -dins pelagonidin Others (alkaloids, nitrogen- containing, organosulfur Isoflavones Genistein compounds)

Figure 2.1. Classification of phytochemicals identified in plants, concentrating mainly on phenolic compounds (Bellik et al., 2012).

2.3. Phenolic compounds

Phenolic compounds constitute a major group of phytochemicals, showing an enormous variety of structures from the simple phenolic acids to polyphenols and polymeric compounds formed from the simpler structures (Khoddami et al., 2013). They are found extensively in fruits, vegetables, herbs, spices, green tea, flowers, seeds, stems and nuts, which are common items of the human diet (Mandal et al., 2017). Phenolic compounds were initially recognized for their role in the sensory quality of edible plants. For instance, they are responsible for the colours of many fruits and vegetables, and the flavour quality like astringency in tea and wine, as well as being the substrates for enzymatic browning (Garcia-Salas et al., 2010). Nowadays, the potential health effects of phenolic compounds have gained considerable attention from the research community (Balasundram et al., 2006, Ramassamy, 2006, Ignat et al., 2011, Huang et al., 2009). Phenolic compounds which are found in Australian native plants are detailed in 2.5.1.

2.3.1. Phenolic acids

Phenolic acids are a major class of phenolic compounds which occur in both free and bound forms in plants (Działo et al., 2016). Phenolic acids are organic compounds containing the phenolic ring with a carboxylic acid functional group. Structurally, they can be divided into two groups: hydroxybenzoic and hydroxycinnamic acids. Caffeic acid, ρ-coumaric acid, and ferulic acid are among the most common hydroxycinnamic acid derivatives while gallic acid is one of the common hydroxybenzoic acids (Manach et al., 2004). Phenolic acids are found widely in different types of foods derived from plant sources such as fruits, vegetables and cereal grains, as well as coffee, herbs and spices (Pawlik and Aladedunye, 2017, Goleniowski et al., 2013). In foods, these acids often exist as esters with carbohydrates or other compounds with hydroxyl groups such as quinic acid (Sakulnarmrat, 2012).

2.3.2. Flavonoids

Flavonoids are the biggest group of phenolic compounds, with more than 8000 flavonoids identified so far (Babu and Liu, 2009). All flavonoids have a basic triple-ring structure, as shown in Figure 2.2., which serves as the backbone of every flavonoid molecule (de la Rosa et al., 2010). Different chemical structures of the triple rings lead to six types of flavonoids, namely flavones, flavonols, flavanols, flavanones, isoflavones and anthocyanidins (Pietta, 2000). The chemical structures of flavonoids can influence their chemical properties and bioactivities. For example, the occurrence of a hydroxyl group at flavonols might provide stronger antioxidant capacities than flavones (Pietta, 2000). The basic structures of the main flavonoids are presented in Table 2.1.

Figure 2.2. Basic chemical structure of flavonoids (Kumar and Pandey, 2013).

Table 2.1. Structures and examples of the main classes of dietary flavonoids

Type of Example Food sources Chemical structure flavonoid compounds Flavones Apigenin Celery Luteolin Parsley Tangeretin Peppermint Chrysin Broccoli Grapefruit Grapes

Orange Flavonols Quercetin Onions Kaempherol Leeks Myricetin Broccoli Isorhamnetin Blueberries Grapes

Flavanones Hesperetin Mint Naringenin Grapefruit Eridictyol Oranges Lemons

Flavanols Catechin Cocoa Epicatechin Chocolate Epigallocatechin Teas Grapes

Anthocyanins Cyanidin Red cabbage Pelagonidin Blueberry Delphinidin Black rice Peonidin Raspberry

Malvidin Isoflavones Daidzin, Soybean Genistin, Green bean Biochanin A Fabaceae Formononetin

(Source: Keen et al., 2005, Meskin et al., 2008, Solanki et al., 2015, Bultosa, 2016, Kang et al., 2018)

Flavonoids are very common in plant sourced foods and beverages such as fruits, vegetables, grains, barks, roots, stems, flowers, tea as well as wine (Panche et al., 2016). In nature, flavonoids can occur in free forms, but also frequently as glycosides, which can alter their chemical properties and bioactivities. This also makes the identification of flavonoids more difficult. 21

Isoflavones are a subclass of flavonoids with structures that resemble estrogens such as β-estradiol (Fan et al., 2015). For this reason, isoflavones are sometimes called“phytoestrogens” although their physiological functions in the body are not yet clear. Isoflavones are found abundantly in legumes, especially in soybeans (Quinhone and Ida, 2015). The health effects of isoflavones are somewhat controversial. While a number of studies have shown that they may have preventive effects on diseases such as atherosclerosis (Liu et al., 2014), cancer (Tan et al., 2011) and ameliorates muscle atrophy (Hirasaka et al., 2016), there are also reports that high intakes of isoflavones may be linked with breast cancer in women (Ghoncheh et al., 2016). However, most of these claims about the health effects of isoflavones, either positive or negative, lack strong supports of clinic research and more studies are needed to clarify the controversy.

Anthocyanidins are a subclass of flavonoids which are largely responsible for the blue, purple and red colours of fruits and vegetables such as berries, cherries, plums, radishes and cabbage (Kris-Etherton and Keen, 2002). Many types of anthocyanins have been identified in plants and most of them are strong antioxidants that could provide considerable health benefits including anti-cancer properties (Faria et al., 2005, Anderson, 2006). For example, studies have shown that anthocyanidins derived from blueberries can suppress the growth of tumour cells (Martin et al., 2003), inhibit angiogenesis (Bagchi et al., 2004) and inducing the apoptosis of cancer cells (Lazze et al., 2004). Anthocyanins from the Australian native fruits Illawarra plums and currants have been reported to possess anti-inflammatory activities (Tan et al., 2011a).

2.3.3. Tannins

Tannins refer to a group of high molecular weight, polyphenolic compounds that occur extensively in plants (Cirkovic et al., 2017). Molecular weights of tannins range from about 500 to over 3,000 Da for gallic acid esters and reach to 20,000 Da for proanthocyanidins (Khanbabaee and Van Ree, 2001, Han et al., 2007). They are divided into two main groups, condensed and hydrolysable tannins (Okuda, 2005). The condensed tannins consist of polyhydroxyflavan-3-ol monomers that are connected as long-chain polymers (Okuda, 2005), whereas esters of gallic acid are defined as hydrolysable tannins. Ellagitannins are a typical example of hydrolysable tannins which can be identified by the occurrence of

22 hexahydroxydiphenoyl (HHDP). The HHDP group is linked by intramolecular bonds between galloyl groups (Aaby et al., 2005). Tannins are common in many plant sourced foods and beverages such as red wine and tea where they give stringency to the products (McGee and Harold, 2004). They are frequently bound with proteins, alkaloids and other compounds to form an insoluble mass, leading to the formation of precipitates in wine and tea (Okuda and Ito, 2011).

2.4. Health beneficial properties of phenolic compounds

2.4.1. Antioxidant activity

Antioxidants refer to the group of compounds that are able to react with free radicals or quench reactive oxygen species, whereby inhibit or retard the process of oxidation that takes place in vitro (e.g., in food products) or in vivo (e.g., in the body) (Aurelia and Gheorghe, 2011). Antioxidants are believed to play an important role in the prevention and mitigation of many diseases such as inflammation, cardiovascular diseases, neurodegenerative disorders and cancer as oxidative stress is shown to be closely associated with the prognosis of these diseases (Pizzino et al., 2017). Sources of antioxidants for the body can be both endogenous and exogenous. Endogenous antioxidants include enzymes such as catalase, superoxide dismutase, glutathione peroxidase and non-enzymatic compounds such as albumin, metallothioneins, uric acid and bilirubin (Pisoschi and Negulescu, 2012). Exogenous or dietary antioxidants come from foods or nutritional supplements that are rich in antioxidant compounds such as vitamin C, vitamin D, vitamin E, carotenoids and phenolic compounds (Pisoschi and Negulescu, 2012, Litescu et al., 2011).

2.4.1.1. The mechanism of antioxidant activity of phenolic compounds

Lipid peroxidation is the chemical process of lipid degradation by oxidation which can occur both in the body tissues and in foods that contain lipids. The process is usually initiated by stimuli such as heat, light irradiation or ions of metals, and proceeds through the mechanism of free radical chain reactions (Shahidi, 1997). Lipid peroxidation is a

23 complex chemical process that involves a series of reactions that can be divided into three stages, namely initiation, propagation and termination, which are detailed in Figure 2.3.

Radical Initiation: • • RH + O2 → R + OH

Radical Propagation: • • R + O2 → ROO ROO• + RH → R• + ROOH RO• + RH → R• + ROH → Oxidation products

Radical Termination: R• + R• → RR R• + ROO• → ROOR

Figure 2.3. The three stages of autoxidation: initiation, propagation and termination. Note: R = side chain; • = free radical/reactive species (Madhavi et al., 1996)

The initiation of lipid peroxidation occurs with the fatty acid molecule losing a hydrogen radical, usually due to the stimulation of heat, light, presence of metals or the action of prooxidants such as hydroxyl radical or lipid radical, with resultant generation of lipid free radical (R•). In the propagation stage, the lipid free radical, which is chemically unstable and very reactive, reacts with oxygen to form a lipid peroxyl radical (ROO•) which, in turn, reacts with another fatty acid molecule, leading to the formation of another lipid free radical. The free radical thus propagates and the reaction becomes self- sustaining. Termination reactions are the final stage in which free radicals combine their electrons to produce new substances or when they encounter antioxidants such as vitamins C, D or E or phenolic compounds, which leads to the formation of termination products. The lipid peroxidation process is well understood and has been extensively reviewed (Brettonnet et al., 2010, Lorenzo and Pateiro, 2013, Martino et al., 2014, Olgun and Yıldız, 2014, Böttcher et al., 2015, Ahmed et al., 2016).

Antioxidants are electron donor compounds which can offer their own electrons to free radicals. By doing so they react with free radicals or quench reactive oxygen species, forming stable products and thus terminating the oxidative process of lipids (Aurelia and Gheorghe, 2011). Phenolic compounds can inhibit the oxidizing chain reactions in several ways, including direct reaction with reactive oxygen species, quenching free radicals, and chelation of metal ions such as Fe3+ and Cu+ and inhibition of oxidative enzymes (Leopoldini et al., 2004). Such abilities of phenolic compounds are related to their molecular structure, especially the number of hydroxyl groups and the presence of conjugation and resonance effects (Rice-evans et al., 1995). Quantum chemical modeling of flavanols demonstrate that the planar conformation of the triple rings permits electronic delocalization extending between adjacent rings, whereby enhance electron donating capacity of the molecules (Russo et al., 2007).

The antioxidant activities of phenolic compounds are closely associated with their health beneficial properties. This is because the metabolic processes of the body can generate a number of reactive oxygen species (ROS) and reactive nitrogen species (RNS), including superoxide (O2•-), alkoxyl (RO•), nitric (NO•), hydroxyl (HO•) and peroxyl (ROO•) radicals (Valko et al., 2006). These highly reactive oxygen and nitrogen species can attack body components and tissues (e.g., cell membrane, DNA and proteins), resulting in organ damage (Klaunig and Kamendulis, 2004). Accumulation of the reactive species can lead to oxidative stress, which has been recognized as a leading cause of many chronic diseases, including inflammation, atherosclerotic cardiovascular disease, cancers, cataract degeneration, aging, obesity, hypertension and type 2 diabetes (Halvorsen et al., 2002, Liu and Finley, 2005, Eberhardt and Jeffery, 2006, Halliwell, 2007). As potent antioxidants, phenolic compounds can prevent the generation and accumulation of these reactive species, and alleviate the body from the oxidative stress, thereby preventing the occurrence of these chronic disorders (Aurelia and Gheorghe, 2011). For example, a number of studies have shown that phenolic acids, tannins and flavonoids can effectively retard lipid peroxidation (Seddik et al., 2010, Maqsood and Benjakul, 2014). Epidemiological studies have demonstrated that consumption of high natural antioxidant diets such as fruits and vegetables can lead to lower risk of chronic diseases including cardiovascular diseases and several cancers (Serafini et al., 2002, Pellegrini et al., 2006, McCullough et al., 2003, Hatamnia et al., 2013).

2.4.2. Antimicrobial Activity

Antimicrobial substances are important to the food industry as they constitute an integral part of the industry’s overall strategy in combating pathogenic and spoilage microorganisms in foods to ensure the food products are not only safe, but also can meet the required shelf-life. Currently, chemical preservatives such as nitrite, nitrate, SO2, parabens and organic acids are the major antimicrobial substances used by the industry (Rai et al., 2016). However, due to consumer concerns about chemical additives, there is a demand in the industry for natural preservatives which might be less harmful to humans and the environment (Gutierrez et al., 2008). From this perspective, the antimicrobial activity of phenolic compounds have been extensively investigated. For example, Puupponen et al. (2012) studied the antimicrobial activity of 17 pure phenolic compounds including both flavonoids and phenolic acids, as well as eight berry extracts, against selected Gram- positive and Gram-negative bacteria, including foodborne pathogens and probiotic bacteria. It was found that different bacterial species, and different strains from the same species, exhibit different levels of susceptibility to different phenolic compounds. Pereira et al. (2007) investigated antimicrobial properties of phenolic compounds extracted from the olive leaves against a number of microorganisms including some gut bacteria and those which may cause human respiratory tract infections (Pseudomonas aeruginosa, Escherichia coli and Klebsiella pneumoniae), Gram positive bacteria (Bacillus cereus, B. subtilis and Staphylococcus aureus), and fungi (Candida albicans and Cryptococcus neoformans). Seven phenolic compounds, namely apigenin 7-O-glucoside, verbascoside, luteolin 7-O- glucoside, caffeic acid, oleuropein, luteolin 4’-O-glucoside and rutin, were identified and quantified. The extracts from olive leaves showed synergistic antibacterial and antifungal effects. Additionally, Mandal et al. (2017) investigated a number of phenolic compounds for their antibacterial effect on Pseudomonas aeruginosa and Staphylococcus epidermidis. Among them, rutin, tannic acid, eugenol, and epigallocatechin gallate displayed the highest antibacterial efficacy.

Phenolic compounds exert their antimicrobial activity through several mechanisms. Interactions with cell membrane is a key antimicrobial mechanism where the compounds could disrupt the membranes with resultant increase in membrane permeability which leads to the loss of the cellular material and, eventually, death of cell (Moreno et al., 2006, Slavin et al., 2017). Interaction of phenolic compounds with cell membrane could occur

26 in two ways. Firstly, phenolics such as flavonoids can block the lipid bilayer and penetrate it. Secondly, phenolics can induce membrane fusion, leading to the leakage of intramembranous materials and aggregation, with consequent cell death (Ikigai et al., 1993, Burt, 2004). Another mechanism that has been proposed is related to the substitution of the alkyl in the phenol nucleus or the aromatic ring of the compounds, which leads to the formation of phenoxyl radicals (Dorman and Deans, 2000). These radicals can interact with the isomeric alkyl substituents present on the surface of the bacterial cell, thus inhibiting its growth. Lou et al. (2012) investigated the antimicrobial activity of ρ- coumaric acid against several Gram-negative bacteria namely E.coli, Salmonella typhimurium and Shigella dysenteriae. It was found that the phenolic acid altered the permeability of the cell membrane and bound with cellular DNA, thus inhibiting the cellular functions. The antimicrobial activity of flavonoids has been widely investigated (Cushnie and Lamb, 2005). It is found that the B ring in the structure of flavonoids can suppress nucleic acid reaction by hydrogen bonding, which results in inhibition on the synthesis of DNA and RNA (Anandh et al., 2014).

2.4.3. Anti-cancer activity

Cancer can essentially be regarded as the uncontrolled growth of cells that eventually invade and spread to other parts of the body (Cooper and Hausman, 2000). Both endogenous and exogenous factors can cause cancer, with the former including inherited mutations, hormones and immune conditions, and the latter including diet, smoking, viral and bacterial infections, chemical and physical carcinogens (Anand et al., 2008). Anti- cancer compounds have the ability to inhibit or mitigate these factors, preventing cancer from occurring or inhibiting the growth of cancer cells. The mechanism of anti-cancer compounds is complex, which depends on the cancer cell itself as well as the compound type (Ali et al., 2013). There are different mechanisms in each substance. For instance, polyphenols that have inhibitory effects on tumorigenesis can be divided into three groups based on their protective actions in the carcinogenic process (Surh, 2003). In the first group, the compounds can interfere with the formation of carcinogens from the precursor substances. In the second group, the representing compounds act as blocking agents by preventing the carcinogens to reach the critical sites in target cells. The third group of inhibitors, often called suppressing agents, exert the effect by delaying, impairing or

27 reversing the malignancy expression post exposure to carcinogens (Surh, 2003). Over the last few decades, a huge number of studies have been carried out to explore the anticancer properties of phenolics or plants extracts that are rich in these phytochemicals, which has accumulated a growing body of evidence in this area (Sakulnarmrat, 2012). For example, catechin, which is the most abundant component (20-30% on a dry weight basis) in green tea, has been found to be able to prevent the melanoma metastasis in murine models. The tumour formation can be inhibited by this compound to increase the animals’ life span (Isemura et al., 2015). Rahmani et al., (2015) showed that catechins can effectively reduce oxidative stress and could play a role in decreasing the incidence of cancer. Proanthocyanidins have been reported to exhibit many health beneficial properties including anti-cancer effects. They are found to prevent the development of breast cancer cells in vivo (Agarwal et al., 2000) and in vitro (Kim et al., 2004, Mantena et al., 2006). Meeran and Katiyar (2007) reported that these substances affect the death of apoptotic cell in a dose-dependent manner. Proanthocyanidins extracted from grape seeds can prevent the growth of epidermal factors and mitogen-activated protein kinase (MAPK), leading to the inhibition of prostate cancer (DU145) cell growth in mice and tissue culture (Tyagi et al., 2003). Extracts from grape seeds can prevent the proliferation of colorectal carcinoma cells, which is found to be related to their pro-apoptotic ability including both mitochondrial membrane loss and caspase-3 activation (Hsu et al., 2009). Another group of phenolic compounds, ellagitannins, extracted from berries have been shown to suppress cancer proliferation (Stoner et al., 2008).

2.4.4. Inhibitory activity on digestion enzymes

Modern industrialized societies face a number of health challenges; chief among them is the over consumption of energy-intensive foods, with resultant overweight and obesity which has become a growing epidemic (Bahadoran et al., 2015). Overweight and obesity could lead to a number of metabolic syndromes including insulin deregulation (Bahadoran et al., 2015, Marlatt et al., 2016), which can cause diabetes and increase the risk of several other diseases including cancer (Dandona et al., 2005). Phenolic compounds are not only well recognized for their antioxidant capacity; they are also shown to be potential inhibitors for several metabolically important digestive enzymes, such as α-amylase, α- glucosidase and pancreatic lipase. These enzymes are responsible for breaking starch and

28 fats into smaller molecules, allowing them to be promptly absorbed by the human body (Sakulnarmrat and Konczak, 2012). Inhibition of these enzymes is one of the strategies to manage overweight and obesity by reducing the digestion and absorption of fats and starch, hence energy consumption by the body.

2.4.4.1. α-Amylase and α-glucosidase inhibitory activities

α-Amylase and α-glucosidase are the key enzymes in the digestive system that are responsible for the digestion of dietary carbohydrates (Muhammad et al., 2016). α- Amylase randomly cleaves the α-(1,4)-glycosidic bonds of starch, generating oligosaccharides while α-glucosidase, which is mostly located in the small intestine, ultimately hydrolyzes the oligosaccharides into glucose that can be easily absorbed by the body (Simpson et al., 2012). The combined action of α-amylase and α-glucosidase is quite efficient in the digestive system. After a starchy meal, especially those with high glycemic indices, the starch can be rapidly degraded into glucose which is immediately absorbed into the body and circulates in blood vessel, resulting in postprandial hyperglycemia (Kwon et al., 2006). This can be quite dangerous for diabetic patients whose body lacks the ability to produce insulin to metabolize glucose efficiently (Tadera et al., 2006). Inhibitors of α- amylase and α-glucosidase can delay the hydrolysis of carbohydrates in the system and, thus, prevent postprandial hyperglycemia (Martins et al., 2015). Currently, α-amylase and α-glucosidase inhibitors such as acarbose, are standard prescription drugs for the management of type 2 diabetes mellitus. However, these synthetic drugs can have a range of side effects including diarrhea, flatulence and nausea (Silavwe et al., 2015). Therefore, it is recognized that inhibitors of carbohydrate-degrading enzymes derived from natural sources could be a supplement or alternative to synthetic drugs (Hayes and Tiwari, 2015). Studies have shown that a range of natural materials, including fruits and vegetables, herbs and spices, medicinal plants and seaweeds contain inhibitory activities against α- amylase and α-glucosidase that could potentially be used for managing diabetes (Bo-wei et al., 2017).

The inhibitory activities of plants against α-glucosidase and α-amylase are generally related to their phenolic compounds (Apostolidis et al., 2006, Kwon et al., 2006, Cazzola et al., 2011, El-Beshbishy and Bahashwan, 2011, Wongsa et al., 2012). For example, the

29 total phenolic content of a number of fruits and herbs has been shown to be significantly correlated with their α-glucosidase and α-amylase inhibitory activities (Wongsa et al., 2012). Several studies have investigated the α-glucosidase inhibitory activity of individual phenolic compounds. Of the flavonoids, quercetin is found to be an especially potent α- glucosidase inhibitor (IC50 105.7 µg/ml) that is more than 20 times stronger than the antidiabetic drug acarbose (IC50 2549.0 µg/ml) under the experimental conditions (Quan et al., 2019). For phenolic acids, caffeic acid showed greater potency in inhibiting α- amylase activity than ρ-coumaric acid, whereas both phenolic acids contribute good α- glucosidase inhibitory activities. Kwon et al. (2006) examined the influence of pH on the inhibitory effects of phenolic compounds on α-glucosidase and reported that pH can have a significant influence on the α-glucosidase inhibitory activity. It is observed that pH 6.5-7.5 elicited the highest inhibitor activity (>80%) for rosmarinic acid, caffeic acid and catechin, while pH in the range of 3.5-4.5 produced low activities. However, some phenolic acids, i.e., hydroxybenzoic, vanillic and protocatechuic acid, exhibited higher activities under acidic conditions.

