Studies on Clinacanthus Nutans

Genetic, phytochemical and bioactivity studies of Clinacanthus nutans (Burm. f.) Lindau (Acanthaceae)

A thesis submitted in fulfilment of the requirements for the degree of Doctor of Philosophy (Applied Biology and Biotechnology)

Siat Yee Fong

BSc (Biotechnology) (Monash), MAppSc (Molecular Biotechnology) (USYD)

School of Applied Sciences

College of Science, Engineering and Health

RMIT University

August 2015

Declaration

I certify that except where due acknowledgement has been made, the work is that of the author alone; the work has not been submitted previously, in whole or in part, to qualify for any other academic award; the content of the thesis is the result of work which has been carried out since the official commencement date of the approved research program; any editorial work, paid or unpaid, carried out by a third party is acknowledged; and, ethics procedures and guidelines have been followed.

Siat Yee Fong 31 August 2015

Acknowledgements

I thank my supervisors, Dr Tien Huynh, Associate Professor Terrence Piva and Dr Sylvia Urban for their supervision, advice, encouragement, help and support during my entire PhD tenure. Thank you for your insight and wisdom in manuscripts and thesis. I am also grateful to Professor Ann Lawrie, whose advice has been generously given on a number of occasions.

I acknowledge TKC farm, Kui Min Liau, Vun Jhit Lim and Rattanapong Charntawong for the provision of plant materials. My sincere thanks to Robert Brkljaca and Dr Chaitali Dekiwadia for their help in HPLC and TEM work, respectively. Thanks are also extended to Layla Mehdi Alhasan, Griff D’Costa, Xian Yang Chan, Alaa Banjar and Sushil Talda for providing the cells and reagents used in this study.

I am very appreciative for the provision of scholarship by the Ministry of Higher Education Malaysia and also the financial support from Universiti Malaysia Sabah. Without these, it would have been impossible for me to pursue my PhD.

Last but not least, I am extremely grateful to my parents, family and friends for their unfailing love and support, mentally and spiritually throughout. Sorry I had to make you listen to all my complaints. Special thanks to Dilani Wimalasiri for the friendship. I am thankful that we could share our ups and downs together during these 3 years.

Papers and communications arising from this study

FONG, S. Y., PIVA, T., URBAN, S. & HUYNH, T. 2014. Genetic homogeneity of vegetatively propagated Clinacanthus nutans (Acanthaceae). J. Med. Plants Res., 8, 903-914. http://www.academicjournals.org/journal/JMPR/article-abstract/629A21546085. (Presented as Chapter 2).

FONG, S. Y., PIVA, T., URBAN, S. & HUYNH, T. Variation in total phenolic and flavonoid contents and anticancer activity of Clinacanthus nutan (Burm. f.) Lindau (Acanthaceae) leaves against human D24 melanoma cells. (Manuscript).

FONG, S. Y., PIVA, T., URBAN, S. & HUYNH, T. Effect of postharverst processes on the total phenolic and flavonoid contents and anticancer activity of Clinacanthus nutans (Burm. f.) Lindau (Acanthaceae) against selected human cancer cell lines. (Manuscript).

FONG, S. Y. & HUYNH, T. Assessment of genetic variation between and within Clinacanthus nutans and Andrographis paniculata using RFLP, RAPD and microsatellite markers. RMIT School of Applied Sciences Annual Research Day, Melbourne, Australia. 14 June 2013.

FONG, S. Y., BRKLJACA, R., URBAN, S., PIVA, T., & HUYNH, T. Variation in total phenolic and flavonoid content in leaves of Clinacanthus nutans. 2nd International Conference and Exhibition on Pharmacognosy, Phytochemistry & Natural Products, Beijing, China. 25-27 August 2014.

FONG, S. Y., BRKLJACA, R., URBAN, S., PIVA, T. & HUYNH, T. Variation in total phenolic and flavonoid content in leaves of Clinacanthus nutans. RMIT SEH College HDR Student Conference, Melbourne, Australia. 17 October 2014.