2.4.4.2. Lipase inhibition activity

Pancreatic lipase is the main enzyme that is involved in the digestion of lipids. It hydrolyzes triacylglycerols into free fatty acids and intestinal lumen monoacylglycerols, and the free fatty acids can then be absorbed into the body (Yang et al., 2014). However, over absorption of fats can lead to accumulation of them in adipocytes, and with time, this can result in chronic weight gain and obesity, which has become one of the leading causes for many disorders in the society (Ghigliotti, et al., 2014). Lipase inhibitors have the capacity to reduce fat hydrolysis and absorption in the gastrointestinal tract with the undigested fats excreted in feces rather than being used as a source of energy, which, if persists, can lead to weight loss to the individual (Tushuizen et al., 2007). Recently, the pancreatic lipase inhibitors have received considerable amount of research for this reason as well as the potential for pancreas to help restore the organ for regular insulin production and reverse the symptoms of diabetes (Tushuizen et al., 2007, You et al., 2012, Yang et al., 2014).

Many phenolic compounds have been reported to exhibit pancreatic lipase inhibitory activities (Bustanji et al., 2010, Ikarashi et al., 2011). For instance, phenolic acids (e.g., gallic, ellagic and ferulic) and flavonoids (e.g., luteolin, kaempherol, genistein and quercetin) have been found to be good inhibitors of pancreatic lipase (Sergent et al., 2012). Coumarin derivatives, flavonoid glycosides in peanut shell, flavan-3-ols, phenolic acids, phenolic esters and mangiferin in mango leaf are also considered as strong pancreatic lipase inhibitors (Birari and Bhutani, 2007). A number of flavonols and tannins have also been reported to exhibit lipase inhibition properties (Kurihara et al., 2006).

2.4.5. Anti-inflammatory activity

Inflammation is a basic mechanism of the body to respond to injury and infection, while pain, swelling and fever are the most common symptoms of the inflammatory process (Miliani, et al., 2006). Furthermore, excessive oxidative stress may cause chronic inflammation, which has been recognized as a causative factor in various cancers including oesophageal, gastric, intestinal, prostate, bladder and thyroid cancers (Balkwill and Mantovani, 2001, Mantovani et al., 2008). Because oxidative stress is a major cause of chronic inflammation, antioxidants, due to their capacity to neutralize the reactive species, could have a major role in inflammation lessening (Reuter et al., 2010). As discussed in section 2.4.1, phenolic compounds are potent natural antioxidants and, hence, can help mitigate the responses to inflammation generated from the reactive species (Peter, 2012). However, the cause of inflammatory processes can also come from enzyme-related pathways, involving pro-inflammatory enzymes such as hyaluronidase, cyclooxygenase-1 (COX-1), cyclooxygenase-2, COX-2 and inducible nitric oxide synthases (iNOS) as well as the products from them, i.e., prostaglandin E2 (PGE2) and nitric oxide (NO) (Soberón et al., 2010, Tan et al. 2011d, Guo et al., 2014). Hyaluronidases are a family of enzymes that catalyze the hydrolysis of hyaluronic acid, a major carbohydrate polymer of the extracellular matrix. Hydrolysis of hyaluronic acid lessens the intercellular viscosity and facilitates cell proliferation in tissues, which can consequently lead toward extracellular matrix degradation and inflammation (Jeong et al., 2000). Inhibition of hyaluronidase can be thus be used as a method for mitigating and preventing inflammation (Jeong et al., 2000, Salmen, 2003). Over the last few decades, a number of studies have been carried out to investigate the hyaluronidase inhibiting activities of plants including fruits, vegetables and herbs. For example, Ippoushi et al. (2000) 31 evaluated the hyaluronidase inhibiting activity of 46 different herbs and vegetables. It was found that herbs in the Lamiaceae family have stronger inhibitory effects than other families of herbs and vegetables studied. Lemon balm was observed to have the highest efficiency and the authors identified rosmarinic acid as the main hyaluronidase inhibitor in the herb. Soberón et al. (2010) investigated the inhibitory effect of flavonoids against hyaluronidase and found that quercetin glycosides exhibited the highest activity compared to several other flavonoids, including hyperoside, isoquercitrin and rutin.

2.5. Phenolic compounds in Australian native plants and their health properties

2.5.1. Phenolic compounds identified in Australian native plants

Australia has a unique ecosystem that consists of a large and diverse range of native plants. Many of these plants have been used as food or for medicinal purposes by the indigenous population for thousands of years (Sakulnarmrat, 2012). In recent decades, native plants have received increasing attention for their potential health beneficial properties. A number of studies have sought to identify the phenolic compounds and other phytochemicals in the plants in an attempt to elucidate the chemical basis of their health properties (Balasundram et al., 2006). Initially, the investigations are focused on native fruits. In recent years, however, studies have extended to native vegetables, herbs and spices. Table 2.2 lists phenolic compounds that have been found in Australian native plants.

Table 2.2. Phenolic compounds found in Australian native plants

Phenolic Common name Amount References Compound (Scientific name) (mg/g DW) Chlorogenic acid Bush tomato 0.42 (Konczak et al., 2010) (Hydroxycin- (Solanum centrale) namic acid) Anise Myrtle 25.7 (Sakulnarmrat, 2012) (Syzgium anisatum) Lemon Myrtle 35.3 (Backhousia citriodora) Tasmannia pepper 359 (Tasmannia lanceolata) Rabbiteye blueberry 160 (Vaccinium ashei Reade) Quandong 259 (Tang et al., 2015) (Santalum acuminatum) River mint 15.4 (Mentha australis) Caffeic acid Bush tomato P (Sommano et al., 2013, (Hydroxycin- Finger lime Konczak et al., 2010) namic acid) (Citrus australasica) (Sommano et al., 2013) Tasmannia pepper 1.1 (Konczak et al., 2010) River mint Trace (Tang et al., 2015) Mint bushes Trace (Tang et al., 2017) (Prostanthera rotundifolia) ρ-Coumaric acid Bush tomato P (Konczak et al., 2010) (Hydroxycin- Mint bushes Trace (Tang et al., 2017) namic acid) Glucose ester of ρ- Mint bushes1 24.0 (Tang et al., 2017) coumaric acid Ellagic acid Anise Myrtle 152.9 (Sakulnarmrat, 2012) (Hydroxycin- Davidson’s plum 36.4 namic acid) (Davidsonia pruriens) Lemon Myrtle 102.0 Ferulic acid Bush tomato 0.82 (Sommano et al., 2013) (Hydroxycin- Wattle seed (Konczak et al., 2010) namic acid) (Acacia victoriae) 1-O-β-ᴅ Mint bushes 14.6 (Tang et al., 2017) glucopyranosyl sinapate Rosmarinic acid River mint 160.4 (Tang et al., 2015) (Hydroxycin- namic acid) Verbascoside River mint 127.1 (Tang et al., 2017) (Hydroxycin- namic acid) 4methoxycinnamic Mint bushes 94.7 (Tang et al., 2017) acid Vanilic acid Finger lime (Sommano et al., 2013) (Dihydroxybenzoic Lemon myrtle acid) Biochanin A River mint 9.6 (Tang et al., 2015) (Isoflavone)

Table 2.2. (Continued) Phenolic compounds found in Australian native plants

Phenolic Common name Amount References Compound (Scientific name) (mg/g DW) Catechin Kakadu plum (Sakulnarmrat, 2012) (Flavanols) (Terminalia ferdinandiana) Anise Myrtle 17.3 (Sommano et al., 2013) Lemon myrtle Hesperetin Anise Myrtle 4.10 (Sakulnarmrat, 2012) (Flavanone) Lemon myrtle 3.82 (Konczak et al., 2010) River mint Trace (Tang et al., 2015) Hesperidin Bush tomato (Sommano et al., 2013) (Flavanone) Davidson’s plum Mint bushes Trace (Tang et al., 2017) Kaempferol Quandong 0.61 (Konczak et al., 2010) (Flavonol) Riberry 0.03 (Syzygium luehmannii) Luteolin Wild lime (Sommano et al., 2013) (Flavone) (Adelia ricinella) Myricetin Davidson’s plum 9.87 (Sommano et al., 2013, (Flavone) Lemon myrtle 3.61 Sakulnarmrat, 20012) Anise Myrtle 4.14 (Konczak et al., 2009) Naringenin Bush tomato (Sommano et al., 2013) (Flavanone) Davidson plum Kakadu plum Lemon myrtle Wattle seed Wild lime River mint Trace (Tang et al., 2015) Mint bushes Trace (Tang et al., 2017) Narirutin River mint 27.2 (Tang et al., 2015) (Flavanone) Neoponcirin River mint 145.0 (Tang et al., 2015) (Flavanone) Quercetin Davidson’s plum 6.08 (Konczak et al., 2010, (Flavonol) Quandong 9.9 Sakulnarmrat, 2015) Anise Myrtle 3.38 (Sakulnarmrat, 2012) Tasmannia pepper 0.86 (Konczak et al., 2009) Chrysanthemin Tasmannia pepper 23.95 (Konczak et al., 2009) (Anthocyanins) Quandong 5.76 Davidson’s plum 4.3 (Netzel et al., 2007, Native pepper berries 11.19 Konczak et al., 2010a) (Tasmannia lanceolata) Antirrhinin Tasmannia pepper 55.26 (Konczak et al., 2009) (Anthocyanins) Peonidin 3- Finger lime 31.6 (Netzel et al., 2007) glucoside (Anthocyanins) Cyanidin 3,5- Native pepper berries 88.1 (Netzel et al., 2007) diglucoside (Anthocyanins) 1Values are expressed as mg gallic acid equivalent (GE) per g purified extract for compounds. P – Possible (confirmation required)

Phenolic compounds identified in Australia native plants include phenolic acids and flavonoids. Hydroxycinnamic acid is the main type of phenolic acids found in the plants, with chlorogenic and caffeic acids being the most commonly identified compounds. Chlorogenic acid is found in several Austrian native herbs and spices including anise myrtle, lemon myrtle, river mint and Tasmanian pepper as well as the native fruits bush tomato, rabbit eye blueberry and quandong. The level of chlorogenic acid is particularly high in the native Tasmannia pepper, where it reaches 359 mg/g dry weight (DW). It is also fairly high in the native fruits rabbit eye blueberry and quandong (160 and 259 mg/g DW), but the level is rather low in bush tomato (0.42 mg/g DW).

Caffeic acid has been identified in the native fruits bush tomato and finger lime, the native vegetable mint bush, the native mint and the native pepper, but mostly at low or trace levels. Other phenolic acids identified include ρ-coumaric, ellagic, ferulic and rosmarinic acids. Ellagic acid is present at a high level in anise myrtle and lemon myrtle while rosmarinic acid is present in river mint at a high level. In comparison, the number of dihydroxybenzoic acids reported for Australian native plants is relatively small. Vanillic acid is one example which is found in finger lime and lemon myrtle. Verbascoside, which is a caffeoyl phenylethanoid glycoside, has recently been identified in the native mint at the concentration of 127.1 mg/g DW (Tang et al., 2017). Verbascoside, also known as acteoside, has been found in various medicinal plants and reportedly possess antimicrobial (Pardo et al., 1993), antioxidant (Bilia et al., 2008) and anti-inflammatory (Akdemir et al., 2011) activities.

Australian native plants contain a number of flavonoids, which included flavones (apigenin, luteolin and myricetin), flavanols (catechin, kaempferol, quercetin and rutin), flavanones (hesperetin, hesperidin, naringenin and narirutin), isoflavone (biochanin A) and anthocyanins and their glycosides (chrysanthemin, antirrhinin, peonidin 3-glucosides and cyanidin 3, 5-diglucoside). Naringenin has been found in the largest number of plants, eight in total, but it is present only at trace levels in them. Chrysanthemin is the second most commonly identified flavonoid, found in five native plants (four of which being fruits), with the levels ranging from 4.3 mg/g in Davidson plum to 23.95 mg/g DW in Tasmanian pepper. Quercetin and myricetin have each been identified in four native plants, with the former ranging from 0.86 to 9.9 mg/g and the latter 3.61 to 9.87 mg/g DW. The other flavonoids are reported in three native plants or less. Of all the flavonoids 35 identified in Australia native plants, neoponcirin was found to occur at the highest level. This flavanone is present in river mint at 145.0 mg/g DW; however, to date, it has not been reported in other Australian native plants. Other abundant flavonoids include cyaninin 3, 5- diglucoside, antirrhinin, peonidin 3-glucosides and chrysanthemin in Tasmanian pepper berries (88.1, 55.26, 31.6 and 23.95 mg/g DW, respectively), narirutin in river mint (27.2 mg/ g DW) and catechin in anise myrtle (17.3 mg/g DW). The other flavonoids are present in the plants at levels mostly lower than 10 mg/g DW or trace amounts.

2.5.2. Health related biological properties of Australian native plants

Traditionally, Australian native plants have served as a major source of folk medicine for the aboriginal tribes (Barr et al., 1988). The native people have an extensive knowledge about how to use Australian native plants for curative functions on various illnesses and diseases. This knowledge was accumulated from years of observations and passed down through the word of mouth from generation to generation. However, the scientific basis of the curative effects of the native plants is not known to the indigenous populations. In recent decades, with worldwide interest in the pharmacological properties of phytochemicals, Australian native plants have also received increasing attention and a large number of studies have been conducted to determine their health benefits as presented in Tables 2.3.-2.6.

2.5.3. Antioxidant activities of Australia native plants

Many studies have been carried out to determine the antioxidant activities of Australian native fruits, herbs and spices (Table 2.3.). Generally, the studies involved extracting the phenolic fraction from the plant materials, usually using a mixture of water and alcohol, such as methanol and ethanol, at various proportions and the antioxidant activity of the extract is then determined using several assay procedures with ABTS (2, 2’-azino-bis(3- ethylbenzothiazoline-6-sulphonic acid) diammonium salt) and DPPH (Di(phenyl)-(2,4,6- trinitrophenyl) iminoazanium) radical scavenging, FRAP ) Ferric reducing antioxidant power( and ORAC (Oxygen radicals absorbance capacity) being most common. Among the different methods, FRAP has been used in all the studies, which has made a comparison of the results possible. 36

Table 2.3. Antioxidant capacity in Australia native plant

Antioxidant Capacity (Dry weight basis) Common name ABTS DPPH FRAP TPC References (Scientific name) μmol μmol μmol mg GE/g TE/g TE/g Fe2+/g Anise myrtle 2158 55.9 (Konczak et al., (Syzgium 2010b) anisatum) Bush tomato 206 12.4 (Konczak et al., (Solanum centrale) 2010b) Lemon myrtle 1225 31.4 (Konczak et al., (Backhousia 2010b) citriodora) Mint bushes 262.4 110.2 606 42.1 (Tang et al., 2017) (Prostanthera rotundifolia) River mint 398.5 178.9 1933 76.3 (Tang et al., 2015) (Mentha australis) Spearmint 403.5 168.2 1859 83.1 (Tang et al., 2017) (Mentha spicata) Mountain pepper 1315 102.1 (Konczak et al., (Tasmannia 2010b) lanceolata)

Antioxidant Capacity (Fresh weight basis) Common name ABTS DPPH FRAP TPC References (Scientific name) μmol μmol μmol mg GE/g TE/g TE/g Fe2+/g Anise myrtle 8054 729 (Sakulnarmrat et al., 2012) Peppermint 13.2 (Konczak et al., 2010b) Tasmannia 4444 912 (Sakulnarmrat et pepper al., 2012) (Tasmannia lanceolata) Lemon myrtle 5025 661 (Sakulnarmrat et (Backhousia al., 2012) citriodora) Bay leaf 3040 319 (Sakulnarmrat et (Laurus Nobilis) al., 2012) Australian desert 34.8 10.8 (Konczak et al., lime 2010a) (Citrus glauca) Brush cherry 27 12.6 (Netzel et al., (Syzygium 2007) australe) Blueberries 4.5 (Konczak et al., 2010b)

Table 2.3. (Continued) Antioxidant capacity in Australia native plant

Antioxidant Capacity (Fresh weight basis) Common name ABTS DPPH FRAP TPC References (Scientific name) μmol μmol μmol mg GE/g TE/g TE/g Fe2+/g Burdekin plum 192 27.1 283.4 17.1 (Netzel et al., 2006, (Pleiogynium Netzel et al., 2007) timorense) Cedar bay cherry 129.5 9.6 233.3 65.0 (Netzel et al., 2006, (Eugenia Netzel et al., 2007) reinwardtiana) Davidson’s plum 36.5 3.2 49.3-53.9 15.9-16.8 (Konczak et al., (Davidsonia 2010a, Netzel et al., pruriens) 2007) 9258 949 (Sakulnarmrat et al., 2012) Finger Lime 12.6)g( 12.6 )g( (Konczak et al., (Citrus 23.3 )p( 23.3 )p( 2010a) australasica) 8.7 )r( 13.8 )r( 10.9 )y( 16.2 )y( (Netzel et al., 2007) Illawarra plum 122.8 8.9 214.8 68.2 (Netzel et al., 2006, (Podocarpus Netzel et al., 2007, elatus) Tan et al., 2011b) Kakadu plum 204.4 691 74.7-160 (Konczak et al., (Terminalia 2010a, Tan et al., ferdinandiana) 2011b) 27.2 (Netzel et al., 2006) Lemon aspen 14.0 10.8 (Konczak et al., (Acronychia 2010a) acidula) Molucca 45.1 5.3 66.6 21.9 (Netzel et al., 2006, raspberry Netzel et al., 2007) (Rubus moluccanus) Mountain pepper 123.2 11.8 186.7 82.5 (Konczak et al., berry 2010b) (Tasmannia 14.0 (Netzel et al., 2007, lanceolata) Netzel et al., 2006) Muntries 123.8 15.4 267.6 67.1 (Netzel et al., 2006, (Kunzea pomifera) Netzel et al., 2007, Tan et al., 2011b) Quandong 123 50.4 (Konczak et al., (Santalum 2010a) acuminatum) 3225 543 (Sakulnarmrat et al., 2012) Riberry 28.1 33.2 7.5 (Konczak et al., (Syzygium 2010a, Netzel et al., luehmannii) 2007) Rabbit eye 6098 504 (Sakulnarmrat et (Vaccinium ashei) al., 2012)

Table 2.3. (Continued) Antioxidant capacity in Australia native plant

Antioxidant Capacity (Fresh weight basis) Common name ABTS DPPH FRAP TPC References (Scientific name) μmol μmol μmol mg GE/g TE/g TE/g Fe2+/g Native currant 1256 1402 (Tan et al., (Acrotriche 2011b) depressa) Southern 4811 551 (Sakulnarmrat highbush et al., 2012) (V.darrowii x V. corymbosum) 1GE: gallic acid equivalent; TE, trolox equivalent; Fe2+: iron (II) equivalent; TPC: total phenolic content; FRAP: ferric reducing antioxidant power. Data expressed in dry and fresh weight basis are in brackets. Finger limes: green (g), pink (p), red (r) and yellow (y).

The FRAP values for the different native plants ranged from the low value of 14.0 μmol GE/g the native fruit lemon aspen to the very high value of 9258 μmol GE/g for the fruit Davidson plum (as reported by Sakulnarmrat, et al., 2012). Apart from Davidson plum, a number of other native plants also gave high FRAP values including, in descending order, anise myrtle, rabbit eye berry, quandong, river mint and mountain pepper; the FRAP values of these plants are greater than 1000 μmol GE/g. It should be pointed out, however, that the variation in the reported FRAP values for the same plants can vary greatly in different studies. For example, Konczak et al. (2010b) reported a FRAP value of 2158 μmol GE/g for anise myrtle, while a value of 8054 was reported by Sakulnarmrat, et al. (2012) for the same herb. Similarly, much higher FRAP values were reported by Sakulnarmrat and co-workers for several other native plants than those reported by the Konczak group. The apparent reason for the large discrepancies is not clear, but could be partly related to conditions of the samples (source, location, harvest time, etc) and, perhaps, how the assays were performed in the different laboratories. Further research is therefore needed to identify the causes of the discrepancies.

In most of the studies on the antioxidant capacity of Australian native plants, the phenolic content of the plants was usually measured alongside the antioxidant capacities, because it is well established that the antioxidant activity of alcoholic extracts was mainly due to their phenolic content. Australian native plants tend to have a higher total phenolic content than those found in the leaves of blueberries (4.5 mg GE/g, FW) and peppermint (13.2 mg GE/g, DW) (Konczak et al., 2010b). However, Tang et al. (2016) reported that 39 the total phenolic contents of the native river mint (76.3 mg RE/g, DW) and mint bush (42.1 mg RE/g, DW) were lower than the common spearmint (83.1mg RE/g, DW). Using the data in Table 2.3., a correlation analysis was carried out between the total phenolic content of the Australian native plants and their antioxidant capacities obtained by the ABTS radical scavenging and FRAP assays and the results are presented in Table 2.4.

Table 2.4. Correlation analysis between the total phenolic contents and antioxidant capacities of Australian native plants

TPC FRAP ABTS TPC 1.000 0.9192** 0.9139** FRAP 1.000 0.8221* ABTS 1.000 *Correlation is significant at the 0.05 level; ** Correlation is significant at the 0.01 level. TPC: total phenolic content, FRAP: ferric reducing capacity power and ABTS: 2,2’-azino-bis (3ethylbenzothiazoline- 6-sulphonic acid).

As can be seen, there are significant positive correlations between the total phenolic content and antioxidant capacities obtained by both the ABTS and FRAP assays (p < 0.01). The antioxidant capacities obtained by ABTS and FRAP assays are also significantly correlated (p<0.05). This indicates that phenolic compounds in the Australian native plants had a direct relationship with their antioxidant capacities in both ferric chelation and free radical scavenging.

2.5.4. Inhibitory properties of Australian native plants on digestive enzymes

The inhibitory properties in Australian native plants on digestive enzymes are presented in Table 2.5. Several studies have examined the inhibitory effects of native plants on α- amylase and α-glucosidase, and most of the plants examined were found to exhibit a moderate level of such activities compared with common fruits and vegetables. Of the native plants studied, southern highbush was found to be the strongest inhibitor of α- glucosidase, followed by rabbit eye berry, lemon myrtle, Davidson’s plum. Only three studies have determined α-amylase inhibitory activity of two native plants, the river mint and Tasmania pepper. Both were found to exhibit some inhibitory activities on the

40 enzyme but the activities were not very strong. Sakulnarmrat and Konczak (2012) examined the α-glucosidase inhibitory activity of phenolic compounds extracted from several native plants. It was found that lemon myrtle, anise myrtle and Tasmannia pepper leaf displayed higher activities than the common herb bay leaf. Sakulnarmrat et al. (2012) showed that phenolic extracts from quandong and Davidson’s plum were potent inhibitors of α-glucosidase with IC50 of 0.39 mg/mL and 0.13 mg/mL, respectively. In a more recent study, Tang et al. (2017) examined the α-glucosidase inhibitory activity of the phenolic extract from Australian native herb Prostanthera rotundifolia and found that the crude extract had a medium level of potency on the enzyme (24.6% inhibition), which was higher than the common spearmint (15.6%).

A few studies have investigated the pancreatic lipase inhibitory activities of Australian native plants. Among the native plants examined, mint bush was found to be the most potent inhibitor with an IC50 of 0.39 mg/ml, followed by Tasmannian pepper and quandong, both of which had the same IC50 of 0.6 mg/ml. These plants have a strong inhibitory activity on pancreatic lipase than the common herb spearmint (IC50, 0.61 mg/ml) and much higher than bay leaf (IC50, 6.3 mg/ml). Sakulnarmrat et al., (2012) also reported that native plants are more potent lipase inhibitors than blueberries.

Table 2.5. Inhibitory properties of Australian native plants on digestive enzymes

1 Common name Enzyme inhibition [IC50 (mg/mL)] References (Scientific name) Pancreatic α-glucosidase α-amylase lipase Anise myrtle 0.3 - 1.55 (Sakulnarmrat et al., (Syzgium 2012) anisatum) Bay leaf2 3.21 - 6.3 (Sakulnarmrat et al., (Laurus Nobilis) 2012) Davidson’s plum 0.13 - 1.74 (Sakulnarmrat et al., (Davidsonia 2012) pruriens) Lemon myrtle 0.13 - 2.51 (Sakulnarmrat et al., (Backhousia 2012) citriodora) Mint bushes 242 - 0.39 (Tang et al., 2017) (Prostanthera rotundifolia) River mint 5.6 54 1.10 (Tang et al., 2015) (Mentha australis) Spearmint2 164 184 0.61 (Tang et al., 2017) (Mentha spicata) Tasmannia 0.83 214 0.60 (Sakulnarmrat et al., pepper 2012) (Tasmannia lanceolata) Quandong 0.39 - 0.60 (Sakulnarmrat et al., (Santalum 2012) acuminatum) Rabbit eye berry 0.097 - 0.94 (Sakulnarmrat et al., (Vaccinium ashei) 2012) Southern 0.091 - 1.02 (Sakulnarmrat et al., highbush 2012) (V.darrowii x V. corymbosum) 1 IC50: half maximum inhibitory concentration. 2These are not native plants, but common herb used as control in the studies. 3-: not determined. 4: activity expressed as % inhibition

2.5.5. Anti-inflammatory properties in Australian native plants

Only a small number of studies have investigated the anti-inflammatory properties of Australian native plants using various in vitro methods (Table 2.6.). Some studies have investigated anti-inflammatory activities by measuring the production of nitrite and prostaglandin E2 (PGE2). For example, Tan et al. (2011) examined the effects of purified extracts of Kakadu plum and Illawarra plum on the production of nitrite and prostaglandin E2 (PGE2) in LPS-activated murine macrophages. Kakadu plum was a much more efficient inhibitor of NO• and prostaglandin E2 (PGE2) production than Illawarra plum. Another study has reported that purified extracts obtained from a number of native Australian herbs (anise myrtle, lemon myrtle, Tasmannia pepper leaf, bay leaf) and fruits (Davidson’s plum, quandong, rabbit eye blueberry, southern highbush blueberry) inhibited the accumulation of nitric oxide (NO•) and release of PGE2 in LPS-activated hepatocellular carcinoma (HepG2) cells (Sakulnarmrat et al., 2012). Lemon myrtle, anise myrtle and quandong were found to be the most efficient inhibitors of NO• accumulation, while Tasmannian pepper leaf and lemon myrtle had the greatest potential to suppress the release of PGE2. However, using a different cell line, Guo et al. (2014) found that bay leaf and anise myrtle were more efficient than Tasmannia pepper leaf and lemon myrtle in preventing the accumulation of NO• accumulation and release of PGE2. Tang et al. (2017) investigated the anti-inflammatory activities of purified extracts of Australian native mints (mint bushes, river mint) using the hyaluronidase inhibitory assay. River mint was found to be a much more efficient inhibitor of the enzyme than mint bush as well as the common spearmint.

Table 2.6. Anti-inflammatory properties of Australian native plants

Anti-inflammatory properties Common name Nitric oxide Prostaglandin Hyaluronidase • References (Scientific name) (NO ) (PGE2) (% inhibition) Concentration production (µM) (pg/ml) Anise myrtle 1 37 (Sakulnarmrat et (Syzgium anisatum) al., 2012) 27 1900 (Guo et al., 2014) Bay leaf 1 10 (Sakulnarmrat et (Laurus Nobilis) al., 2012) 18 2000 (Guo et al., 2014) Lemon myrtle 0.5 8 (Sakulnarmrat et (Backhousia al., 2012) citriodora) 35 3000 (Guo et al., 2014) Mint bushes 6 (Tang et al., (Prostanthera 2017) rotundifolia) River mint 62 (Tang et al., (Mentha australis) 2017) Spearmint 28 (Tang et al., (Mentha spicata) 2017) Tasmannia pepper 2 5 (Sakulnarmrat et (Tasmannia al., 2012) lanceolata) 32 2150 (Guo et al., 2014) Illawarra plum 14 1300 (Tan et al., (Podocarpus elatus) 2011)

Kakadu plum 13 1050 (Tan et al., (Terminalia 2011) ferdinandiana) Davidson’s plum 1.8 100 (Sakulnarmrat et (Davidsonia al., 2012) pruriens) Quandong 1.4 57 (Sakulnarmrat et (Santalum al., 2012) acuminatum) Rabbit eye 2.1 55 (Sakulnarmrat et (Vaccinium ashei) al., 2012) Southern highbush 3 50 (Sakulnarmrat et (V.darrowii x V. al., 2012) corymbosum)

2.5.6. Antimicrobial properties of Australian native plants

Several studies have investigated the antimicrobial activities of native Australian plants although the number of such studies is much smaller than that on the antioxidant activity. Two common methods are used in these studies to assess the antimicrobial activity of the plant extracts: the agar disk diffusion method and the minimum inhibitory concentration method (MIC) (Burt, 2004). The agar disk diffusion method measures the size of the area in an agar plate where the microbial growth is inhibited (known as inhibition area) by the extract while the MIC method determines the minimum concentration of the extract which inhibits the growth of the target organism (Burt, 2004). Dupont et al. (2006) examined extracts of five Australian native plants (lemon iron bark, anise myrtle, strawberry gum, cut leaf and ringwood) obtained with three different solvents (water, ethanol and hexane) for their antimicrobial activity against several Gram positive and Gram negative bacteria. It was found that all the herbs were able to suppress the G+ S. aureus, with anise myrtle demonstrating the best result with a MIC of 7.8 μg/mL, followed by strawberry gum (15.6 μg/mL) and cut leaf (15.6 μg/mL). High inhibitory activities on the G- P. aeruginosa and Sal. Enteritidis were found in lemon iron bark with a MIC of 31.3 μg/mL for both organisms.

Zhao and Agboola (2007) investigated 18 native bush foods for their antimicrobial activities against pathogenic and spoilage bacteria and spoilage yeasts. The native plants were extracted with three different solvents: methanol, water and hexane. The methanol extracts showed much stronger antimicrobial activities than the water and hexane extracts for all the plants. Tasmannia pepper berry and leaf displayed the highest activities. Aniseed myrtle, quandong, eucalyptus oil and wild limes also exhibited high antimicrobial activities. Cock (2008) also reported that the methanolic extract of lemon myrtle had a moderate level of inhibiting activity against bacteria such as Aeromonas hydrophilia (8.3 mm), B. bacillus (7.6 mm), B. subtilis (7.6 mm) and P. fluorescens (7.6 mm). These results were comparable with those of Zhao and Agboola (2007) for the same organisms. Weerakkody et al. (2010) compared the antimicrobial activities of Tasmannian pepper leaf and lemon ironbark with common herbs such as rosemary. The two native plants showed higher activities against L. monocytogenes with an inhibition diameter of 25.6 and 14.4 mm, respectively, followed by S. aureus with 21.8 and 13.2 mm, respectively. Winnett et al. (2014) compared the MIC of native pepper berry, leaf and peppercorn on

S. aureus and B. cereus and found that the berry had the highest activity with MICs of 77 and 93 μg/mL for the two microorganisms, respectively.

2.5.7. Other health related properties of Australian native plants

Australian native plants have also been found to possess other health-related biological properties including anti-cancer and hypertensive activities. Tan et al. (2011) investigated the proapoptotic anticancer effects of purified extract of Kakadu plum and Illawarra plum using the HT-29 cell line. It was found that Kakadu plum induced DNA damage and apoptosis in HT-29 cells. Sakulnarmrat et al. (2012) investigated the anticancer activity of purified phenolic extract of Australian native herbs including anise myrtle, lemon myrtle, Tasmanian pepper leaf, bay leaf, and fruits which were Davidson’s plum, quandong, rabbit eye blueberry and southern high bush blueberry. Several cell lines were used to assess the anti-cancer effects including the human gastric adenocarcinoma (AGS), human bladder transitional cell carcinoma (BL13), human colorectal adenocarcinoma (HT-29) and hepatocellular carcinoma (HepG2) cell lines. All the phenolic extracts demonstrated a decrease in cell activities in all the cancer cell lines. The most effective antiproliferative effect was found in anise myrtle on HepG2 cells with an IC50 value of 0.38 mg/mL. Strong antiproliferative activity was also observed for Tasmania pepper leaf against BL-13 while lemon myrtle showed good antiproliferative activity against all the cancer cell lines.

Sakulnarmrat et al. (2012) also investigated the angiotensin-converting enzyme (ACE) inhibitory activity of purified extracts of Australian native herbs and fruits. The results showed that Davidson’s plum was the most potent inhibitor with 91.3% inhibition, followed by Tasmannia pepper leaf, while the other native plants showed low or no ACE- inhibitory effects.

2.6. Plants investigated in this study

2.6.1. Samphire (Sarcocornia quinqueflora)

Samphire (Sarcocornia quinqueflora) is also commonly known as beaded samphire, bead weed, beaded glasswort or glasswort. The genus Sarcocornia was discovered by Scott (1977) and was distinguished from similar plants such as Arthrocnemum Moq and Salicornia L. on morphological characteristics. Sarcocornia is a perennial plant, with the plants of different sizes staying typically in a horizontal arrangement (Kadereit et al., 2007). Figure 2.4 shows the morphological characteristics and growth forms of Sarcocornia.

Figure 2.4. Morphological characteristics and growth forms of Sarcocornia (Source: Steffen et al., 2015).

The spike-like inflorescence which is a thyrse (A, B, D) with 3–12 strongly decreased flowers per cym (Alonso and Crespo, 2008, Steffen et al., 2010, De la Fuente et al., 2013). (A) Sarcocornia quinqueflora, Yalgorup National Park, Western Australia; (B) S. pacifica, Tomales Bay, California; with a Cuscutagrowing on it; (C) S. littorea, Pearly Beach, Western Cape, South Africa; (D) S. xerophila, Knersvlakte, Western Cape, South Africa (A–D all erect, broomy or spreading shrubs); (E) intensive cushions forming clonal S. pulvinata, Moquegua, Peru; (F) loose carpet-forming clonal S. tegetaria, Langebaan Lagoon, Western Cape; (G) short-lived prostrate herb S. natalensis subsp. affinis,

West Coast, Western Cape, South Africa; (H) loose spreading spider-like clonal S. capensis, West Coast, Western Cape, South Africa.

Plants in the genus often show clonal growth patterns, forming mat-like bushes. The decumbent sections of the plants consist of fleshy, cylindrical internodes and narrow nodes that often carry adventitious roots (C, D, F–H). Species of Sarcocornia found in Australia, their distribution, habitat, growth pattern and number of chromosomes are presented in Table 2.7.

Table 2.7. Australian species of Sarcocornia, their distribution, habitat, growth pattern and number of chromosomes

Australian Sarcocornia

Species S. quinqueflora S. globosa S. blackiana

Distribution Eastern, southern and Western Australia Southern and western western coasts of coasts of Australia, Australia, coasts of coasts of Tasmania, Tasmania, coasts. coasts.

Habitat Intertidal habitats, Elevated edges of Rarely flooded saline supratidal flats, edges saline lakes and habitats of inland salt lakes alluvia and pans

Growth Mat-forming, Erect shrub Erect shrub pattern decumbent subshrub

Chromosome 36 (tetraploid) 18 (diploid) – no. (2n)

No. of 2 1 2 accessions

References Shepherd and Yan Shepherd and Yan Wilson (1980, 1984) (2003), Wilson (2003), Wilson (1980, 1984) (1980, 1984) (Adapted from Wilson, 1980, 1984, Shepherd and Yan, 2003)

In Australia, there are mainly three species of Sarcocornia, namely S. quinqueflora, S. globosa and S. blackiana, which are widely distributed throughout the Australian, Tasmanian and Caledonian coasts (Scott, 1977, Wilson, 1980). Sarcocornia blackiana is found in rarely flooded saline habitats, whereas S. quinqueflora generally grows in intertidal areas, but also has been found at supratidal flats and along the inland pans and salt lakes. The last species, S. globosa, is found mostly in inland areas or on the upper parts of saline lakes and alluvia of Western Australia (Datson, 2002).

Samphire (Sarcocornia quinqueflora) has long been regarded as an edible plant in Australia and there are several books and TV programs that have promoted its consumption (Cribb and Cribb, 1975, Lear and Turner, 1977). Samphire plants collected from coastal wetlands have been used in Australian restaurants although, at the present, it is considered not to be very economical to harvest them in mangroves and saltmarshes in large scale )Saintilan, 2009(. In restaurants, it is considered as a unique and delicious vegetable. Raw samphire can be used in salad if well rinsed with water since it could be too salty to consume directly. Cooking of the plant is also common, which is usually done by stirfrying, simmering in water or in a steamer, like most vegetables. Due to its salty taste, crunchy texture and unique flavour, samphire is also used as an ingredient to be added to dishes alongside the main fish, other seafood and meat dishes. In recent years, samphire has become available in many Australian supermarkets all year round as an exotic vegetable.

Samphire contains about approximately 3.1% of protein, 9.1% of carbohydrate and 0.5% of dietary fibre. It is rather rich in calcium (150 mg/100g) and several other minerals including potassium )50 mg), sodium )870 mg( and iron )1 mg( (Seward, 2019). Low calorie (approximately 100 calories per 100 gram serving) and low fat are other benefits of samphire, and it can be a nutritious and delicious vegetable for regular consumption (Staughton, 2017). Moreover, samphire has been found to contain fucoidans, a group of sulfated polysaccharides generally found in brown seaweeds, which reportedly have many health benefits including anti-inflammatory (Christenses and Brandt, 2006) and antioxidant effects (Holtkamp et al., 2008).

2.6.2. Saltbush (Atriplex nummularia)

Saltbush (Atriplex nummularia), also commonly called oldman saltbush, is a member of the saltbush family. The Atriplex genus contains more than 250 species where most of them are found in the subtropical and temperate regions. Figure 2.5 shows the salt land site and saltbush (Bartel, 2010).

Figure 2.5. Salt land site and saltbush (Source: Bartel, 2010)

In Australia, nearly 61 species of the genus exist. A. nummularia is quite adaptable to the environment and can grow in most soils, but they are more widely found in the semi-arid and arid regions of the Australian mainland on heavy clays and saline soils in the low lying areas with flood plains. Australian saltbushes can grow up to 3 m high and 2-4 m wide with 1-3 cm long leaves which are covered by a scaly coat with a sliver grey colour. The shape of the leaves can vary from elliptical to almost circular, with the variation occurring both in the same plant and in other plants of the species (Wrigley and Fagg, 2003). Saltbush can grow in rainfall zones from 175 mm to 500 mm per year. It is tolerant to frost and drought and is well adapted to the low rainfall conditions which are prevalent in the interior regions of Australia. Rainfalls stimulate their growth, and they can endure intermittent flooding (Richards, 2014). They are also tolerant to acid soil conditions and can survive in highly acid soils with pH<5.

Saltbush has a high protein content (12-16%) and is rich in several minerals and vitamins such as vitamin E (Milton, 2005). Atriplex nummularia is quite edible although currently it is mainly used as stock fodder. The bush is economically significant to the wool industry in several regions of Australian states where saltbushes and bluebushes are required for its survival (QGDAF, 2019). Saltbush is also used to improve the soil conditions of saline sites, which can improve the growth of other plants (Puccinellia, 2006(. For food use, large fresh saltbush or blanched saltbush leaves are used as wrapper for fish, meat or tofu or as a leafy bed in meat or vegetable grills. It is also stir fried with garlic, soy sauce, ginger and other condiments, or used in salads and for battered and tempura cooking style. Dried flakes of saltbush are used as a seasoning for pasta, grills and bread, or added to sea salt as meat seasoning rub or marinade (Jennifer, 2012(. To date, there is no report on the phytochemicals and bioactivities of saltbush (Atriplex nummularia).

2.6.3. Sea parsley (Apium prostratum)

Sea parsley is a native plant of Australia, which can be found abundantly along the southern coastline. It has narrow green leaves similar to the flat leaf of parsley. However, it has a different flavour which is rather similar to celery (Peter, 2006). It can be used similarly like parsley or celery, for instance on salads, for flavoring in cooking and as a garnish (Herbst, 2001). Although an annual plant, sea parsley has a robust tap root similar to carrot, giving it a semi-perennial capacity (Gamble, 2019). The morphology of sea parsley is shown in Figure 2.6. The plant was mentioned as far back as in 1788 in Captain Cook’s landing in Australia where at the time it provided the crew with a welcome flavour for the soups and stews (Grossberg and Fox, 2008).

Figure 2.6. Sea parsley morphology (Source: Lehmuskallio, 2017)

There are a number of studies that have investigated the phytochemicals and health related biological activities of the common vegetable, celery, which belongs to the same Apium genus as sea parsley. These studies have identified an array of phenolic compounds in celery (Sowbhagya et al., 2010) and found that it can provide several significant health functionalities such as antimicrobial (Mišić et al., 2008), anticancer (Sultana et al., 2005) and anti-inflammatory activities (Ovodova et al., 2009), as well as the capacity to lower the level of serum cholesterol (Tsi et al., 2000). However, to date, there has been no report on the study of phenolic compounds and bioactivities of sea parsley (Apium prostratum). Some general information about sea parsley is presented in Table 2.8.

Table 2.8. General information, properties and uses of sea parsley

Common Name: Sea parsley, sea celery. Other Names: Bladder seed, lavose, lovage, love parsley, smallage. Botanical Apium prostratum Genus: Apium Family: Apiaceae Species: Prostratum Native location: Southern coastline of Australia Parts used: Leaves, stems, roots, seeds, oil, rhizones, fruit Properties: A bitter-sweet, sedative herb, pungently aromatic, that induce better digestion, relaxes spasms, increases perspiration, and acts as a diuretic and expectorant. It may have antimicrobial activity against pathogenic organisms. Medicinal uses: Internally for indigestion, colic, gas, poor appetite, kidney stones, cystitis, painful menstruation and slow labour. It can be used to treat aphthous ulcers and sore throat, as well as indigestion, gastric conditions, flatulence, and menstrual problems, and to prevent kidney gravel. Culinary uses: Young shoots and leaf stalks are blanched and eaten as a vegetable stalks are diced. Seeds are added to biscuits, soups, and bread. Leaves are added to savoury dishes, soups, stews, and salads. Dried leaves are mixed with tea. Economic Uses: Oil is used in commercial alcoholic drinks, in perfumery and food flavoring. (Adapted from Brown, 1995, Grossberg and Fox, 2008)

2.7. Conclusion

Phenolic compounds are a huge and diverse group of phytochemicals, which are found extensively in foods derived from plant sources such as fruits, vegetables, whole grains, herbs, spices, green tea and coffee. The potential health effects of phenolic compounds have gained considerable attention from the scientific community in recent decades. Due to its unique climate and ecosystem, Australia is home to a large and diverse collection of plant species, many of them being native to Australia. Native plants in Australia have served as food and medicine for the indigenous people for thousands of years. In recent years, a large number of studies have been conducted to assess the potential health benefits of these plants. These studies have gathered a growing body of evidence about the health benefiting properties of the phenolic compounds in Australian native plants, including antioxidant, antidiabetic, antimicrobial and anticancer properties. However, these studies have only covered a dozen or so Australian native plants, most of which are plants that are commercially cultivated. There are many Australian native plants which have shown potential to be used as vegetables or herbs, and many of which are becoming available in supermarkets and restaurants. There is therefore a need to examine the health beneficial properties of the emerging native plants to prepare them for introduction to the Australian market as a regular food item. In this thesis, three native plants, namely samphire, saltbush and sea parsley, were selected for such studies.