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Table of Contents Declaration ...... i Acknowledgements ...... ii Papers and communications arising from this study ...... iii List of figures ...... x List of tables ...... xii List of Abbreviations and acronyms ...... xiii Abstract ...... xvii CHAPTER 1 ...... 1 Introduction and review of literature ...... 1 1.1 Importance of medicinal plants ...... 1 1.2 Acanthaceae ...... 3 1.2.1 Family characteristics ...... 3 1.2.2 Important members ...... 4 1.3 Clinacanthus nutans (Burm. f.) Lindau ...... 7 1.3.1 Vernacular names ...... 7 1.3.2 Geographical distribution/ecology ...... 7 1.3.3 Botanical characteristics ...... 7 1.3.4 Propagation ...... 9 1.3.5 Genetic diversity ...... 10 1.3.6 Phytochemical constituents ...... 16 1.3.7 Traditional medicinal uses of C. nutans ...... 37 1.3.8 Pharmacological activities of C. nutans ...... 38 1.3.9 Cancer ...... 42 1.4 Overview and aims of study ...... 58 CHAPTER 2 ...... 61 Genetic homogeneity of vegetatively propagated Clinacanthus nutans (Burm. f.) Lindau (Acanthaceae) ...... 61 2.1 Introduction ...... 61 2.2 Materials and methods ...... 64 2.2.1 Plant materials ...... 64 2.2.2 DNA extraction...... 66 2.2.3 PCR-RFLP ...... 66 2.2.4 RAPD amplification ...... 68

2.2.5 Microsatellite amplification ...... 68 2.2.6 Data collection and analysis ...... 69 2.3 Results ...... 70 2.3.1 PCR-RFLP analysis ...... 70 2.3.2 RAPD analysis ...... 71 2.3.3 Microsatellite analysis ...... 74 2.4 Discussion ...... 78 2.4.1 Genetic homogeneity in C. nutans ...... 78 2.4.2 Genetic differences between C. nutans, C. siamensis and A. paniculata ...... 80 2.4.3 Genetic variation in A. paniculata ...... 81 2.5 Conclusion ...... 82 CHAPTER 3 ...... 83 Variation in the total phenolic and flavonoid content of Clinacanthus nutans (Burm. f.) Lindau (Acanthaceae) leaves obtained from different locations ...... 83 3.1 Introduction ...... 83 3.2 Materials and methods ...... 86 3.2.1 Plant materials ...... 86 3.2.2 Preparation of crude extracts ...... 87 3.2.3 Determination of the TPC...... 88 3.2.4 Determination of the TFC...... 88 3.2.5 HPLC analysis ...... 89 3.2.6 Statistical analysis...... 90 3.3 Results ...... 90 3.3.1 Comparison of the TPC and TFC in leaf samples of C. nutans obtained from different locations within and between countries ...... 90 3.3.2 Correlation between the colorimetric and HPLC methods to investigate the TPC and TFC in C. nutans leaves ...... 93 3.4 Discussion ...... 96 3.4.1 The TPC varies in C. nutans leaves collected from different locations ...... 96 3.4.2 The TFC varies in C. nutans leaves collected from different locations ...... 98 3.4.3 Relationship between the colorimetric and HPLC methods ...... 100 3.5 Conclusion ...... 102

CHAPTER 4 ...... 103 Effect of drying temperature and extraction solvent on the total phenolic and flavonoid content of Clinacanthus nutans (Burm. f.) Lindau (Acanthaceae) leaves ...... 103 4.1 Introduction ...... 103 4.2 Materials and methods ...... 106 4.2.1 Plant materials ...... 106 4.2.2 Postharvest drying treatment ...... 107 4.2.3 Postharvest crude extraction ...... 107 4.2.4 Determination of the TPC...... 108 4.2.5 Determination of the TFC...... 109 4.2.6 HPLC analysis ...... 109 4.2.7 Statistical analysis...... 109 4.3 Results ...... 109 4.3.1 Effect of drying temperature on the TPC and TFC in C. nutans leaves ...... 109 4.3.2 Effect of extraction solvents on the TPC and TFC in C. nutans leaves...... 112 4.3.3 HPLC analysis and its correlation with the colorimetric methods ...... 114 4.4 Discussion ...... 118 4.4.1 Changes in the TPC and TFC in C. nutans leaves at different drying temperatures ...... 118 4.4.2 The TPC and TFC vary among the different crude extracts of C. nutans leaves .. 119 4.5 Conclusion ...... 121 CHAPTER 5 ...... 123 In vitro cytotoxic effect of different crude leaf extracts of Clinacanthus nutans (Burm. f.) Lindau (Acanthaceae) on selected human cancer cell lines and anticancer activity of the crude cold aqueous leaf extract against D24 melanoma cells ...... 123 5.1 Introduction ...... 123 5.2 Materials and methods ...... 127 5.2.1 Plant materials ...... 127 5.2.2 Preparation of crude extracts ...... 128 5.2.3 Cell line and culture conditions ...... 128 5.2.4 Cytotoxicity assay...... 128