CHAPTER 3 Materials and Methods

3.1. Plant materials

Saltbush (Atriplex nummularia) was purchased from Hello Plants & Garden Supplies, Melbourne, Victoria; sea parsley (Apium prostratum) was purchased from Outback Pride at Cobb&Co Penola Pty Ltd, Penola, South Australia; and samphire (Sarcocornia quinqueflora) from Flowerdale Farm, Melbourne, Australia. The plants were grown locally and harvested in winter. For saltbush and sea parsley, the young shoots and leaf stalks were used for the study; for samphire, the whole plant was used in the study. In all cases, the fresh plant materials were transferred by air freight to our laboratory using an insulated cool box with ice packs, where they were immediately freeze-dried (-109°C, 0.015 kPa; Lab gene ScanVac Coolsafe 110-4 Pro Freeze Dryer, Lynge, Denmark), vacuumed packed (Vacumatic Pty. Ltd. Melbourne, Australia) and kept at -80°C until use. Fresh unfrozen samples and herbarium specimen samples of sea parsley, samphire and saltbush were confirmed as the species by the Royal Botanic Gardens through comparison with the specimens kept at the National Herbarium of New South Wales in September 2017.

3.2. Chemicals and reagents

The chemicals that were used for sample preparation are methanol and Amberlite® resin (XAD-7) which were purchased from Sigma Aldrich (Sydney, Australia).

The following chemicals and reagents were used for antioxidant capacity assays and were purchased from Sigma Aldrich, (Sydney, Australia): Folin-Ciocalteu phenol reagent, ABTS (2, 2’-azino-bis(3ethylbenzothiazoline-6-sulphonic acid) diammonium salt), DPPH (di(phenyl)-(2,4,6trinitrophenyl)-iminoazanium), potassium persulfate, TPTZ (2,4,6- tripyridin-2-yl1,3,5-triazine), sodium acetate trihydrate, iron(III) chloride hexahydrate, trolox and gallic acid.

The following chemicals and reagents were used for enzyme-based assays and were purchased from Sigma Aldrich (Sydney, Australia): acarbose, orlistat, 4-dimethylaminobenzaldehyde (DMAB), dimethyl sulfoxide (DMSO), dinitrosalicylic acid, α-glucosidase, α-amylase, hyaluronidase, hyaluronic acid, pancreatic lipase, 4-methylumbelliferyl oleate, p-nitrophenyl- β-ᴅ-glucopyranoside (P-NPG) and sodium potassium tartrate tetrahydrate.

The following chemicals and reagents were used for high performance liquid chromatography (HPLC) and liquid chromatography-mass spectrometry (LC-MS) analysis: acetonitrile (HPLC grade) was purchased from Sigma-Aldrich Corporation (Sydney, Australia); methanol (HPLC grade) from Burdick & Jackson (Muskegon, MI, USA); glacial acetic acid from Ajax Finechem Pty. Ltd. (Sydney, Australia). Reference standards of phenolic compounds used were: caffeic acid, catechin, catechin hydrate, ferulic acid, chlorogenic acid, ρ-coumaric acid, sinapic acid and syringic acid, which were purchased from Sigma Aldrich (Sydney, Australia); apigenin, hesperetin, naringin, naringenin, quercetin, luteolin and apiin were purchased from Aladdin Chemistry Co. Ltd (Shanghai, China). Milli-Q water (Millipore, Sydney, Australia) was used in all the experiments.

3.3. Extraction of phenolic compounds

3.3.1. Preparation of crude phenolic extracts

Samples of freeze-dried plant materials were first crushed with a rotor mill (Pulverisette 14, Fritsch GmbH, Idar-Oberstein, Germany) to pass 0.5 mm mesh. After that, 10 g of sample was extracted with 80% methanol (v/v) at a 1:20 g/mL ratio for 10 min in a sonicated water bath, and centrifuged at 3000g for 10 min. The pellet was extracted two more times under the same conditions, and the supernatants were pooled. The supernatant was filtered through a 0.45 μm polytetrafluoroethylene (PTFE) membrane to obtain the crude extract, which was kept at -20°C until used for analysis. Antioxidant activity assays of the extracts were usually conducted within 3 days after they were prepared. For enzyme inhibition studies, 50 mL of the crude extracts were first concentrated for 30 min by vacuum rotary evaporation (37°C; Orme Scientific Ltd., Manchester, UK), and then freeze-dried and kept at -20°C. Freeze-dried plant samples were used within two weeks. The yield of the crude extracts was calculated as following:

Weight of crude extract (g) Crude yield (%) = × 100 Weight of sample (g)

3.3.2. Preparation of purified extracts

Freeze-dried samples (5 g) of the three plants were extracted 3 times, each time with 100 mL of 80% (v/v) methanol. For the first extraction, the plant and solvent mixture was sonicated for 10 min, then stirred with a magnetic stirrer for 2 h at 4°C, and finally centrifuged at 10,000 g for 15 min (Thermo Scientific Fiberlite, Sydney, Australia). The second and third extractions were performed with magnetic stirring overnight at 4°C. The supernatants were pooled and subjected to vacuum rotary evaporation (40°C; Orme Scientific Ltd., Manchester, UK) to remove the methanol solvent. Undissolved residual material in the remaining aqueous extract was removed by centrifugation at 3,500 g for 30 min.

The purification of the aqueous extract was carried out following the procedure of Konczak and Sakulnarmrat (2012) using the non-ionic aliphatic acrylic polymer adsorbent, Amberlite® XAD-7 resin. Amberlite® resin adsorbs polar substances from non-polar solvents, but non-polar substances from aqueous solutions. The resin was packed in a 300 x 25 mm liquid chromatography (LC) column. Aliquot (50 mL) of the aqueous extract, obtained as described above, was loaded onto the column and prewashed with Milli-Q water until the elute became colourless, which indicates that the impurities in the sample had been mostly removed. The column was then eluded with 80% ) v/v) methanol to obtain the purified phenolic extract. Afterwards, methanol was eliminated via vacuum rotary evaporation at 40°C, and the remaining aqueous extract was lyophilized (-109°C, 0.015 kPa; Labogene ScanVac Coolsafe 110-4 Pro Freeze Dryer, Lynge, Denmark). The freeze dried powder was stored at -20°C and homogenised before bioassays and chemical analysis. The yield of the purified extract was calculated as follows:

Weight of purified extract (g) Purification yield (%) = × 100 Weight of sample (g)

3.4. Proximate analysis 3.4.1 Moisture content

The moisture content of the plant material was determined by the AACC method (44-01, 2000). The moisture content of fresh plant samples, which was packed by modified atmosphere packaging and maintained in refrigerator at 3 °C, was determined within three days after arrival. Before the analysis, aluminum dishes were dried and kept in a desiccator. Two duplicate plant samples (5 g) were put into the pre-weighed dishes, which were placed in a preheated vacuum drying oven (Townson & Mercer Ltd, Croydon, England) set at 70°C until the weight became constant. The moisture content on a wet basis (MCwb) was calculated as follows:

Wi − Wf MCwb(%) = × 100 Wi

Where, Wi = weight of initial sample; Wf = weight of final sample

3.4.2. Ash content

The ash content was determined according to the approved AACC method (08-01, 2000(. Before the analysis, aluminium dishes were dried and kept in a desiccator. Fresh plant samples (2 g) were transferred to the dishes and put in the furnace for 5 h at 520°C. The content of ash was estimated as follows:

Wi − Wf Ash (%) = × 100 Wi

Where, Wi = weight of initial sample; Wf = weight of final sample.

3.4.3. Lipid analysis

The content of crude lipid was determined via a Soxhlet system using the approved AACC (30-25, 2000( method. Two duplicate samples of freeze-dried plants (2 g) were extracted with diethyl ether at ambient temperature for 8 h. After that, the solvent extracts were placed in an oven at 105°C until constant weight was reached. The content of crude lipid was estimated based on the percentage of residue per gram of sample weight.

Weight of extract Crude lipid (%) = × 100 Weight of sample

3.4.4. Protein content

The content of protein was determined by a LECO TruSpec CN Analyser (LECO Corporation, St Joseph, MI, USA) based on nitrogen content in the sample, using the approved AACC (46-30, 2000( method (LECO, 2006(. Aliquots (0.2 g) of lyophilized plant samples was filled into the loading head, and the atmospheric gases were purged. The device was set in the furnace at 950°C and flushed rapidly with oxygen, followed by further combustion at 850°C. A thermal conductivity cell was used to determine the nitrogen content (N %). Protein content of the plant sample was determined from the amount of nitrogen, with a conversion factor of 5 to calculate the protein content (Yeoh and Wee, 1994). The analysis was performed in duplicates.

3.4.5. Carbohydrate content

The content of carbohydrate was obtained by difference, which was calculated as follows:

Carbohydrate (%) = 100 – [Moisture (%) + Protein (%) + Lipid (%) + Ash (%)]

3.5. Assay of total phenolic content and antioxidant capacity

3.5.1. Preparation of plant extract solutions

The lyophilized crude extract, obtained as described above, was dissolved (40 mg/ml) in phosphate buffer (75 mM, pH 7.2(. The lyophilized purified extract was first dissolved in the same phosphate buffer (1 mg/mL) and then diluted with the same buffer at the ratio of 1:25. These plant extract solutions were used for all the following assays for total phenolic content and antioxidant capacity.

3.5.2. Total phenolic content

The Folin-Ciocalteu assay described by Dorman et al. (2003( were used, with minor modifications, to determine the amount of total phenolic content in plant extracts. Aliquots (30 µL) of the plant extract solutions, prepared as described in section 3.5.1, were mixed with Folin-Ciocalteu phenol reagent (140 µL), left at ambient temperature for 5 min, and placed onto a 96-well microplate (Sarstedt Australia, Technology Park, SA, Australia). A 20% sodium carbonate solution (30 µL) was added to the mixture which was shaken for 5 s to terminate the reaction. After that, the mixture was left at room temperature for 2 h and the absorbance was measured at 765 nm with a SpectraMax M2 microplate reader (Molecular Device, Sunnyvale, CA, USA) against the phosphate buffer blank. A reagent control was analysed in the same way, but the phosphate buffer was used to replace sample extracts. Gallic acid at the concentration range of 1-100 mg/mL in the same phosphate buffer were analysed exactly the same way to create a standard curve (Appendix 1(, which was used to estimate the total phenolic content in the sample, which was expressed milligram gallic acid equivalent (GE) per milligram of plant extract material (mg GE/mg). The assay was repeated at least five times (n = 5) for each extract.

3.5.3. Ferric Reducing Antioxidant Power (FRAP) assay

FRAP assay followed the method of Konczak et al. (2010b) with minor modifications. The FRAP reagent prepared by mixing 20 mL of acetate buffer (300 mM, pH 3.6) 2 mL of TPTZ (10 mM) in hydrochloric acid and 2 mL of iron (III) chloride hexahydrate (20 mM). Aliquotes (20 μL) of the extract solutions, prepared as described in section 3.5.1,

60 were mixed with the FRAP reagent (150 μL) and the mixtures were left in darkness for 8 min. The absorbance at 600 nm was then measured with a SpectraMax M2 microplate reader (Molecular Device, Sunnyvale, CA, USA) against a blank of Milli-Q water. The reagent blank was assayed in the same way except Milli-Q water was used instead of the extract solution. Trolox solutions (25-1000 μM) were assayed in the same way to construct a standard curve and FRAP values of crude and purified extracts were expressed as mM Trolox equivalent per milligram of dry matter of the plant extract (µM TE/mg). The FRAP assay was repeated at least five times (n = 5) for each extract.

3.5.4. DPPH radical scavenging capacity assay

The DPPH (di (phenyl)-(2,4,6-trinitrophenyl) iminoazanium) radical scavenging capacity assay was carried out according to the procedure described by Danh et al. (2012( with some minor modifications. The plant extracts (40 μL), prepared as described in section 3.5.1, were mixed with DPPH methanolic solution (160 μL, 0.2 mM), and the mixture was left at room temperature for 30 min under dark conditions. Absorbance at 517 nm was then measured via a SpectraMax M2 microplate reader (Molecular Device, Sunnyvale, CA, USA) against a phosphate buffer blank. The reagent blank was assayed in the same way except Milli-Q water was used instead of the extract solution. Gallic acid solutions at the concentration range of 1-100 mg/mL in the same phosphate buffer were analysed exactly the same way to create a standard curve. DPPH radical scavenging activity of crude and purified extracts was referred as gallic acid micromolar equivalent (GE) per milligram of dry extract (μM GE/mg). The DPPH radical scavenging assay was repeated at least five times (n = 5).

3.5.5. ABTS radical scavenging capacity assay

The ABTS (2, 2’-azino-bis (3ethylbenzothiazoline-6-sulphonic acid) diammonium salt) radical cation scavenging capacity assay was performed following the method of Danh et al. (2012). ABTS stock solution was prepared by mixing 19.2 mg ABTS powder with 3.5 mL of Milli-Q water, followed by the addition of 11.04 mg potassium persulfate powder and another 20 mL of Milli-Q water. The mixture was left for 12-16 h at room temperature without any light. Working solution of ATBS was prepared by mixing 1 mL of the ABTS 61 stock solution with 39 mL Milli-Q water just before analysis. For the assay, the plant extracts (10 μL), prepared as described in section 3.5.1, and were transferred to a 96-well microplate, followed by the addition of 190 μL of the ABTS working solution. The mixtures were shaken for 5 s, and then left in darkness for 5 min. The absorbance of mixtures at 734 nm was measured on a SpectraMax M2 microplate reader (Molecular Device, Sunnyvale, CA, USA) against a blank of phosphate buffer. The reagent control was assayed in the same way except that Milli-Q water was used instead of the extract solution. Trolox solutions at the concentration range of 1-100 mg/mL in the same phosphate buffer were analysed in exactly the same way to create a standard curve. ABTS radical scavenging activity of crude and purified extracts was referred to as micromolar Trolox equivalent per milligram of dry extract (µM TE/mg). This assay was repeated at least five times (n = 5).

3.6. Assays of enzyme inhibitory activities

3.6.1. Preparation of plant extract solutions

Solutions of the lyophilized crude and purified extract at the concentration range of 0.125- 20 mg/mL were prepared by dissolving appropriate amount of the extract in phosphate buffer (75 mM, pH 7.2(. These plant extract solutions were used for all the following assays of enzyme inhibitory activities.

3.6.2. Hyaluronidase inhibitory activity assay

The assay of hyaluronidase inhibitory activity was conducted according to Muckenschnabel et al. )1998( with some minor modifications. Lyophilized crude and purified extracts were dissolved in DMSO in a range of concentrations (0.125-20 mg/mL). Aliquots (13.5 µL) of the extract solutions were added with citrate-phosphate buffer (150 µL, 100 mM, pH 5(, bovine serum albumin solution (37.5 µL, 0.2 g/L), hyaluronic acid (37.5 µL) and Milli-Q water (61.5 µL). After addition of hyaluronidase enzyme (37.5 µL, 100 U/mL), the reaction mixture was incubated at 37°C for 1 h. An alkaline solution was used as the terminating agent, which was prepared by mixing 1 mL of a potassium carbonate solution

(8 g K2CO3 in 10 mL Milli-Q water) with 10 mL of a borate solution (17.3 g H3BO3 and

7.8 g KOH in 100 mL Milli-Q water). The solution was heated for 5 min, cooled down and 90 µL of which was transferred to each well of the microplate to terminate the enzymatic reaction. After that, 110 µL of 4-dimethylaminobenzaldehyde (DMAB) was added and the mixture incubated at 37°C for 20 min. The DMAB solution was prepared by dissolving DMAB (20 g) in HCl (25 mL) and glacial acetic acid (75 mL). The solution was diluted with four volumes of glacial acetic acid before use and the absorbance at 590 nm was measured by a SpectraMax M2 microplate reader (Molecular Device, Sunnyvale, CA, USA) against an enzyme blank. The control solution was assayed in the same way except that DMSO was used instead of the extract. This experiment was repeated five times (n = 5). The hyaluronidase inhibition activity was estimated as follows:

A − A Inhibition (%) = S SB × 100 AC − ACB

Where, AC = absorbance of control, ACB = absorbance of control blank, AS = absorbance of sample, ASB = absorbance of sample blank

3.6.3. Pancreatic lipase inhibitory assay

The pancreatic lipase inhibitory assay was performed according to the method of Sakulnarmrat Konczak (2012( with minor modifications. The lipase solution was prepared by mixing lipase (0.085 g) with 1 mL of McIlvaine’s buffer (pH 7.4). The solution was centrifuged at 10,000 g for 10 min, and the supernatant was collected. Solutions of lyophilized crude and purified extracts at the concentration range of 0.125-20 mg/mL were prepared with McIlvaine’s buffer containing disodium phosphate (0.2 M) and citric acid (0.1 M) with the pH adjusted to 7.4. Orlistat (0.125-20 mg/mL) was prepared in dimethyl sulfoxide (DMSO). For the assay, sample extracts (50 µL) were added to 100 µL of 4- methylumbelliferyl oleate (0.1 mM, in DMSO), followed by the pancreatic lipase enzyme (50 µL, 0.085 g/mL), and the mixture was left at 37 °C for 20 min. After that, 2 mL of 0.1 M sodium citrate and 1 mL of 0.1 N hydrochloric acid were added to terminate the reaction. The fluorescence absorbance of the mixture was measured at the emission wavelength of 460 nm and excitation wavelength of 320 nm by a Varian Cary Eclipse fluorescence reader (Agilent Technologies Inc., Sydney, Australia) against a buffer blank.

The control solution was assayed in the same way except that the buffer solution was used instead of the extract. This assay was repeated five times (n=5). The lipase inhibition was expressed as % of inhibition and determined using the following equation:

(F − F ) − (F − F ) Inhibition (%) = 1 − C CB S SB × 100 (FC − FCB)

Where, FC = Fluorescence of control, FCB = Fluorescence of control blank, FS = Fluorescence of sample, FSB = Fluorescence of sample blank.

3.6.4. α-Amylase inhibitory activity assay

The assay of α-amylase inhibition was performed following the method of Namboothiri et al. (2011) with some minor modifications. The dinitrosalicylic acid colour reagent was prepared by dissolving dinitrosalicylic acid (472.6 mg) in Milli-Q water (20 mL), followed by the addition of 12 mL of 2 M sodium hydroxide, 12 g sodium potassium tartrate tetrahydrate and Milli-Q water (12 mL). The mixture was heated in a shaking water bath overnight in darkness to dissolve all the constituents. Lyophilized crude and purified and extract solutions (0.125-20 mg/mL) were prepared with sodium phosphate buffer (0.02 M, pH 6.9 with 0.006 M sodium chloride). Starch solution (100 µL, 1% v/v in the same buffer) was mixed with the extract solution (100 µL) and the mixture was kept at 25°C for 10 min. After that, α-amylase (100 µL, 0.5 mg/mL, 200 U) was added and left at 25°C for a further 10 min. Dinitrosalicylic acid colour reagent (200 µL) was then added and the mixture was incubated at 100°C for 5 min to terminate the reaction. The mixture was left at room temperature until it was cooled down, and 50 µL of which was transferred to a microplate and mixed with 200 µL Milli-Q water. The absorbance at 540 nm was then measured with a SpectraMax M2 microplate reader (Molecular Device, Sunnyvale, CA, USA) versus a reagent blank (no enzyme). The control was assayed in the same way except that the phosphate buffer was used instead of sample extracts. Acarbose solutions (0.125-20 mg/mL) were assayed in the same way to produce a standard curve. This assay was repeated five times (n = 5). The α-amylase inhibition activity was expressed as percentage inhibition using the following equation:

(A − A ) − (A − A ) Inhibition (%) = C CB S SB × 100 AC − ACB

Where, AC = absorbance of control, ACB = absorbance of control blank, AS = absorbance of sample, ASB = absorbance of sample blank.

The inhibition activity was determined for 10 different concentrations of each extract between 0.125 and 20 mg/mL. An inhibition v. extraction curve was constructed and the half inhibition concentration (IC50) was calculated from the curve for each extract.

3.6.5. α-Glucosidase inhibitory activity assay

The assay of α-glucosidase inhibition was performed following the method of Zhang et al. (2013(. All solutions were prepared at ambient temperature. Lyophilized crude and purified extracts (0.125-20 mg/mL) were prepared in sodium phosphate buffer (100 mM, pH 6.9(. α-Glucosidase (30 µL, 0.1 U/mL) was mixed with the extracts (30 µL) in a 96- well microplate. The mixture was maintained at 37°C for 10 min, and the substrate, p- nitrophenyl-β-D-glucopyranoside (P-NPG) (60 µL, 0.375 mM) was added. The mixture was incubated at 37°C for 30 min and Na2CO3 (120 µL, 0.2 M) was added to terminate the reaction. The absorbance at 405 nm was measured on a Spectra Max M2 microplate reader (Molecular Device, Sunnyvale, CA, USA) against an enzyme blank (no enzyme was used). The control was assayed in the same way except that the phosphate buffer was used instead of sample extracts. Acarbose solutions (0.125-20 mg/mL) were assayed in the same way to produce a standard curve. This assay was repeated five times (n = 5). The α-glucosidase inhibitory activity was expressed as percentage inhibition using the following equation:

(A − A ) − (A − A ) Inhibition (%) = C CB S SB × 100 AC − ACB

Where, AC = absorbance of control, ACB = absorbance of control blank, AS = absorbance of sample, ASB = absorbance of sample blank.

3.7. Identification of and quantification of phenolic compounds

3.7.1. High Performance Liquid Chromatography-Photodiode Array Detector (HPLC- DAD) analysis

HPLC-DAD analysis was conducted on a Shimadzu Prominence Ultra-Fast Liquid Chromatography (UFLC) system (Shimadzu, Japan), equipped with a SIL-20A HT autosampler, a DGU-20A5 degasser, two LC-20AD pump, a CTO-20A column oven, a SPD-M20A photodiode array detector, a CBM-20A bus module and a FC-10A fraction collector. The column used was a Phenomenex (Sydney, Australia) Luna 5 µm C18 )2( (250 mm x 4 mm i.d.), which was kept at 30°C during analysis. The HPLC procedure was based on Wang et al. (2004(. The gradient elution system consisted of 2.5% v/v aqueous acetic acid (solvent A) and 100% acetonitrile (solvent B). All solvents were filtered by 0.45 µm PTFE membrane (Grace Davidson Discovery Sciences, Melbourne, Australia) before use. The gradient program was as follows: 15% B (0 min), 15% B (6 min), 29% B (8.5 min), 31.5% B (17.5 min), 90% B (19 min), 90% B (21.5 min), 15% B (27 min), 15% B (34 min). The flow rate was 0.8 mL min-1. The sample solution was prepared by dissolving lyophilised purified extracts (1 mg) in 1 mL methanol (HPLC grade) and 20 µL of sample solution was injected into the HPLC system using the autosampler. The detection wavelength used was 280nm for hydroxybenzoic acids and flavanols, 320nm for hydroxycinnamic acids and 370 nm for flavonols. The analysis was repeated at least five times for each sample (n = 5(.

3.7.2. Quantification of phenolic compounds by HPLC-PDA

Quantification of phenolic components was carried out by HPLC-PDA using the external standard method where a standard curve was constructed for each compound using the reference standards. Good linearity (r2 > 0.99) was obtained for all the standards. Quantification run for each phenolic compound was independently repeated at least three times (n = 3). 66

3.7.3. Liquid Chromatography-High Resolution Mass Spectrometry analysis

LC-HRMS analysis was carried out on an Orbitrap LTQ XL instrument (Thermo Fisher System, San Jose, CA), equipped with an Accela pump. The column was an Acquity uplc C18 (150 mm x 2.1 mm i.d, 1.7 µm, Waters XBridge, Sydney, Australia). The mobile phases were 5 mM ammonium formate in water, pH 7.4, adjusted with ammonium hydroxide (solvent A) and 5 mM ammonium formate in 90% methanol, pH 7.4 (solvent B). The solvents were filtered through a 0.45 µm PTFE membrane (Grace Davidson Discovery Sciences, Melbourne, Australia) before use. The gradient program was as follows: 10% B (0 min), 30% B (2.5 min), 30% B (4.5 min), 45% B (5.5 min), 45% B (7 min), 75% B (12 min), 90% B (14 min), 20% B (14.1 min), 10% B (19 min). The flow rate was 200 µL/min-1. The LC-HRMS conditions used were as follows: HESI and APCI sources were used in negative mode to generate ions from the analytes in solution. Data was acquired in Q3 full scan mode and the product scans were performed on analyte ions of purified extracts and reference standards. The injection volume for the extract samples was 20 µL and each analysis was repeated at least three times. Ions were measured over a mass range of m/z 120-950 over 20 min (full scan mode).

The instrument of LC-HRMS was optimised for sensitivity on both solvent and compound using LC tuning software. The Qual Browser on Xcalibur 2.2 software (Thermo Fisher Scientific, San Jose, CA) was used for analysis of mass spectrometry data. The LOD for LC- HRMS was 0.625 ng of apigenin. This experiment was independently repeated at least three times (n = 3(.

3.8. Statistical Analysis

Each experiment was independently repeated at least twice and each assay was typically replicated at least three times except the proximate analysis where the analysis was replicated two times. Means and standard deviations were calculated based on all the independent measurements. One-way ANOVA was used to compare differences between sample treatments and Tukey’s test was used to identify the differences between the means. The Pearson’s correlation coefficient was used to analyse correlation. Differences were considered to be significant when p < 0.05. All statistical analysis was performed with the Graphpad Prism 6 (Graphpad Software, CA, USA) software. 67

CHAPTER 4 Phenolic Composition and Antioxidant Activities of Saltbush, Samphire and Sea Parsley

4.1. Introduction

Phytochemicals such as phenolic compounds have received a lot attention over the last few decades because of their potential health benefits (Balasundram et al., 2006). Alongside this general rush in the study of phytochemicals, the phenolic compounds in Australian native plants have also gained considerable interests and a number of Australian native fruits, vegetables, herbs and spices have been investigated for this purpose. However, as mentioned in Chapter 2, Australia is home to over 24,000 species of plants and over 5000 of which are edible or can be used for medicinal purposes (Cooper, 2004), but previous research has covered only a fraction of them. Meanwhile, there is an increasing number of the native fruits, vegetables, herbs and spices being launched into Australian special shops and supermarkets or become available in restaurants, not only for their exotic and unique flavour but also for their perceived health benefits and therapeutic properties (Sakulnarmrat et al., 2012). Clearly, more research is needed to investigate phenolic compounds in these Australian native plants in order to establish the scientific foundation of their health beneficial properties and therapeutic functions. In this thesis, three native plants were selected for investigation, namely saltbush (Atriplex nummularia), samphire (Sarcocornia blackiana) and sea parsley (Apium prostratum).

Investigation of phenolic compounds usually starts with extraction of these compounds, followed by determination of the total phenolic content and antioxidant activities of the extracts, and this is the general approach used in this chapter. Because the phenolic extracts usually contain a large proportion of non-phenolic impurities such as carbohydrates, proteins, lipids and pigments which are often co-extracted with the phenolic components (Tang et al., 2006), a purification step is included to remove these impurities. The objectives of this chapter were to determine the general phenolic content of saltbush, samphire and sea parsley using the Folin-Ciocalteu procedure, and to evaluate the antioxidant capacities of the three plant samples using three different methods, namely ABTS (2,2'-azino-bis(3-ethylbenzothiazoline-6-sulphonic acid) radical cation scavenging

68 capacity assay, DPPH (di(phenyl)-(2,4,6-trinitrophenyl)iminoazanium) radical scavenging assay and ferric reducing antioxidant power (FRAP) assay.

4.2. Results and discussion

4.2.1. Proximate composition of saltbush, samphire and sea parsley

Results of proximate analysis for moisture, protein, crude lipid, ash and carbohydrate of Australian native saltbush, samphire and sea parsley are given in Table 4.1. Proximate analysis of these three plants have not been reported before. Sea parsley showed the highest moisture content (86.2%) followed by samphire (84.7%) and saltbush (76.9%) and the same trend was observed for the level of crude lipid in the plants. Saltbush had the highest level of protein content (4.2%), followed by sea parsley (3.3%) and samphire (2.8%) while samphire had the highest ash content at 5.9%, compared to 5.4% and 4.3% for saltbush and sea parsley respectively and carbohydrate content.

Table 4.1. Proximate composition of Australian native saltbush, samphire and sea parsley (w/w, fresh weight1)

Plant species Moisture Protein Crude Lipid Ash Carbohydrate2 (%( (%( (%( (%( (%( Saltbush 76.9 ± 2.54 4.2 ± 0.26 1.5 ± 0.11 5.4 ± 0.14 12.0 ± 0.12 Sea parsley 86.2 ± 1.95 3.3 ± 0.14 2.1 ± 0.22 4.3 ± 0.35 4.10 ± 0.24 Samphire 84.7 ± 3.15 2.8 ± 0.21 0.6 ± 0.23 5.9 ± 0.43 6.00 ± 0.21 1Protein conversion factor used was 5 (Yeoh and Wee, 1994). Moisture (n = 3); all other data (n = 2). 2Carbohydrate was calculated by difference: % carbohydrate = [100 – (moisture (%) + protein (%) + lipid (%) + ash (%))].

4.2.2. Yield of phenolic extracts from saltbush, samphire and sea parsley

Freeze-dried leaves of the three native plants were ground to powder and extracted with 80% (v/v) aqueous methanol three times to obtain the crude extracts, which were purified by liquid chromatograph using Amberlite® resin (XAD-7). The yield of crude and purified extracts of the three plants are given in Table 4.2. Overall, samphire produced the highest crude yield (12.05% DW), followed by sea parsley (9.69% DW) and saltbush

(9.14% DW), which gave similar yields. Samphire also gave the highest yield of purified extract at 6.21%, which was slightly over half of the crude yield for this plant. Sea parsley and saltbush had similar yields of purified extracts at 3.94% and 3.41% respectively, which were about 40% of their crude yields. Similar results have been reported for other Australian native plants. For example, Tang et al. (2016) reported crude yields of 12.2%, 10.6% and 10.1% for mint bush, river mint and spearmint, respectively. The purified yields for the three plants were 6.7% for mint bush, 5.8% for spearmint and 4.3% for river mint, where the purified yields were about 42-54% of the crude yields. Similarly, Konczak and Roulle (2011) reported purified yields of 5.74% for lemon myrtle, 4.93% for anise myrtle and 5.24% for bay leaf. The impurities in the crude extracts include soluble proteins, carbohydrates, lipids and pigments, which are removed by the purification process using liquid chromatography with Amberlite® resin (XAD-7) (Dai and Mumper, 2010, Tang et al., 2016).

Table 4.2. Yield of crude and purified extracts from native Australian saltbush, samphire and sea parsley (%(, DW1

Plant species Crude Purified (%( (%( Samphire 12.05 ± 0.48a 6.21 ± 0.16a Saltbush 9.14 ± 0.12c 3.41 ± 0.04c Sea Parsley 9.69 ± 0.41b 3.94 ± 0.05b 1Yield was expressed as percent of the dry plant matter (%, DW). The data represent mean ± standard deviation of at least 5 independent experiments.

4.2.3. General discussion on the extraction and purification of saltbush, samphire and sea parsley

Researchers have used a variety of different solvents to extract phenolic compounds from plants, including polar solvents such as water, methanol, ethanol and acetone at various concentrations and, less commonly, non-polar solvents such as hexane, as well as combinations of these solvents (Obermeyer et al, 1995, Mazur and Adlercreutz, 1998, Alhakmani et al., 2013, Shahidi and Naczk, 2004, Wong-Paz et al., 2015, Tang et al., 2016). The yield of phenolic compounds can be strongly affected by the selection of solvents as studies have found that different types of phenolic compounds may be preferentially 70 extracted by different solvent. For example, acetone has been shown to extract higher amounts of flavonols, whereas methanol is more efficient in extracting polyphenols overall (Dai and Mumper, 2010). Among the solvents, aqueous methanol has been one of the most commonly used for extracting phenolic compounds from plants including Australian native plants. Zhao and Agboola (2007) investigated the efficiency of various solvents, including hexane, methanol and water, for extracting phenolic compounds from a number of Australian native plants, and found that methanol was the most efficient for this purpose. In our laboratory, 80% (v/v) has been found to be generally satisfactory for the extraction of phenolic components from Australian native plants (Tang, et al, 2016, 2017), and therefore this solvent was also used in the present study.

Apart from phenolic compounds, other substances including carbohydrates, lipids, vitamins and pigments can also be extracted by aqueous methanol. The presence of these components can significantly interfere with subsequent analyses such as identification of the phenolic compounds by HPLC and LC-MS. Therefore, purification of crude phenolic extracts is an essential step prior to the characterization of the chemical components and bioactivities of phenolic compounds in the extracts (Dai and Mumper, 2010, Tang et al., 2016). Liquid chromatography is one of the most commonly used methods for the purification of phenolic compounds (Pyrzynska and Biesaga, 2009, Dai and Mumper, 2010). This method employs adsorbent materials such as silica-based resins (e.g., C18 resin) or polymeric adsorbents (e.g., Amberlite® XAD-7 and XAD-16), which have different affinities to compounds based on their polarities. Non-polar compounds such as carbohydrates, lipids, vitamins and pigments can be easily removed due to their low affinities with the resins (Sakulnarmrat and Konczak. 2012). This technique is simple, fast and relatively inexpensive, and therefore has been used extensively for the purification of phenolic compounds (Aehle et al., 2004). Both XAD-7 (Tan et al., 2011a, 2011b, 2011c, Tang et al., 2016, 2017) and XAD-16 (Sakulnarmrat and Konczak, 2012, Sakulnarmrat et al., 2013, Guo et al., 2014) have been used to purify phenolics in Australia native plants. For these reasons, this technique was also used for this purpose in the present study.

Extreme care was exercised during the extraction and purification process to minimize potential damage to phenolic compounds that may occur due to oxidation. The entire extraction process was carried out at 4°C and the extract samples were stored at -20°C and in darkness. The purified extracts were purged by nitrogen gas before the storage. 71

Samples were usually analyzed within 14 days, and for samples that were to be stored longer than 14days, they were vacuum-packed and stored at -80°C.

4.2.4. Phenolic composition of Australian native saltbush, samphire and sea parsley

Total phenolic content of crude and purified extracts of saltbush, samphire and sea parsley samples was determined using the Folin-Ciocalteu procedure )Table 4.3(. Of the three plants, sea parsley was found to always have the highest total phenolic content in both the crude (154.59±7.64 mg GAE/g dry weight of the extract, the same in the following sections) and purified extracts (362.28±18.57 mg GAE/g). However, the data trends were slightly more complex for the other two plant samples. For the crude extracts, samphire had a greater yield (88.94 ± 3.15 mg GAE/g) than saltbush (79.65 ± 2.48 mg GAE/g), but the reverse was true for the purified extracts (127.33 ±11.14 and 212.79 ± 18.24 mg GAE/g for samphire and saltbush, respectively). Purification resulted in marked increases in phenolic content for all three plant samples. For saltbush and sea parsley, the phenolic contents in the purified extracts were more than 2-fold higher than the respective crude extract, for samphire, the phenolic content in the purified extract was 1.43 times greater than the crude extract. These results are expected as the purification step was aimed at removing non-phenolic components which were extracted alongside the phenolic compounds. As shown by previous researchers, the non-phenolic components were likely composed of sugars, organic acids, proteins and fats which are soluble in aqueous methanol (Dai and Mumper, 2010, Oszmiański et al., 2011). The greater increases in the phenolic content in purified extracts of saltbush and sea parsley than that of samphire suggest that saltbush and sea parsley may have a greater amount of non-phenolic impurities in the crude extract. This agrees with the results in Table 4.2., where it is shown that yield differences between the purified and crude yields were much greater for saltbush and sea parsley than for samphire.

Table 4.3. Total phenolic content of crude and purified extracts of saltbush, samphire and sea parsley samples in dry weight of the extract (mg GAE /g, Extract)

Plant Crude Purified Increase of purified species (mg GAE /g, Extract) (mg GAE /g, Extract) over crude extract (times) Samphire 88.94 ± 3.15b 127.33 ± 11.14b 1.43 Saltbush 79.65 ± 2.48c 212.79 ± 18.24c 2.67 Sea Parsley 154.59 ± 7.64a 362.28 ± 18.57a 2.34 The data represent the mean ± standard deviation of at least 5 independent experiments. Means with different letters in the same column were significantly different with p < 0.05 (n = 5).

Priecina and Karklina (2015) reported a total phenolic content of 441.76 ± 13.86 mg GAE/100g DW for the extract of celery roots, which is higher than that for sea parsley which belongs to the same genus. A total phenolic contents of 10.12 mg GAE/g DW has been reported for the methanolic extracts of Atriplex halimus, which is considerably lower than that of saltbush which belongs to same genus (Benhammou et al., 2009). A number of factors could influence the phenolic content of a plant, including genotypes within a species, environmental factors such climatic and geographical conditions, different parts of the plant tested, time of sample collection, preparation of samples, drying conditions, and solvents and methods used for extraction (Shan et al., 2005, Mimica-Dukic and Bozin, 2008). Furthermore, the methods used for measuring the total phenolic content could have also a significant impact on the results (Şahin et al., 2012). These factors can potentially lead to large differences in phenolic content in the similar plant species, and this may partly explain the differences between the results of the present study and those reported for plants in the same genus.

4.2.5. Antioxidant capacity of saltbush, samphire and sea parsley

4.2.5.1. ABTS free radical scavenging capacity

ABTS radical cation scavenging capacity is one of the most commonly used methods for measuring antioxidant activities of phytochemicals. The assay is based on the capacity of the phytochemical to transfer electron or hydrogen to ABTS radical cation with the consequent formation of a stable compound (MacDonald‐Wicks et al., 2006). In this

73 study, ABTS free radical scavenging activity assay was used to assess the antioxidant activity of Australian native saltbush, samphire and sea parsley samples (Table 4.4). Overall, sea parsley showed considerably higher ABTS radical cation scavenging capacities than samphire and saltbush for both the crude and the purified extracts. The activity of the crude extract of sea parsley was 122.78±9.64 µM TE/g DW of the extract (the same expression was used in all cases in this section(, which was about 65% higher than samphire at 74.32±2.71 µM TE/g and over 80% higher than saltbush at 68.15±3.17 µM TE/g, respectively. For the purified extracts, the ABTS radical cation scavenging activity of sea parsley was 286.19±13.65 µM TE/g which was more than 3-fold higher than saltbush (96.18±8.64 µM TE/g) and two-fold higher than samphire (135.169±10.26 µM TE/g). Purification has resulted in a significant concentration of phenolic antioxidants in the extracts with the purified extracts showing considerably higher ABTS radical cation scavenging activities than the respective crude extracts. This was particular so for sea parsley where the ABTS radical cation scavenging activity of the purified extract was 2.34 times higher than the crude extract. For samphire and saltbush, the activity of the purified extracts was 1.82 and 1.41 time higher, respectively, than the crude extract. These results are broadly consistent with those in Tables 4.2 and 4.3.

Table 4.4. ABTS free radical scavenging capacities of crude and purified extracts from native Australian saltbush, samphire and sea parsley1.

Plant Crude Purified Increase of purified species (µM TE/g, Extract (µM TE/g, Extract over crude extract DW) DW) (times) Samphire 74.32±2.71b 135.17±10.26b 1.82 Saltbush 68.15±3.17c 96.18±8.64c 1.41 Sea Parsley 122.78±9.64a 286.19±13.65a 2.34 1Antixidant activity is expressed as micromole trolox equivalent (TE) per gram dry weight of the extract. The data represent mean ± standard deviation of at least five independent experiments. Means with different letters in the same column were significantly different with p < 0.05 (n = 5).

4.2.5.2. DPPH free radical scavenging capacity

DPPH free radical scavenging capacity is another commonly used method for evaluating the antioxidant activity of phytochemicals, which was also used in the current study to determine the antioxidant activity of the three native plants (Table 4.5.). The data trends 74 are very similar to those of the ABTS free radical scavenging capacities (Table 4.4.). Sea parsley showed the highest DPPH free radical scavenging capacity for both the crude and purified extracts, followed by samphire while saltbush displayed the lowest activities. As expected, the purified extracts showed marked higher DPPH free radical scavenging capacities than the crude extracts for all three plant samples, ranging from 1.74 times for saltbush to 2.38 times for sea parsley. These results demonstrated the effectiveness of the purification procedure in removing impurities and concentrating phenolic compounds.

Table 4.5. DPPH free radical scavenging capacities of crude and purified extracts from native Australian saltbush, samphire and sea parsley1

Plant Crude Purified Increase of purified species (µM TE/g, Extract (µM TE/g, Extract over crude extract DW) DW) (times) Samphire 97.16±5.34b 221.96±11.16b 2.28 Saltbush 89.65±7.26c 156.23±8.41c 1.74 Sea Parsley 124.59±9.08a 297.13±10.99a 2.38 1Antixidant activity is expressed as micromole trolox equivalent (TE) per gram dry weight of the extract. The data represent mean ± standard deviation of at least five independent experiments. Means with different letters in the same column were significantly different with p < 0.05 (n = 5).

4.2.5.3. Ferric reducing antioxidant power (FRAP) assay

FRAP is an assay that measures the total reducing power of the electron donating antioxidants in a reaction system and is frequently applied to measure the antioxidant capacity of food and food components. As shown in Table 4.6 and unexpectedly, samphire showed the highest FRAP values for both the crude and purified extracts. The FRAP value of the crude extracts of samphire was 416.54±20.72 µM TE/g extract DW, followed by sea parsley )295.75±21.22 µM TE/g extract DW and saltbush 292.56±29.54 µM TE/g extract DW, which were very similar. The purified extracts followed a similar trend with samphire exhibiting the highest ferric reducing activity, but the FRAP value of sea parsley was significantly higher than that of saltbush, which is somewhat different from the trend for the crude extracts. The data trends of FRAP assay were different from those of ABTS and DPPH free radical scavenging capacity assays.

Table 4.6. Ferric reducing antioxidant power of crude and purified extracts from native Australian saltbush, samphire and sea parsley1

Plant Crude Purified Increase of species (µM TE/g, Extract (µM TE/g, Extract purified over crude DW) DW) extract (times) Samphire 416.54±20.72a 792.88±53.36a 1.90 Saltbush 292.56±29.54c 423.93±31.36c 1.45 Sea Parsley 295.75±21.22b 473.03±39.69b 1.60 1Antixidant activity is expressed as micromole trolox equivalent (TE) per gram dry weight of the extract. The data represent mean ± standard deviation of at least five independent experiments. Means with different letters in the same column were significantly different with p < 0.05 (n = 5).

4.3. Correlation analysis

Pearson correlation analysis was carried out to determine the correlations between the total phenolic and the antioxidant capacities obtained by three different assays and between the three assays (Table 4.7). The total phenolic content was found to be significantly correlated with all three assays, with the correlations being particularly strong with ABTS and DPPH radical cation scavenging capacities (P<0.01). The results of ABTS and DPPH radical cation scavenging capacities were also strongly correlated (p<0.01). However, neither the results of ABTS nor DPPH radical cation scavenging capacities were significantly correlated with those of FRAP assay (P>0.05).