5.2.5 Determination of half maximal effective concentration (EC50) ...... 129 5.2.6 Observation of D24 cell morphology by phase contrast microscopy ...... 130 5.2.7 Evaluation of apoptosis by Muse cytofluorimetric analysis ...... 130

5.2.8 Detection of apoptosis and necrosis by Annexin V/PI double staining and confocal microscopy...... 131 5.2.9 Ultra structural analysis by TEM...... 132 5.2.10 Statistical analysis...... 133 5.3 Results ...... 133 5.3.1 Cytotoxic effect of the different crude C. nutans leaf extracts on the melanoma and breast carcinoma cells ...... 133 5.3.2 Cytotoxicity of the crude cold and hot aqueous C. nutans leaf extracts against the D24 melanoma and NHDF cells ...... 135 5.3.3 Morphological changes in the D24 melanoma cells treated with the crude old aqueous C. nutans leaf extract ...... 137 5.3.4 Induction of mode of cell death in the treated D24 melanoma cells by the crude cold aqueous C. nutans leaf extract ...... 138 5.3.5 Ultrastructural changes in the D24 melanoma cells treated with the crude cold aqueous C. nutans leaf extract ...... 142 5.4 Discussion ...... 144 5.4.1 The crude C. nutans leaf extracts exhibit different effects on the viability of melanoma and breast carcinoma cells ...... 144 5.4.2 The crude cold and hot aqueous C. nutans leaf extracts exhibit selective cytotoxicity against the D24 melanoma cells ...... 146 5.4.3 The crude cold aqueous C. nutans leaf extract induces apoptosis in the treated D24 melanoma cells ...... 147 5.5 Conclusion ...... 149 CHAPTER 6 ...... 151 Comparative cytotoxicity of Clinacanthus nutans (Burm. f.) Lindau (Acanthaceae) leaves obtained from different locations and anticancer activity of the crude methanol leaf extract against D24 human melanoma cells ...... 151 6.1 Introduction ...... 151 6.2 Materials and methods ...... 154 6.2.1 Plant materials ...... 154 6.2.2 Preparation of crude extracts ...... 155 6.2.3 Cell line and culture conditions ...... 155 6.2.4 Cytotoxicity assay...... 155

6.2.5 Determination of EC50 ...... 155 6.2.6 Observation of D24 cell morphology by phase contrast microscopy ...... 156 6.2.7 Evaluation of apoptosis by Muse cytofluorimetric analysis ...... 156

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6.2.8 Detection of apoptosis and necrosis by Annexin V/PI double staining and confocal microscopy...... 156 6.2.9 Ultra structural analysis by TEM...... 156 6.2.10 Statistical analysis...... 156 6.3 Results ...... 156 6.3.1 Comparison of cytotoxicity among the crude methanol extracts of C. nutans leaves obtained from different locations against the D24 melanoma cells ...... 156 6.3.2 Cytotoxicity of the crude methanol C. nutans leaf extract against the D24 melanoma and NHDF cells ...... 158 6.3.3 Morphological changes in the D24 melanoma cells treated with the crude methanol C. nutans leaf extract ...... 160 6.3.4 Induction of mode of cell death in the treated D24 melanoma cells by the crude methanol C. nutans leaf extract ...... 161 6.3.5 Ultrastructural changes in D24 melanoma cells treated with the crude methanol C. nutans leaf extract ...... 165 6.4 Discussion ...... 167 6.4.1 Cytotoxicity against the D24 melanoma cells varies among the C. nutans leaves obtained from different locations and it correlates with the environment and concentration of phytochemical content ...... 167 6.4.2 The crude methanol C. nutans leaf extract exhibits selective cytotoxicity against the D24 melanoma cells ...... 168 6.4.3 The crude methanol C. nutans leaf extract induces apoptosis in the D24 melanoma cells ...... 169 6.5 Conclusion ...... 171 CHAPTER 7 ...... 173 Synopsis and future directions ...... 173 7.1 Influence of collection sites with different environmental characteristics on the phytochemical constituents and anticancer activity of C. nutans leaves ...... 173 7.2 Influence of postharvest processes on the phytochemical constituents and anticancer activity of C. nutans leaves ...... 175 7.3 Future directions ...... 177 References ...... 181 Appendices ...... 206 Appendix 1 ...... 206 Appendix 2 ...... 207 Appendix 3 ...... 208