Previous studies have reported a range of trolox equivalent antioxidant capacity (TEAC) values for phenolic extracts of commercially edible Australian native plants. For example, Shan et al. (2005) reported TEAC values, obtained by using ABTS radical scavenging capacities assay, for crude extracts of rosemary (378.0 µmol TE/g, DW), basil (295.9 µmol TE/g, DW), coriander (70.2 µmol TE/g, DW), dill (63.6 µmol TE/g, DW) and the common parsley (63.1 µmol TE/g, DW). The TEAC values of the three native plants in the current study fall between these values. Interestingly, TEAC value by the ABTS assay for sea parsley (122.78±9.64) was almost twice that for the common parsley, which has a similar look and taste to sea parsley but belongs to a different genus.

Table 4.7. Pearson correlation coefficient (r) and significance level (p) for relationships between total phenolic content, and antioxidant assays of Australian native plants

TPC ABTS DPPH FRAP TPC 1.000 0.896 0.798 -0.532 0.000** 0.000** 0.041* ABTS 1.000 0.965 -0.185 0.000** 0.509 DPPH 1.000 0.038 0.894 FRAP 1.000

*Correlation is significant at the 0.05 level; ** Correlation is significant at the 0.01 level. TPC: total phenolic content, ABTS: 2,2’-azino-bis (3ethylbenzothiazoline-6-sulphonic acid), DPPH: di(phenyl)- (2,4,6-trinitrophenyl) iminoazanium, FRAP: ferric reducing capacity power.

Note should be made that the TEAC values obtained by the FRAP assay for the three plants were substantially greater than those produced by the ABTS and DPPH assays. Similar results have been reported by several other researchers including Nilsson et al. (2005) for fruits and vegetables and Wong et al. (2006) for herbs. Wong et al. (2006) suggested some phenolic compounds are more efficient in reducing ferric irons than scavenging free radicals, which could explain the different results obtained by FRAP and ABTS and DPPH assays. Benzie and Choi (2014) reviewed the advantages and limitations of the commonly used methods for measuring antioxidant capacities. They conclude that no one method is perfect and can adequately accommodate all the different antioxidants, and suggest that more than one methods should be used in the measurement to obtain a more complete picture of the antioxidant capacity in the samples.

4.4. Conclusion

Results presented in this chapter demonstrated that the three native Australian plants, namely samphire, saltbush and sea parsley possessed relatively high levels of phenolic contents and antioxidant capacities, as assessed by three different assays, when compared with Australian native plants in general. Of the three plant samples, sea parsley contained the highest amount of phenolic compounds and exhibited the highest antioxidant capacity when determined by the ABTS and DPPH free radical scavenging capacity assays. Saltbush had the lowest amount of phenolic compounds and showed the lowest antioxidant activities when assessed by all three assay methods. Samphire had the second highest phenolic content and also displayed the greatest FRAP value. Purification of the extracts by liquid chromatography with Amberlite® resin (XAD-7) resulted in a significant concentration of the phenolic compounds with the total phenolic content increased by 1.43-2.67 times in the purified extracts. Similarly, the purification caused a marked increase in the antioxidant activity of the extracts, with the antioxidant capacities increased by 1.41-2.38 times depending on the plants and assay methods. Correlation analysis shows that the phenolic content was significantly correlated with the antioxidant capacities as measured by all three assays as well as between the results of ABTS and DPPH free radical scavenging capacity assays (p<0.01), but neither was correlated with the FRAP assay results.

CHAPTER 5 Inhibitory Effects of Saltbush, Samphire and Sea Parsley on Health Related Enzymes

5.1. Introduction

One of the major health challenges of modern industrialized societies is the prevalence of the so called “metabolic syndrome”, which is a cluster of adverse medical conditions including overweight and obesity, hypertension, hyperglycemia and high serum triglycerides (Dandona et al., 2005). Metabolic syndrome is associated with heightened risk of heart disease, stroke and type 2 diabetes, which are the leading causes of death in many countries (Brand-Miller and Holt, 1998). Unhealthy life style plays an essential role in the development of metabolic syndrome, with poor diet such as over consumption of energy intense foods, e.g., carbohydrates and fats, being a major part of it (Bahadoran et al., 2015, Marlatt et al., 2016). Several digestive enzymes, including α-glucosidase, α-amylase and pancreatic lipase, are involved in breaking down carbohydrate and fats into smaller molecules, which are promptly absorbed by the human body. Normally, this process is quite essential for the body, but over absorption of carbohydrates and fats causes excessive accumulation of energy in the body, which can ultimately lead to the development of metabolic syndrome (Sakulnarmrat and Konczak, 2012).

In recent decades, a large number of studies have demonstrated that consumption of plant- based foods, such as fruits, vegetables and whole grains, can help prevent the development of metabolic syndrome, partly due to the capacity of the plants to inhibit the digestive enzymes (Sakulnarmrat and Konczak, 2012). Studies have shown that phenolic compounds are major contributors to the inhibition of the enzymes (Dandona et al., 2005, Bahadoran et al., 2013). A number of studies have investigated the capacity of native Australian plants in inhibiting these metabolically important enzymes. For example, Sakulnarmrat and Konczak (2012) examined the α-glucosidase inhibitory activity of phenolic compounds extracted from native plants and found that some of them, such as lemon myrtle and Tasmannia pepper leaf, displayed significant activities. Sakulnarmrat et al. (2014) studied the α-glucosidase inhibition activity of the phenolic fractions extracted from quandong and Davidson’s plum. It was found that both were potent inhibitors of the enzyme with IC50 values of 0.39 mg/mL and 0.13 mg/mL, respectively.

In the previous chapter, it was demonstrated that the Australian native plants samphire, saltbush and sea parsley contained relatively high levels of phenolic compounds, which displayed high levels of antioxidant activities that are comparable with other Australian native plants. This means that the phenolic extracts of the three plants may also possess inhibitory activities against the metabolically important enzymes. Therefore, in this chapter, the inhibitory activities of the saltbush, samphire and sea parsley on α-amylase, α-glucosidase and pancreatic lipase, as well as on the inflammation-related hyaluronidase were investigated.

5.2. Results and discussion

5.2.1. Inhibition against α-amylase and α-glucosidase

α-Amylase and α-glucosidase are two main enzymes that are responsible for the digestion of dietary starch. α-Amylase cleaves the α-1,4-glycosidic bonds of starch (both amylose and amylopectin) molecules randomly, producing oligosaccharides of various sizes (Gropper and Smith, 2012). One of the major strategies in managing diabetes mellitus, especially the non-insulin dependent type 2, is to prevent or delay glucose absorption and subsequent elevation of postprandial blood glucose level through oral administration of α-amylase and α-glucosidase inhibitors such as Acarbose (Min and Han, 2014). However, these drugs produce a number of side effects including hernias, diarrhea, ulceration and flatulence (Zhang et al., 2017). Thus, there is a need to seek natural alternatives, such as foods that have inhibitory effects on the two enzymes but have no or less side effects.

5.2.1.1. Inhibition against α-amylase

The crude and purified extracts of the three plants were assessed for their inhibitory activity on α-amylase in the concentration range of 0.125 to 20 mg/mL (Figure 5.1). For both the crude and purified extracts of all three plant samples, the inhibition activity exhibited a concentration-dependent effect, i.e. the activities increased with increasing extract concentrations. However, the relationship between the inhibitory activity and the extract concentration was not linear, rather, the relationship resembled a negative exponential curve. In the lower concentration range (0.125-2.5 mg/mL), the inhibitory

80 activity increased sharply with an increase in the extraction concentration. With further increases in the extract concentration, however, the effect became less prominent, which is seen as the curve began to plateau.

Crude Extracts

% Inhibition % 20

0 0 5 10 15 20 25 Concentation (mg/ml)

Sea Parsley Samphire Saltbush

Purified Extracts

70 60 50 40 30

% Inhibition % 20 10 0 0 5 10 15 20 25 Concentation (mg/ml)

Sea Parsley Samphire Saltbush

Figure 5.1. α-Amylase inhibitory activity of the crude and purified extracts obtained from native Australian saltbush, samphire and sea parsley.

Of the three plants, sea parsley showed the highest α-amylase inhibitory activity, followed by samphire while saltbush exhibited the lowest activity. This data trend is true for both the crude and the purified extracts. For all three plants, purified extracts showed significantly 81 higher α-amylase inhibitory activities than the corresponding crude extracts. A better comparison between the relative inhibitory activities of the samples can be made by calculating their IC50 values, which is the concentration of the extract at which 50% of the α-amylase activity was inhibited (Table 5.1). As can be seen, for the crude extracts, sea parsley had the lowest IC50 value of 18.61 ± 1.64 mg/mL, which was followed by samphire (29.88 ± 2.14 mg/mL), while saltbush had the highest IC50 value of 78.36 ± 4.56 mg/mL. Similar trend was observed for the purified extracts, where sea parsley had an

IC50 value of 6.71 ± 0.53 mg/mL, followed by samphire (13.93 ± 1.44 mg/mL) and saltbush (51.84 ± 2.87 mg/mL). Furthermore, the IC50 values of purified extracts were significantly lower than those of the corresponding crude extracts. This demonstrates that the purification process had resulted in a concentration of the compounds with α-amylase inhibitory.

Table 5.1. α-Amylase inhibitory activity of crude and purified extracts obtained from native Australian saltbush, samphire and sea parsley

1 α-Amylase inhibition activity [IC50 (mg/mL)] Plants species Crude extract Purified extract Sea parsley 18.61±1.64c 6.74±0.53 c Samphire 29.88±2.14b 13.93±1.44 b Saltbush 78.36±4.56a 51.84±2.87 a

1 IC50: half maximum inhibitory concentration. The data represent the mean ± standard deviation of at least five independent experiments. Means with different letters in the same column were significantly different with p < 0.05 (n = 5).

5.2.1.2. Inhibition against α-glucosidase

The α-glucosidase inhibitory activity of the crude and purified extracts of the three plants at concentrations between 0.125 to 20 mg/mL was investigated and the results are presented in Figure 5.2. Similar to their effects on α-amylase, both the crude and purified extracts exhibited significant inhibition on α-glucosidase and the effects were dependent on the concentration of the extracts. The relationship between the inhibitory activity and the extract concentration followed a negative exponential curve. In the lower concentration range (0.125-2.5 mg/mL), the inhibitory activity rose steeply with increases in the extraction

82 concentration. With further increases in the extract concentration, however, the increases in the inhibitory activity slowed as the curve began to plateau.

Crude Extracts

70 60 50 40 30

% Inhibition % 20 10 0 0 5 10 15 20 25 Concentation (mg/ml)

Sea Parsley Samphire Saltbush

Purified Extracts

80 70 60 50 40

30 % Inhibition % 20 10 0 0 5 10 15 20 25 Concentation (mg/ml)

Sea Parsley Samphire Saltbush

Figure 5.2. α-Glucosidase inhibitory activity of crude (A) and purified (B) extracts obtained from native Australian saltbush, samphire and sea parsley.

For both the crude and purified extracts of the three plants, sea parsley showed the highest α-glucosidase inhibitory activity, followed by samphire while saltbush exhibited the lowest activity. Purification of the extracts resulted in marked increases in their α- glucosidase inhibitory activities with purified extracts showing significantly higher activities than the corresponding crude extracts for all three plants. However, for a more direct comparison between the relative inhibitory activities of the three plant samples, the data in Figure 5.2 were extrapolated to determine the IC50 values (Table 5.2.). For the crude extracts, sea parsley gave the lowest IC50 value of 9.73 ± 0.45 mg/mL, which was followed by samphire (19.36 ± 0.64 mg/mL), while saltbush had the highest IC50 value of 31.10 ± 1.44 mg/mL. The same trend was observed for the purified extracts, where sea parsley had an IC50 value of 3.05 ± 0.32 mg/mL, followed by samphire (8.84 ± 0.46 mg/mL) and saltbush (15.72 ± 1.07 mg/mL). Moreover, the IC50 values of purified extracts were significantly lower than those of the corresponding crude extracts. These results indicated that the purification process has resulted in a concentration of the compounds with α-glucosidase activities.

Table 5.2. α-Glucosidase inhibitory activity of crude and purified extracts obtained from native Australian saltbush, samphire and sea parsley

1 α-Glucosidase inhibition activity [IC50 (mg/mL)] Plants species Crude extract Purified extract Sea parsley 9.73±0.45c 3.05±0.32 c Samphire 19.36±0.64b 8.84±0.46 b Saltbush 31.10±1.44a 15.72±1.07 a

Many studies have shown that plant extracts exhibit inhibitory activities against α-amylase and α-glucosidase and the effects are usually attributed to the phenolic compounds in the extracts (McDougall et al., 2005, Apostolidis et al., 2006, Kwon et al., 2006, Cazzola et al., 2011, El-Beshbishy and Bahashwan, 2012, Wongsa et al., 2012, Indrianingsih et al., 2015, Irondi et al., 2016(. For example, Indrianingsih et al. (2015( evaluated the total phenolic content and α-glucosidase inhibitory activity of the ethanolic extracts of 18 tropical and 84 subtropical plants. It was found that the total phenol content in the plants was generally correlated with the α-glucosidase inhibitory activities. This trend was also found in the current study where sea parsley, which had the highest total phenol content, also exhibited the strongest inhibitory effect on α- amylase and α- glucosidase. Hu et al. (2017) investigated the α-glucosidase inhibitory activities of a number of medicinal plants as well as their ABTS and DPPH radical scavenging activities, which are closely associated with the phenolic content of the plants. They found that the α-glucosidase inhibitory activities of the plants were correlated with the antioxidant activities, a result which is also observed in the present study when the antioxidant activities were determined by the ABTS and DPPH radical scavenging capacity assays.

Several studies have investigated the inhibitory effects of Australian native plants on carbohydrate-digesting enzymes. Sakulnarmrat and Konczak (2012) examined the α- glucosidase inhibitory activity of phenolic compounds extracted from native herbs and found that lemon myrtle (IC50 = 0.13 mg/mL), anise myrtle (IC50 = 0.30 mg/mL) and

Tasmannia pepper leaf (IC50 = 0.83 mg/mL) displayed relatively high activities while the activity of bay leaf (IC50 = 3.21 mg/mL) was much lower. Sakulnarmrat et al. (2014) studied α-glucosidase inhibitory activity of phenolic extracts from the native fruits quandong and Davidson’s plum and found that they exhibited comparable activities to anise myrtle and lemon myrtle, with IC50 values of 0.39 mg/mL and 0.13 mg/mL, respectively. Compare to these herbs and fruits, the α-glucosidase inhibitory activity of sea parsley (purified extract) was comparable to bay leaf, but lower than the others, while the activities of samphire and saltbush were much lower.

The α-amylase and α-glucosidase inhibitory activities of the three plant extracts are likely due to the phenolic compounds present in them. Previous studies have assessed the inhibitory activities of many individual phenolic compounds. For example, caffeic acid was found to be a particularly good inhibitor of α-glucosidase with 91.3% inhibition of this enzyme at the concentration of 1mg/mL (Kwon et al. 2006). Funke and Melzig (2005) investigated the inhibitory effect of several phenolic compounds on α-amylase activity. Compounds such as apigenin, tannic acid, ferulic acid, fisetin, rosmarinic acid, luteolin, chlorogenic acid and caffeic acid were able to inhibit the activity of α-amylase by their reactivities towards the enzyme. They demonstrated that the inhibition effect on α-amylase and other enzymes are dependent on the concentration of phenolic compounds and 85 suggested that it may also be related to the number and position of hydroxyl groups in the molecules. As detailed in Chapter 6, several of these phenolic compounds, such as apigenin and caffeic acid, are present in the extract of the sea parsley and likely to be mainly responsible for the inhibitory effects of the extract on α-amylase and α-glucosidase.

5.2.2. Inhibition against pancreatic lipase

Pancreatic lipase is an enzyme that hydrolyses dietary fats with the release of fatty acids which can be absorbed by the body (Birari and Bhutani, 2007). Accumulated fat absorption can cause excessive fat deposition in adipose tissues of the body, which leads to weight gains (Garza et al., 2011). Inhibition of pancreatic lipase can prevent this occurrence and, thus, lessen the risk of overweight and obesity (Birari and Bhutani, 2007(. The inhibitory activity of native Australian saltbush, samphire and sea parsley extracts in the concentration range of 0.125- 20 mg/mL on pancreatic lipase was determined and the results are shown in Figure 5.3. Extracts from all three plants exhibited significant and concentration- dependent inhibitory effects on the enzyme. The relationship between the concentration of the extract and the inhibition activity followed a negative exponential curve for all the plants, which is similar to that between the concentration of the extract and the inhibitory activities on α-amylase and α-glucosidase. Of the three plants, sea parsley gave the highest lipase inhibition activity for both the crude and the purified extracts, followed by samphire while saltbush exhibited the lowest activity. The differences in the inhibitory activity between the three plants are seen more clearly when their IC50 values are compared (Table 5.3). The IC50 value of crude extract of sea parsley was about half and 1/3 of that for samphire and saltbush, respectively. For the purified extract, the differences were even more pronounced with sea parsley having an IC50 value of 2.24 mg/mL, compared with the values of 9 .38±0.8 mg/mL and 13.84 mg/mL for samphire and saltbush respectively. As expected, the purified extracts showed marked higher lipase inhibition activities than the corresponding crude extracts. This suggests that phenolic compounds are the ones that are mainly responsible for the lipase inhibition effects and the removal of non-phenolic components resulted in a concentration of the phenolic compounds in the purified extracts (Dai and Mumper, 2010, Oszmianski et al., 2011).

Crude Extracts 70

30 % Inhibition % 20

0 0 5 10 15 20 25 Concentation (mg/ml)

Sea Parsley Samphire Saltbush

Purified Extracts 90 80 70 60 50 40

% Inhibition % 30 20 10 0 0 5 10 15 20 25 Concentation (mg/ml)

Sea Parsley Samphire Saltbush

Figure 5.3. Pancreatic lipase inhibitory activity of crude and purified extracts obtained from native Australian saltbush, samphire and sea parsley.

Table 5.3. Pancreatic lipase inhibitory activity of crude and purified extracts obtained from native Australian saltbush, samphire and sea parsley

1 Pancreatic lipase inhibition activity [IC50 (mg/mL)] Plants species Crude extract Purified extract Sea parsley 7.37±0.52c 2.24±0.31 c Samphire 14.28±0.94b 9 .38±0. 86b Saltbush 20.96±1.34a 13.84± 1.21a

Furthermore, this trend is particularly pronounced for the extract of sea parsley where the

IC50 value of the crude extract was more than 3 times higher than that of the purified extract. One possible explanation for this is that the crude extract of sea parsley contained a greater proportion of impurities compared with the other two plants. However, as shown in Table 4.2 (Chapter 4), the yield of purified extract of sea parsley was slightly over half of that for the crude extract, i.e., about half of the crude extract was impurities for this plant; whereas the yield of the purified extract was less than 40% for the other two plant samples. Therefore, this explanation must be discounted. Another possible explanation is that the phenolic compounds in sea parsley are more potent inhibitors compared with those in the other two plants and this will be explored in the next chapter.

Previous studies have reported that polyphenol-rich extracts can be effective inhibitors of pancreatic lipase (McDougall, 2009, Buchholz and Melzig, 2015). Paulina et al. (2014) studied the methanolic, water and acetic extracts of chokeberry for their inhibition on lipase and found that the acetic extracts gave the highest inhibitory activity with an IC50 of 83.45 mg/ml. Sompong et al. (2016) investigated the inhibitory activity of several herbal medicines on pancreatic lipase. All extracts of the herbs exhibited the inhibitory effects on the enzyme with the IC50 values ranging between 0.015 to 4.259 mg/mL.

Additionally, Dechakhamphu et al. (2015) reported IC50 values of 0.11-0.45 mg/mL for a number of medicinal plants. Tang et al. (2016, 2017) evaluated the effect of purified extracts of native Australian mints (river mint, mint bush and spearmint) on pancreatic lipase and reported IC50 values of 1.90-4.70 mg/mL. In the present study, the purified

88 extract of sea parsley had an IC50 of 2.24mg/mL, which was in the middle of the reported results for medicinal plants and herbs. However, the IC50 values of saltbush and samphire were much higher, indicating that they were less effective inhibitors of lipase. The previous studies have generally demonstrated a significantly positive correlation between total phenolic content and the lipase inhibition activity. This agrees with the results of the present study which showed that the higher phenolic content in sea parsley also led to more effective inhibition of pancreatic lipase. This provided strong supporting evidence demonstrating that phenolic compounds were the main inhibitory agents on pancreatic lipase as well as α-glucosidase and α-amylase (Önal et al., 2005, Adisakwattana et al., 2010, Ngamukote et al., 2011, Worsztynowicz et al., 2014, Dechakhamphu et al., 2015).

5.2.3 Inhibition against hyaluronidase

Hyaluronidase is a family of related enzymes that are known to be associated with inflammation and allergic responses. The enzyme acts by hydrolysing the polysaccharide hyaluronic acid in the extracellular matrix of connective tissues and the excessive activity of the enzyme can result in metabolic imbalances that lead to chronic inflammation (Soberón et al., 2010(. Apart from inflammation, hyaluronidase may also be involved in several other pathological conditions, including increased risks of cancer (Johnson, 2007). Therefore, inhibitors of hyaluronidase may have potential therapeutic effects on inflammation, allergy and other disorders. In this study, the hyaluronidase inhibitory activities of the crude and purified extracts of saltbush, samphire and sea parsley at concentrations between 0.125 to 30 mg/mL were investigated and the results are presented in Figure 5.4.

All three plant extracts displayed significant concentration-dependent inhibitory effects on hyaluronidase. The relationship between the concentration of the extracts and the inhibition effects followed a negative exponential curve for all three plants, a trend that is observed for the inhibitory activities on α-amylase, α-glucosidase and pancreatic lipase. However, different from the effects on the digestive enzymes where sea parsley showed the highest inhibitory activities, here samphire gave the highest hyaluronidase inhibition activity for both the crude and the purified extracts, followed by sea parsley while saltbush

89 exhibited the lowest activity. This data trend is also evident when the inhibitory IC50 values of three samples on the enzyme is compared (Table 5.4.).

Crude Extracts

70 60 50 40 30

20 % Inhibition % 10 0 0 5 10 15 20 25 30 35 -10 Concentation (mg/ml)

Samphire Sea Parsley Saltbush

Purified Extracts

80 70 60 50 40 30

% Inhibition % 20 10 0 -10 0 5 10 15 20 25 30 35 Concentation (mg/ml)

Samphire Sea Parsley Saltbush

Figure 5.4. Hyaluronidase inhibitory activity of crude and purified extracts obtained from native Australian saltbush, samphire and sea parsley.

Table 5.4. Hyaluronidase inhibitory activity of crude and purified extracts obtained from native Australian samphire and sea parsley and saltbush

1 Hyaluronidase inhibition activity [IC50 (mg/mL)] Plants species Crude extract Purified extract Samphire 9.98±0.91c 5 .51±0. 89c Sea parsley 13.06±0.83b 8.70± 0.53b Saltbush 65.51 ±3.44a 4 4 .34± 2.54a

As can been seen, the IC50 value of the crude extract of samphire was the lowest at 9.98 ± 0.91 mg/mL, which was followed by sea parsley at 13.06 ± 0.83 mg/mL, while saltbush had the highest IC50 value of 65.51 ± 3.44 mg/mL. For the purified extract, the difference between the IC50 values of samphire and sea parsley were not as prominent, which were

5.51 ± 0.89 mg/mL and 8.70 ± 0.53 mg/mL respectively. Saltbush had a much large IC50 value of 44.34 ± 2.54 mg/mL. For all three plant samples, the purified extracts had significantly lower IC50 values (higher inhibitory activities) than the respective crude extracts, suggesting the purification step has concentrated the phenolic components which were largely responsible for the hyaluronidase inhibitory activity. It is interesting to note that for both the crude and purified extracts, samphire also showed considerably greater ferric reducing activities (high FRAP values) than the other two plants (Table 4.6., Chapter 4). This appears to suggest that the phenolic compounds in this plant, which were effective in chelating ferric ions, were also good inhibitors of hyaluronidase. However, further research is needed to confirm such association.

For a long time, phytochemicals have been used for the treatment of inflammation and associated diseases (El Beyrouthy et al., 2008, Krishnaswamy, 2008). Tan et al. (2011c) reported that the polyphenolic-rich fractions of native Australian fruits exhibited anti- inflammatory activities while Mueller et al., 2010 showed that phenolic extracts of herbs and spices also showed such activities. Perera et al. (2016) examined the hyaluronidase inhibitory and antioxidant activities of 10 medicinal plants in Sri Lanka and reported moderate inhibitory effects ranging from 30.79 ± 0.31 to 69.35 ± 4.96% on hyaluronidase

91 for the plant extracts. They also demonstrated that the hyaluronidase inhibitory effects were not always correlated with antioxidant activities. Some plants, such as S.cochinchinesis and C.innophyllum, had a high hyaluronidase inhibitory but moderate levels of antioxidant activities compared with other plants. This appears to agree with the current study where samphire had the highest inhibition on hyaluronidase, but lower levels of phenolic content and antioxidant activities as measured by the ABTS and DPPH radical scavenging assays.

5.3. Conclusion

This chapter demonstrated that samphire, saltbush and sea parsley possessed significant, concentration-dependent inhibitory activities against four metabolically important enzymes, namely α-glucosidase, α-amylase, pancreatic lipase and hyaluronidase. The inhibitory effects on the enzymes followed negative exponential curves and the IC50 values of the inhibition are comparable to many plant foods including several native Australian fruits and herbs. Of the three plants, sea parsley exhibited the highest inhibitory activities on α- glucosidase, α-amylase and pancreatic lipase for both the crude and purified extracts, followed by samphire. Samphire exhibited the highest inhibitory activity against hyaluronidase, followed by sea parsley, while saltbush extracts gave the lowest inhibitory activities on all the enzymes. Purified extracts showed greater inhibitory effects than the crude extracts in all cases and the inhibitory activities were generally correlated with the phenolic contents and antioxidant capacities of the plant samples. This demonstrated that phenolic compounds were the main components responsible for the inhibitory effects. Because sea parsley has shown the highest overall biological activities among the three plants, and due to the scope of this study, this plant was selected for identification of the phenolic components, which is described in the next chapter.

CHAPTER 6 Identification and Quantification of Phenolic Compounds in Sea Parsley (Apium prostratum)

6.1. Introduction

Apium is a genus consisting of about 20 species of flowering plants including celery and sea parsley. It is widely distributed in Australia, Asia, Africa, Europe and South America (Cowan and Chapman, 1992). At least four Apium species are endemic to Australia, which are Apium annuum, Apium prostratum, Apium insulare and Apium leptophyllum (de Salas and Baker, 2017). Sea parsley (Apium prostratum) is a native plant of Australia which is widely found along the southern coastline. Dried leaves of sea parsley are used as an Australian spice mix which tastes similar to celery (Peter, 2006). To date, there have been no reported studies that have investigated the phenolic composition of these Australian native plants.

In this thesis, the phenolic composition and in vitro biological activities of sea parsley, together with saltbush and samphire, were systematically investigated for the first time. Results presented in the previous two chapters have demonstrated that sea parsley had the highest phenolic content and exhibited greatest bioactivities among the three plants, including antioxidant capacity and inhibitory activities on α-amylase, α-glucosidase, pancreatic lipase and hyaluronidase. The logical next step in the investigation is to identify the phenolic compounds in the plants. Because the scope of this MPhil study which does not permit the identification of phenolic compounds in all three plants, sea parsley was selected for this purpose.

High-performance liquid chromatography (HPLC) is a separation technique that is widely applied to the analysis of phytochemicals (Chai et al., 2014, Kumar et al., 2015, 2016, Yuan et al., 2016). It is extensively used for the separation, identification and quantification of phenolic compounds (Ignat et al., 2011). HPLC copuled with photodiode array detection (HPLC-PDA) can provide tentative identification of phenolic compounds, especially when reference standards are available for comparison. However, the compounds identified by HPLC-DAD needs to be confirmed by techniques such as liquid chromatography- mass spectrometry (LC-MS), which can provide information on the molecular mass of

93 the phenolic compounds. When the molecular mass of the phenolic compounds matches that of the reference standard, the chemical identity of the compound can usually be confirmed (Justesen, 2000, Ignat et al., 2011).

The main objective of this chapter was to identify and quantify the phenolic compounds in sea parsley (Apium prostratum) by using a combination of HPLC-PDA and LC-MS analyses.

6.2. Results and discussion

6.2.1. Determination of phenolic compounds in sea parsley (Apium prostratum) by high-performance liquid chromatography-photodiode array detector

The identification of phenolic compounds in plant extracts can be a difficult and complex process because the extract is expected to contain a variety of different phenolic compounds. In the present study, the extract of sea parsley was first purified by liquid chromatography using XAD-7 Amberlite® resins. Phenolic compounds in the purified extracts were then identified using a four-step identification and confirmation processes, as illustrated in Figure 6.1.

1. 2. 3. Comparison UV 4. of HPLC Co-chromatogr- absorption Confirmation retention time aphy with standard spectra by LC-HRMS with standard comparison compounds compounds

Figure 6.1. Processes used to identify phenolic compounds in the purified extract of sea parsley (Apium prostratum).

The first step involved comparison of HPLC retention times of the peaks in the sample with those of phenolic standards to attain preliminary identification. Twelve standards were used, including five phenolic acids (caffeic, chlorogenic, ρ-coumaric, ferulic and sinapic acids) and seven flavonoids (apiin, apigenin, catechin, naringenin, hesperetin, luteolin, and quercetin). These compounds were selected based on their availability as well as their possible presence in sea parsley sample according to the results of previous studies on Apium species (Mattila and Hellström, 2007, Li et al., 2014, Kooti and Daraei, 2017). The second step involves verification of the retention times of peaks by co-eluding the sample with suspect compounds. In the third step, the UV spectra of the suspected peaks were compared with those of the reference standards to further verify their possible identity. Finally, the confirmation of the peaks was achieved by comparing their MS spectra with those of the standard compounds using liquid chromatography-high resolution mass spectrometry (LC-HRMS).

Initially, the HPLC analysis of the purified extract of sea parsley (Apium prostratum) was carried out with a 60 min elution program, which resulted in an elution profile as illustrated in Figure 6.2. Under the gradient elution program, the target analytes began to elude from the column after 6 min and all compounds were eluded in about 28 min. Consequently, the elution program was shortened to 30 min in subsequent analytical runs. Absorbance was recorded with three different wavelengths (280, 320 and 370 nm) to ensure that compounds with different absorbance maxima were all recorded. Following this process, 320 nm was found to be the wavelength at which most of the compounds in the extract of sea parsley gave the most prominent peaks. At the wavelength, HPLC profile of the extract showed seven major peaks (Figure 6.3). Seven compounds, namely apiin, apigenin, caffeic acid, ferulic acid, ρ-coumaric acid, luteolin, and catechin, were provisionally identified based on the comparison of peak retention times, co-elution and comparison of UV spectra with reference standards (Table 6.1).

Figure 6.2. Initial HPLC chromatogram of purified sea parsley ) Apium prostratum) extracts with a run time of 60 min and the peaks recorded at 320 nm.

3 5

2 6 7 1

Figure 6.3. HPLC chromatogram of purified sea parsley )Apium prostratum) extract with the peaks recorded at 320 nm. Provisional identity of the peaks: catechin (1), caffeic acid (2), ρ-coumaric acid (3), apiin (4), ferulic acid (5), luteolin (6) and apigenin (7).

Table 6.1. Retention times, co-elution results and UV spectra of the phenolic compounds identified in the methanolic extract of sea parsley (Apium prostratum) as compared with reference standards

Peak No. Retention times Co-elution UV absorption maxima (min) (nm) Compound Sea parsley Standard Sea parsley Standards

1.Catechin 6.92 6.88 Yes 279 279 2.Caffeic acid 10.59 10.57 Yes 244, 321 244,319 3. ρ-Coumaric acid 16.48 16.39 Yes 310 277,310 4.Apiin 17.11 17.14 Yes 267, 335 267,335 5.Ferulic acid 17.43 17.48 Yes 322 322 6.Luteolin 19.17 19.19 Yes 347 292,347 7.Apigenin 25.63 25.69 Yes 268, 334 268,334

As can be seen from Table 6.1., the retention times of the suspect compounds in sea parsley were very close to those of the reference standards, with the differences being between 0.02 and 0.09 min. When the reference standards were individually spiked into the sea parsley extract, they all co-eluded with the respective suspect compounds in the sample. The absorption maxima of peaks 1, 4, 5 and 7 matched those of the reference standards perfectly, while those of the remaining peaks also matched the standards closely. Furthermore, information available in the literature indicates that apiin, apigenin, caffeic acid, ferulic acid, ρ-coumaric acid and luteolin are often present in herbs from the Apium genus (Mattila and Hellström, 2007, Li et al., 2014, Wesam and Nahid, 2017). The chemical identities of the peaks were further confirmed by LC-HRMS (Section 6.2.2).

6.2.2. Identification and confirmation of phenolic compounds in sea parsley (Apium prostratum) by liquid chromatography-high resolution mass spectrometry (LC- HRMS).

Liquid chromatography-high resolution mass spectrometry (LC-HRMS) along with the LC-MS analysis software Xcalibur and the chemical database ChemSpider were employed to confirm the chemical identity of the seven peaks that were detected by HPLC-DAD. The 97 mass spectra recorded by the LC-HRMS instrument have an accuracy of lesser than 2 ppm. Selection of ionization method is crucial to the success of this type of mass spectrometry and this depends on the properties of the analyte molecules (Krauss et al., 2010, Wu et al., 2012). In most of the reports for investigation on phenolic compounds, the positive ion mode was used; nevertheless, it has been reported that the negative ion mode using both electrospray ionisation (ESI) and atmospheric pressure chemical ionisation (APCI) were more responsive in flavonoid analysis (Justesen, 2000, Cuyckens and Claeys, 2004, Shen et al., 2011). In the present study, the negative mode with both heated electrospray ionisation (HESI) and APCI sources were used for analysing the phenolic compounds in the purified extract of sea parsley because previous studies in our laboratory have shown that certain compounds in Australian native herbs demonstrated a higher signal to noise ratio on the mass spectra with the former, while other compounds, such as apigenin and caffic acid, showed a better signal to noise with the APCI sources (Tang et al., 2016). The range for mass-to-charge ratio (m/z) was set between120 to 950, which covers essentially all the known phenolic compounds. The total ion current chromatograms obtained by HESI and APCI are shown in Figures 6.4 and 6.5., respectively. Clearly, atmospheric pressure chemical ionisation (APCI) produced better results as the chromatogram had a better baseline and significantly less noises compared to that obtained by HESI. Based on these results, APCI was used in all subsequent LC-HRMS analyses.

Figure 6.4. LC-HRMS total ion current (TIC) chromatogram of sea parsley ) Apium prostratum) using heated electrospray ionisation with a mass range between m/z 120-950.

Figure 6.5. LC-HRMS total ion current (TIC) chromatogram of sea parsley ) Apium prostratum) using atmospheric pressure chemical ionisation with a mass range between m/z 120-950.

To confirm the chemical identity of the peaks in the sea parsley extract, their mass spectra were compared with those of reference standards obtained by LC-HRMS under the same analytical conditions, and with those generated by the Xcalibur software in conjunction with the ChemSpider mass spectra database. The Xcalibur software produced an empirical formula based on the molecular ions [M-H]- of the peak, from which simulated mass spectra were generated. For example, Figure 6.6.A shows the mass spectra of the peak at 11.94 min of the TIC chromatogram (Figure 6.5.), which had a molecular ion [M-H]- of 563.14. This peak corresponded to peak 4 in the HPLC-PDA chromatogram (Figure 6.3.), which was previously tentatively identified as apiin. Figure 6.6.B shows the mass spectra of apiin reference standard while Figure 6.6.C shows those generated by Xcalibur. As can be seen, the molecular ions in both Figures 6.6.B and 6.6.C were 563.14, which was the same as that for the peak at 11.94 min of the TIC chromatogram (peak 4 in Figure 6.3.), confirming that this peak was apiin. The molecular ions produced by LC-HRMS for the seven peaks in Figure 6.3 are presented in Table 6.2, while their mass spectra as well as those of the corresponding reference standards are provided in Appendix 4-10. Applying the procedure described above, the seven peaks were confirmed to be catechin, caffeic acid, ρ-coumaric acid, apiin, ferulic acid, luteolin and apigenin. The chemical structures of these phenolic compounds are shown in Figure 6.7.

- Figure 6.6. Comparison of molecular ion [M-H] (m/z 563) of apiin (C26H28O14) in sea parsley (Apium prostratum) (A), the mass spectra of apiin standard (B) and the simulated data generated from Xcalibur (C).

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Table 6.2. Molecular mass, empirical formula and HPLC retention times of the phenolic compounds identified in the methanolic extract of sea parsley (Apium prostratum)

Peak Compound Empirical HPLC Retention Molecular mass (HPLC- formula time (min, λmax) DAD) 1 Catechin C15H14O6 6.9 (279 nm) 289

2 Caffeic acid C9H8O4 10.6 (321 nm) 179

3 ρ-Coumaric acid C9H8O3 16.5 (310 nm) 163

4 Apiin C26H28O14 17.1 (267, 335 nm) 563

5 Ferulic acid C10H10O4 17.5 (322 nm) 193

6 Luteolin C15H10O6 19.1 (347 nm) 285

7 Apigenin C15H10O5 25.6 (268, 334 nm) 269

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Apiin Caffeic acid ρ-Coumaric acid

Ferulic acid Apigenin Luteolin

Catechin

Figure 6.7. Chemical structure of compounds identified in purified sea parsley (Apium prostratum) extract.

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6.2.3. Quantification of phenolic compounds by high-performance liquid chromatography- photodiode array detector analysis

High-performance liquid chromatography with a diode array detector (HPLC-DAD) is a method extensively used for the quantification of phenolic compounds (Irakli et al., 2012, Jin et al., 2013, Plaza et al., 2014, Šućur et al., 2014, Pires et al., 2017), and this method was also employed for this purpose in this study. Peaks of the phenolic compounds were recorded and integrated at 320 nm and their quantification was carried out with the external standard method with standard curves of individual compounds established using reference standards (Table 6.3). Apiin was the most predominant phenolic compound in sea parsley, accounting for 48.2% of the total phenolic compounds obtained by the HPLC method. Apigenin was the second most abundant compound, accounting for 24.8%, followed by caffeic acid (6%) and ferulic acid (2%), while ρ-coumaric acid, luteolin, and catechin which were present at trace levels (<1%).

Table 6.3. Quantification of phenolic compounds identified in the methanolic extract of sea parsley (Apium prostratum) by HPLC-PDA

Peak Compound Concentration (mg per g purified extract, DW) 1 Catechin 0.27 ± 0.003 2 Caffeic acid 37.91 ± 0.15 3 ρ-Coumaric acid 11.62 ±0.08 4 Apiin 147.66 ± 1.14 5 Ferulic acid 25.62 ±0.22 6 Luteolin 7.18 ± 0.06 7 Apigenin 75.99 ± 0.84 Total 306.25

The data represent the mean ± standard deviation of three replicates.

The sum of the seven phenolic compounds gives a total amount of 306.25 mg/g extract (DW) in sea parsley. As reported in Chapter 4, the total phenolic content in the purified extract of sea parsley, as determined by the Folin-Ciocalteu assay was 362.28 GE mg/g extract (DW), which was about 18.30% higher than the result obtained by the HPLC 103 method. This discrepancy could be attributed to several factors. First, phenolic content obtained by the Folin-Ciocalteu method is expressed as gallic acid equivalent (GE), while the result obtained by the HPLC-DAD method is the sum of the amount of individual phenolic compounds, which would be a more accurate reflection of the phenolic components in the extract. Second, the purified extract still contained some residual non-phenolic impurities and some of which, like sugars and vitamin, can react with the Folin-Ciocalteu reagent (Naczk and Shahidi, 2004). Therefore, the result of Folin-Ciocalteu assay would represent an overestimation of the phenolic content in the sample. Finally, there are some trace components in the HPLC-DAD profile of the sea parsley extract that were too small to be integrated, which nevertheless would cause a small underestimation of the phenolic content in the sample. Such discrepancies of phenolic content as obtained by the two different methods have been reported previously. For example, (Sakulnarmrat et al., 2012) reported that the phenolic content obtained by the HPLC method for rabbit eye blueberry, southern highbush blueberry, anise myrtle, Tasmannia pepper leaf and lemon myrtle were respectively 30, 37, 55, 60 and 65% less than that obtained by the Folin- Ciocalteu assay.

The phenolic compounds identified in sea parsley have commonly been found in the Apium genus, which includes several varieties of the commercially significant vegetable, celery (Mattila and Hellström, 2007, Yao et al., 2010, Li et al., 2014, Wesam and Nahid, 2017). For example, Yao et al. (2010) investigated the phenolic compounds in 11 celery species and identified two flavonoids (apigenin and luteolin) and three phenolic acids (caffeic, ρ-coumaric and ferulic).

Apiin, or apigenin-7-O -apiosylglucoside, is a diglycoside of apigenin which has been found in the winter-hardy plants such as celery (Gupta and Seshadri, 1952, Li et al., 2014) and parsley (Meyer et al., 2006). It was reportedly isolated from these vegetables for the first time in 1843 )Braconnot, 1843). Mencherini et al. (2007) reported that apiin is the largest constituent of the phenolic extract from Apium graveolens L. var. dulce (common celery) while Li et al. (2014( found that apiin is present as a major phenolic compound in a number of species in the Apium genus. Similar findings were reported by Kaiser et al. (2013) who found that apiin was present in the common parsley at the level of 36.02 mg/g (dried leaves), accounting for 78.2% of the total phenolics. These findings broadly agree with the results of the current study where apiin was found to be most abundant phenolic 104 component in sea parsley by far, accounting for 48.2% of the total phenolic compounds. Apart from celery and parsley, apiin has also been isolated from vegetables such as green bell pepper, and the herbal plants roman chamomile and german camomile (Hostetler et al., 2013).

A few studies have reported on the biological activities of apiin. Li et al. (2013) demonstrated that apiin extracted from celery (Apium graveolens L. var. dulce) possesses good free radical scavenging activities and suggested that the antioxidant activity could protect body tissues from damage by oxidative stress. Li et al. (2014) evaluated the effects of apiin on several enzymes in mice in vivo, such as catalase (CAT), glutathione peroxidase (GSH-Px) and superoxide dismutase (SOD), which play significant roles in the antioxidant systems of organs including kidney, liver, brain, heart and serum in preventing damages caused by the reactive species. It was found that apiin had a significant scavenging activity on maleic dialdehyde (MDA) and lipofuscin (LPF) and also improved the activities of CAT, SOD and GSH-Px. Additionally, apiin and celery extracts displayed significant inhibitory effect on nitrite (NO) production and iNOS expression in LPS-activated J774.A1 cells demonstrating their anti-inflammatory effects (Mencherini et al., 2007). The presence of high levels of apiin in sea parsley indicates that this native Australian vegetable could potentially bring significant health benefits in these respects.

The second most abundant phenolic compound in sea parsley was apigenin, accounting for 24.8% of the total phenolics. Apigenin, or 4',5,7-trihydroxyflavone, is a flavone that is found in many vegetables, fruits, herbs and cereals, including celery, chamomile, maize, onions, oranges, parsley, rice, tea and wheat sprouts (Haytowitz et al., 2006, Patel et al., 2007, Wen et al., 2014, Yan et al., 2014, Reza, 2016). It has been reported as being one of the major bioactive components in celery (Mafuvadze et al., 2013, Li et al., 2014, Liu et al., 2016), which belongs to the same genus as sea parsley. High levels of apigenin have been reported in the Apiaceae family of plants in a number of studies (Justesen et al., 1998, Justesen and Knuthsen, 2001) including the Shengjie celery (A. graveolens L.) which exhibited the highest antioxidant activity among 11 different celery cultivars (Yao et al., 2010, Yao and Ren, 2011). Liao et al. (2017) investigated the antioxidant activity of apigenin existing in the crude extracts of lotus leaf and found the flavonoid to be an efficient scavenger of DPPH free radicals with an IC50 of 8.25 μM. The high levels of 105 apigenin and apiin present in the sea parsley (Apium prostratum) would likely contribute significantly to the strong antioxidant activity of the plant.