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Appendix 4 ...... 209 Appendix 5 ...... 210

List of figures

Figure 1.1 Vegetative characteristics of C. nutans…………………………………... 8

Figure 1.2 Floral characteristics of C. nutans……………………………………….. 9

Figure 1.3 Biosynthetic pathways of secondary metabolites in plants……………… 17

Figure 1.4 Simplified pathway of the synthesis of phenolic compounds……………. 19

Figure 1.5 Classification of phenolic compounds…………………………………… 20

Figure 1.6 The phases of the cell cycle……………………………………………… 42

Figure 1.7 Morphological changes in apoptotic and necrotic cells………………….. 50

Figure 1.8 The extrinsic and intrinsic caspase activation cascades………………….. 51

Figure 2.1 Macromorphology of plants investigated in this study…………………... 63

Figure 2.2 Sampling sites of C. nutans, C. siamensis and A. paniculata in Malaysia, Thailand and Vietnam……………………………………………………. 64 Figure 2.3 PCR-RFLP patterns of Taq I and MspI digested rbcL gene……………... 71

Figure 2.4 RAPD amplification products of OPA-09, OPA-11 and OPA-18……….. 73

Figure 2.5 Microsatellite amplification products of GACA4 and GTG5…………… 76

Figure 2.6 UPGMA cluster analysis of the RAPD and microsatellite data, and PCA analysis of the combined RAPD and microsatellite data………………... 77 Figure 3.1 Collection sites of C. nutans samples from Peninsular and East Malaysia, Thailand and Vietnam………………………………………… 87 Figure 3.2 Scatter plots showing correlations between HPLC peak area and the TPC and the TFC in C. nutans leaves……………………………………. 94 Figure 3.3 HPLC chromatograms of crude methanol leaf extracts of C. nutans samples CE1and CE2 detected at 332 nm……………………………….. 95 Figure 3.4 Total peak areas of 11 C. nutans samples from HPLC method detected at 332 nm………………………………………………………………… 96 Figure 4.1 Collection site of CP2 from Peninsular Malaysia………………………... 106

Figure 4.2 Scatter plots showing correlations between the contents (TPC and TFC) in C. nutans leaves and postharvest processing factors as well as between the two contents of each factor…………………………………. 112 Figure 4.3 HPLC chromatograms of C. nutans crude methanol extract of the leaves dried at 40 and 80°C and crude ethanol and hot aqueous leaf extracts detected at 332 nm……………………………………………………….. 114 Figure 4.4 Total peak areas of leaf samples dried at different temperatures and extracted with five different solvents obtained from HPLC with UV detection at 332 nm………………………………………………………. 116 Figure 4.5 Scatter plots showing correlations between HPLC peak area and the TPC and TFC in C. nutans leaves treated with different drying temperatures and using different solvent extracts………………………... 117