Zeng et al. (2016) investigated the inhibitory effects of apigenin on α-glucosidase and showed that the flavonoid is a potent inhibitor of the enzyme with an IC50 value of 10.5 µM. Proenc et al. (2017) also investigated the inhibitory activity of apigenin on α- glucosidase, but reported a higher IC50 value of 82µM. However, both IC50 values are significantly lower than that for acarbose which had an IC50 of 607 µM, which means that apigenin is a much more potent α-glucosidase inhibitor than this standard drug for managing type 2 diabetes. The α-amylase inhibitory activity of apigenin was investigated by Liao et al (2017) and it was found that the flavonoid is also a potent inhibitor of this metabolically important enzyme with an IC50 value of 25.14 μM. The high level of apigenin present in the methanolic extract of sea parsley is likely a major contributor to its strong inhibitory activities on α-glucosidase and α-amylase.

Other than antioxidant activities and inhibitory effects on the digestive enzymes, apigenin and its derivatives have been shown to possess a number of other health beneficial properties including anti-inflammatory and anti-carcinogenic effects (O’Prey et al., 2003, Škerget et al., 2005, Dimitrios, 2006, Madunić et al., 2018(, and these bioactive activities of apigenin tend to be more potent compared with other structurally related flavonoids (Gupta et al., 2001). These health effects have been demonstrated in several mammalian systems both in vitro and in vivo (Liu et al., 2004, Ali et al., 2014). In addition, apigenin has been found to induce apoptosis in several cancers in studies using both animal models and cell lines (Das et al., 2006, Nicholas et al., 2007, Kaur et al., 2008, Choi, 2009). It has been demonstrated that apigenin decreases inflammation associated with oxidative stress in tissues by modulating numerous interleukins, blood enzyme markers, oxidative stress markers and expression of several related enzymes (Ali et al., 2016). The high level of apigenin in sea parsley suggests that consumption of this native vegetable could bring significant health benefits to the consumers.

The methanolic extract of sea parsely contained three phenolic acids, namely caffeic, ferulic and ρ-coumaric acids, with caffeic acid being the most abundant of the three at 37.91 mg/g extract DW and representing about 6% of total phenolics in the extract. Caffeic acid is one of the most common phenolic acids in fruits and vegetables and also

106 coﬀee (Cliﬀord, 2000, Matei et al., 2006, Hao et al., 2014). The reported range of caffeic acid in vegetables, fruits and coffee are 1.2- 49.0 mg/100 g, 0.85- 23.4 mg/100 and 87 mg/100 g respectively (Matei et al., 2006, Mattila and Hellström, 2007), respectively, while the reported values for the common celery were between 8 and 13 mg/100 g FW (Herrmann, 1977). The level of caffeic acid in sea parsley was lower than that in coffee but higher than most fruits and vegetables. Caffeic acid is a potent antioxidant, which is partly due to the conjugation in the propenoic side chain, as well as electron delocalization by resonance between the aromatic ring and the propenoic group (Wojdylo et al., 2007). Masek et al. (2007) investigated the antioxidant potential of the caffeic acid using the DPPH and ABTS radical scavenging and FRAP assays. Caffeic acid was found to cause an inhibition of 28.5% and 32.1% for DPPH and ABTS radicals respectively and a 30.8% reduction of the iron ions at the concentration of 30 µg/ml. Similar results were reported by Gulcin, (2006) who also reported that the antioxidant capacities of caffeic acid were comparable with the common antioxidants butylated hydroxytoluene (BHT), butylated hydroxyanisole (BHA) and α-tocopherol but significantly higher than trolox. The presence of caffeic acid in sea parsley at a relatively high level would contribute to the strong antioxidant activity of the plant.

Additionally, caffeic acid has been reported to exhibit several other biological activities that are beneficial to health. These activities include anti-inflammatory (Toyoda et al., 2009, Aoshima et al., 2012), anti-obesity, (Piazzon et al., 2012), anti-mutagenic (Yamada and Tomita, 1996) and anti-tumor effects against several cancers (Chung et al., 2004, Lee et al., 2005, Karthikesan and Pari, 2008, Wu et al., 2011). Aoshima et al. (2012) reported an IC50 value of 66 µM for caffeic acid on hyaluronidase, while an IC50 value of 0.52 mg/mL and 1.81 mg/mL was reported for α-glucosidase and α-amylase, respectively, which were lower than those for acarbose (Chiou et al., 2017). Thus, the presence of caffeic acid in sea parsley would contribute to its inhibitory effects on these metabolically important enzymes, as presented in Chapter 5.

Ferulic acid is the second most abundant phenolic acid in see parsley. It is a hydroxycinnamic acid found in the cell walls of many cereals including brown rice, oats and whole wheat. Other than cereals, ferulic acid is also present in many vegetables and occur in especially high concentrations in bamboo shoots and popcorn (Zhao et al., 2008, Kumar and Pruthi, 2014, Bilge Ertekin et al., 2017). Ferulic acid has been demonstrated to possess strong 107 antioxidant activities, which is attributed to its carbon side chain groups and the position of the hydroxyl and methoxyl groups in the ring (Karamać et al., 2017). As a potent antioxidant, ferulic acid is also known to exhibit significant health benefits, similar to caffeic acid (Shiyi and Kin, 2004, Oresajo, 2008). In vitro studies have demonstrated that ferulic acid, along with caffeic acid, were the most potent inhibitors of pancreatic lipase among the common phenolic acids (Karamać and Amarowicz, 1996). It is also a potent inhibitor of α-glucosidase and α-amylase with IC50 values of 4.9 mM and 9.5 mM, respectively (Adisakwattana et al., 2009, Jeong et al., 2012). In vivo studies have shown that ferulic acid was able to help regulate the blood sugar levels in diabetic rats (Balasubashini et al., 2004). Furthermore, a combination of ferulic acid and resveratrol have been shown to mitigate the damage of liver, kidney and pancreas in diabetic rats, possibly by reducing inflammation (Ramar et al. 2012). In addition, Alam et al. (2013) reported that ferulic acid can enhance the function and structure of the blood vessels, heart, liver and kidneys in rats suffering hypertension. Ferulic acid occurs in sea parsley only at a moderate concentration of 25.62 ±0.22 mg/g extract DW; nevertheless, it is expected to make a moderate contribution to the antioxidant capacity, the inhibitory effects on the metabolic enzymes, as well as the potential health benefits of the plant.

ρ-Coumaric acid is found at a relatively low concentration in sea parsley compared with the other two phenolic acids. ρ-Coumaric acid is widely found in a variety of foods, including fruits and vegetables, whole cereal grains and nuts (Kováčová and Malinová, 2008). It has also been identified in the Australian native plants including Tasmannia pepper, bush tomato and mint bush at low levels (Konczak et al., 2010, Tang et al., 2017). A number of studies have shown that ρ-coumaric acid is a potent antioxidant with a strong free radical scavenging capacity (Castelluccio et al., 1995, Zang et al., 2000, Rao & Muralikrishna, 2004, Ferguson et al., 2005, Kováčová and Malinová, 2008, Gani et al., 2012, Roy & Prince, 2013, Yoon et al., 2013, Kiliç and Yeşiloğlu, 2013, Krishina et al., 2014). Owing to its strong antioxidant activity, ρ-coumaric acid has been reported to decrease the formation of the carcinogenic nitrosamines in the stomach (Ramadoss et al., 2015). Other studies have shown that it exhibits anti-inflammatory, anticancer (Wadhwa et al., 2016), neuroprotective (Vauzour et al., 2010), hepatoprotective (Venkatesan et al., 2011) and antidiabetic (Amalan et al., 2016) effects.

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Luteolin is found to be present in trace amount in sea parsley. Luteolin has been found in many vegetables including parsley )50+ mg/100 g FW(, celery )5-10 mg/100 g FW( and green peppers )3.9 mg/100 g FW( and herbs such as rosemary )< 5 mg/100 g FW( while fruits such as oranges, grapefruit, and lemons are especially rich in this polyphenol (Mecocci et al., 2014, Reza, 2016). Belonging to the subgroup of flavone in the flavonoids group, luteolin possesses four hydroxyl groups at carbons 5, 7, 3’, and 4’ positions, which gives it a strong antioxidant activity (Ross and Kasum, 2002). The hydroxyl moieties and the 2−3 double bond of luteolin are believed to be closely linked to its biological activities, including antioxidant, anticancer, anti-inflammation and anti-allergy properties (Chan et al., 2003, Galati et al., 2004, Lin et al., 2008). Luteolin has been reported to exhibit several other health functions including enhancement of muscle function and improvement of the nerve function (Reza, 2016).

Catechin was also detected in trace amount in sea parsley (< 1 mg per g purified extract). Belonging to the subgroup of flavanols, catechins have been found widely identified in numerous foods, especially fruits such as apples, blueberries, gooseberries, grape seeds, kiwi and strawberries, but also in cocoa, chocolate, green tea and red wine. Catechin is well documented to be a potent antioxidant that can decrease the generation of free radical, lipid peroxidation, oxidative stress and oxidation of low-density lipoprotein (LDL) cholesterol (Suksamrarn et al., 2003, Mandel et al., 2005, Augustyniak et al., 2010). It has also been shown to exert anti-aging effect and may help avert cardiovascular complications (Suksamrarn et al., 2003, Mandel et al., 2005, Augustyniak et al., 2010). These minor phenolic components of sea parsley, even though present at relatively low concentrations, compared with apiin, apigenin and caffeic acid, would nevertheless contribute to the biological activities and health functions of sea parsley.

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6.3. Conclusion

This chapter presented the identification and quantification of the phenolic compounds present in the methanolic extract of sea parsley (Apium prostratum) using a 4-step identification and confirmation process. The first three steps are: 1) comparison of HPLC retention times, 2) co-eluding and 3) comparison of UV absorption maxima with reference standards, which were accomplished with HPLC-DAD. The compounds tentatively identified were then confirmed by comparing their MS spectra with those of reference standards by using liquid chromatography-high resolution mass spectrometry (LC-HRMS) analysis. Seven phenolic compounds were identified in this Australian native plant species, with the main ones being apiin, apigenin, caffeic acid and ferulic acid, while the minor compounds were ρ-coumaric acid, luteolin and catechin. The phenolic fraction of the plant was dominated by apiin and apigenin, which accounted for 48.2% and 24.8% of the total phenolic content, respectively. These compounds, especially apiin, apigenin and caffeic acid, are expected to contribute to the biological activities of sea parsley, including antioxidant capacities and inhibition activities on metabolically important enzymes, as reported in Chapters 4 and 5.

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CHAPTER 7 Conclusions and Recommendations

This MPhil study was carried out with the broad aim to extend the knowledge on the phenolic composition and health-promoting properties of Australian native plants. Three plant species were selected for study, namely saltbush (Atriplex nummularia), samphire (Sarcocornia quinqueflora) and sea parsley (Apium prostratum), which have not been investigated previously. The key conclusions of the study are presented in the following sections together with some recommendations for future research.

The project began with a determination of the proximate composition of the three plants. The moisture levels of the three plants varied from 76.9% to 84.7% (w/w). The ranges of the other main components for the three plants were: protein (2.8-5.4%), lipids (0.6- 8.2%), ash (4.3-5.9%) and carbohydrates (2.21-3.40%). Sea parsley had the highest levels of moisture, lipids and carbohydrates; saltbush had the highest level of protein while samphire had the highest ash content. The proximate compositions of the three plants were broadly similar to most vegetables.

Next, the phenolic components of the three plants were extracted with 80% (v/v) methanol and purified by liquid chromatography using XAD-7 Amberlite® resin. The total phenolic contents of the extracts were determined by the Folin-Ciocalteu procedure and their antioxidant activities were measured by three methods, namely ABTS and DPPH free radical scavenging capacity assays and Ferric reducing antioxidant power (FRAP) assay. The three native Australian plants possessed relatively high levels of phenolic contents and antioxidant activities when compared with Australian native plants in general. Of the three plant samples, sea parsley contained the largest amount of phenolic compounds and exhibited the highest antioxidant capacities when determined by the ABTS and DPPH free radical scavenging capacity assays. Saltbush had the lowest amount of phenolic compounds and displayed the lowest antioxidant activities as assessed by all three assay methods. Samphire had the second highest phenolic content and also showed the greatest FRAP value. Purification of the extracts by liquid chromatography with Amberlite® resin (XAD-7) resulted in a significant concentration of the phenolic compounds with the total phenolic content increased by 1.43-2.67 times in the purified

111 extracts. Similarly, the purification caused a marked increase in the antioxidant activity of the extracts, with the antioxidant capacities increased by 1.41-2.38 times depending on the plants and assay methods. Correlation analysis showed that the phenolic content was significantly correlated with the antioxidant capacities as measured by all three assays as well as between the results of ABTS and DPPH free radical scavenging capacity assays (p<0.01), but neither was correlated with the FRAP assay results.

The project then moved to the determination of the inhibitory activities of the plant extracts on four metabolically important enzymes, namely α-glucosidase, α-amylase, pancreatic lipase and hyaluronidase. The extracts were found to exhibit significant, concentration-dependent inhibitory effects on these enzymes. The inhibitory effects followed negative exponential curves and the IC50 values of the inhibition are comparable to many plant foods including several native Australian fruits and herbs. Overall, the three plants were stronger inhibitors of pancreatic lipase than the other enzymes. Of the three plants, sea parsley exhibited the highest inhibitory activities on α-glucosidase, α-amylase and pancreatic lipase for both the crude and purified extracts, followed by samphire. Samphire exhibited the highest inhibitory activity against hyaluronidase, followed by sea parsley, while saltbush extracts were the least potent inhibitor on all the enzymes. Purified extracts showed greater inhibitory effects than the crude extracts in all cases and the inhibitory activities were generally correlated with the phenolic contents and antioxidant capacities of the plant samples. This demonstrated that phenolic compounds were the main components responsible for the enzyme inhibitory effects.

In the final chapter of the thesis, the phenolic compounds in the purified extract of sea parsley were identified and quantified by a combination of HPLC-PDA and LC-HRMS analyses. Sea parsley was selected for the analyses because it exhibited the highest overall biological activities among the three plants, and due to the limited scope of this study. The identification of the phenolic compounds in the extract of sea parsley followed a 4- step process with the first three steps being: 1) comparison of HPLC retention times, 2) co-eluding; and 3) comparison of UV absorption maxima with reference standards, which were accomplished with HPLC-DAD. The compounds tentatively identified were then confirmed by comparing their MS spectra with those of reference standards by using liquid chromatography-high resolution mass spectrometry (LC-HRMS) analysis. Seven phenolic compounds were identified in this Australian native plant species, with the main 112 ones being apiin, apigenin, caffeic acid and ferulic acid, while the minor compounds were ρ-coumaric acid, luteolin and catechin. The phenolic fraction of the plant was dominated by apiin and apigenin, which accounted for 48.2% and 24.8% of the total phenolic content, respectively. These compounds, especially apiin, apigenin and caffeic acid (6%), are expected to contribute significantly to the biological activities of sea parsley, including their antioxidant capacities and inhibitory activities on the four metabolically important enzymes. In summary, the three native Australian plants contained a relatively high level of phenolic compounds, possessed strong antioxidant capacities and exhibited potent inhibitory activities on four metabolically important enzymes. Furthermore, sea parsley contained seven phenolic compounds that have been shown to have the capacity to mitigate a number of diseases prevent in modern society. This means that consumption of these vegetables can bring significant health benefits to the consumer.

A number of future research directions can be suggested for further studies on this topic. First, due to the limited scope of this MPhil project, only phenolic compounds in sea parsley were identified and quantified. Future studies could seek to identify and quantify the phenolic compounds in saltbush and samphire as well. Second, the assessment of the health-related biological activities of the three plants was accomplished using in vitro methods. While this is usually the first step in assessing the health functions of plants, the results obtained could only be regarded as preliminary. The findings need confirmation by ex vivo studies using tissue cell lines, for example, the HepG2 cells. Such studies will provide information not only on whether the in vitro results are valid in a cellular environment but also the biosorption of the phenolic compounds by the tissue cells. Third, the strong bioactivities and the presence of phenolic compounds such as apiin and apigenin in high concentrations in sea parsley suggest that phenolic extract of this plant may have chemopreventive and therapeutic effects on certain diseases such as diabetes and cancer. Thus, it could be worthwhile to investigate such potential health benefits of sea parsley by in vivo methods using animal models, followed by clinic studies using human volunteers if promising results are obtained. Finally, as home to a huge range of unique plants, there are hundreds more native Australian plants that have traditionally served as a source of food and medicine for the indigenous people. This study, and those before it, have only covered a small proportion of this vast food and pharmaceutical resource. Future studies should continue to systematically investigate the phytochemicals

113 in more Australian native plants. Such investigations will not only serve to catalogue phytochemicals in Australian native plants and provide a more complete understanding of their bioactivities, but also may provide the opportunity to discover new phytochemicals with curative effects on human diseases, thus bring significant health benefits the human society.

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APPENDICES

Assay Equation R 2 Folin-Ciocalteu assay y = 0.0021x + 0.0316 0.9937 ABTS radical scavenging capacity y y = -0.0004x + 0.2057 0.9902 DPPH radical scavenging capacity y y = 0.0022x + 0.0289 0.9913 Ferric reducing antioxidant power y = 0.0008x + 0.0301 0.9956 α-Glucosidase inhibitory activity y =- 516.15x + 827 0.9799 α-Amylase inhibitory activity y = 0.0089x + 0.0013 0.9766 Pancreatic lipase inhibitory y = 538.3x + 1511.5 0.9859 Hyaluronidase inhibitory activity y = 2.5755x + 4.0928 0.9902

Appendix 1. Regression line equation for antioxidant and enzyme-based assays

141

mAU mAU

8 1500

7 A 1250 B

6 1000 5 750 4

500 3

2 250

1 0 250 260 270 280 290 300 nm 250.0 275.0 300.0 325.0 350.0 375.0 nm

mAU mAU(x100)

900 2.5 800 C D 700 2.0 600

500 1.5

400 1.0 300

200 0.5 100

0 0.0

300.0 325.0 350.0 375.0 400.0 nm 280 290 300 310 320 330 340 350 360 370 380 nm

mAU(x100) mAU(x100) 4.5

4.0 2.0

3.5 E 1.5 F 3.0

2.5 1.0

2.0 0.5 1.5

1.0 0.0

0.5 -0.5 0.0

250.0 275.0 300.0 325.0 350.0 375.0 400.0 nm 275.0 300.0 325.0 350.0 375.0 nm

mAU(x100) 8.0 7.0 G 6.0 5.0 4.0

3.0 2.0 1.0

0.0 275.0 300.0 325.0 350.0 375.0 400.0 nm

Appendix 2. Spectra of catechin (A), caffeic acid (B), ρ-coumaric acid (C), apiin (D), ferulic acid (E), luteolin (F), apigenin (G).

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A B

C D

E F

Appendix 3. Retention times of catechin (A), caffeic acid (B), ρ-coumaric acid (C), apiin (D), ferulic acid (E), luteolin (F), apigenin (G).

143

- Appendix 4. Comparison of molecular ion [M-H] (m/z 289) of catechin (C15H14O6) in Sea Parsley (Apium prostratum) (A) and the mass spectra of catechin standard (B) by LC- HRMS.

144

- Appendix 5. Comparison of molecular ion [M-H] (m/z 179) of caffeic acid (C9H8O4) in Sea Parsley (Apium prostratum) (A), the mass spectra of caffeic acid standard (B) by LC- HRMS.

145

Appendix 6. Comparison of molecular ion [M-H] - (m/z 163) of ρ-coumaric acid

(C9H8O3) in Sea Parsley (Apium prostratum) (A), the mass spectra of ρ-coumaric acid standard (B) by LC-HRMS.

146

- Appendix 7. Comparison of molecular ion [M-H] (m/z 563) of apiin (C26H28O14) in Sea Parsley (Apium prostratum) (A) and mass spectra of apiin standard (B) by LC-HRMS.

147

193.0507 NL: 100 9.78E6 MixSTD_4_20190117#1 90 97-221 RT: 3.55-3.93 AV: 25 F: FTMS - p APCI corona Full ms 80 [120.00-950.00]

40 178.0273 RelativeAbundance 223.0609

20 208.0378 195.0548 10 191.0564 197.0454 225.0655 181.0507 187.0402 205.0505 210.0419 219.0509 233.1545 237.0767 241.0504 0 193.0506 NL: 100 8.88E5

C 10 H 10 O4 +H: 90 C 10 H 9 O4 pa Chrg -1

80 A

197.0616 0 175 180 185 190 195 200 205 210 215 220 225 230 235 240 m/z

- Appendix 8. Comparison of molecular ion [M-H] (m/z 193) of ferulic acid (C10H10O4) in Sea Parsley (Apium prostratum) (A) and the mass spectra of ferulic acid standard (B) by LC-HRMS.

148

- Appendix 9. Comparison of molecular ion [M-H] (m/z 285) of luteolin (C15H10O6) in Sea Parsley (Apium prostratum) extract (A) and the mass spectra of luteolin standards (B) by LC-HRMS.

149

- Appendix 10. Comparison of molecular ion [M-H] (m/z 269) of apigenin (C15H10O5) in Sea Parsley (Apium prostratum) extract (A) and the mass spectra of apigenin standards (B) by LC-HRMS.

150