Figure 5.1 Percentage of viable D24, MM418C1, MCF7 and BT474 cells treated with the different crude C. nutans leaf extracts………………………….. 134 Figure 5.2 Cytotoxic effect of the crude cold and hot aqueous C. nutans leaf extracts on the D24 and NHDF cells……………..……………………… 136 Figure 5.3 Phase contrast images showing morphological changes in the D24 cells treated with the crude cold aqueous C. nutans leaf extract……………… 137 Figure 5.4 Cytofluorimetric analysis of the effect of the crude cold aqueous C. nutans leaf extract on the D24 cells………………..…………………….. 139 Figure 5.5 Confocal images of the untreated cells and the induction of early and late apoptosis/necrosis in the D24 cells after treatment with the crude cold aqueous C. nutans leaf extract……………………………………… 141 Figure 5.6 Transmission electron micrographs of the untreated and treated D24 cells with the crude cold aqueous C. nutans leaf extract………………… 143 Figure 6.1 Effect of the crude methanol leaf extracts of C. nutans samples from 11 different locations on the viability of D24 cells…………………………. 157 Figure 6.2 Cytotoxic effect of the crude methanol C. nutans leaf extracts on the D24 and NHDF cells…………………………………………………….. 159 Figure 6.3 Phase contrast images showing morphological changes in the D24 cells treated with the crude methanol C. nutans leaf extract…………………... 160 Figure 6.4 Cytofluorimetric analysis of the effect of the crude methanol C. nutans leaf extract on the D24cells……..……………………………………….. 162 Figure 6.5 Confocal images of the untreated cells and the induction of early and late apoptosis/necrosis in the D24 cells after treatment with the crude methanol C. nutans leaf extract………………………………………….. 164 Figure 6.6 Transmission electron micrographs of the untreated and treated D24 cells with the crude methanol C. nutans leaf extract…………………….. 166 Figure A1 Standard curves of gallic acid and quercetin…………………………….. 208

Figure A2 Scatter plots showing correlations between environmental factors and the TPC and TFC of C. nutans as well as between the TPC and TFC…... 209 Figure A3 Scatter plots showing correlations between cytotoxicity of the crude methanol leaf extracts of 11 C. nutans samples and different parameters, including elevation, annual mean temperature, TPC, TFC and phytochemical (in general) levels as determined by HPLC at 332 nm….. 210

List of tables

Table 1.1 List of medicinally important members of Acanthaceae and their traditional uses…………………………………………………………… 5 Table 1.2 A comparison of the different types of molecular marker techniques…… 12

Table 1.3 Molecular markers used in genetic studies of Acanthaceae members…… 13

Table 1.4 Phytochemical compounds present in different parts of C.nutans………. 23

Table 1.5 HPLC parameters used for the detection and quantification of compounds in C. nutans…………………………………………………. 35 Table 1.6 Pharmacological activities of C. nutans…………………………………. 39

Table 1.7 Oncogenes, tumour suppressor genes and their involvement in related cancers………...…………………………………………………………. 44 Table 1.8 Representatives of Acanthaceae able to induce apoptosis in human cancer cell………………………………………………………………... 48 Table 1.9 Plant-derived drugs currently used in cancer treatment………………….. 55

Table 2.1 The taxon, sample codes, collection sites, environmental and growth conditions of collected samples………………………………………….. 65 Table 2.2 Restriction endonuclease digestions of the rbcL 1F and 724R PCR products…………………………………………………………………... 67 Table 2.3 The five primers and the number of corresponding fragments used in the RAPD and microsatellite analyses……………………………………….. 69 Table 2.4 Restriction endonuclease digestions of rbcL 1F-724R PCR products…… 70

Table 3.1 The TPC and TFC of 11 C. nutans samples from different locations…… 92

Table 4.1 The TPC and TFC in C. nutans leaves dried at different drying temperatures and extracted with different solvents……………………… 110 Table 5.1 Comparison of morphological features of apoptosis and necrosis………. 127

Table 6.1 Melanoma therapeutic compounds obtained from natural sources (dietary components, phytochemicals and crude extracts)………………. 153 Table A1 Genetic similarity matrix generated from the RAPD data……………….. 206

Table A2 Genetic similarity matrix generated from the microsatellite data………... 207

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List of Abbreviations and acronyms

7-AAD 7- aminoactinomycin D

AFLP Amplified fragment length polymorphism

AlCl3 Aluminium chloride

ANOVA Analysis of variance bp Base pair

BRAF Beta rapidly accelerated fibrosarcoma gene

BT474 Human breast carcinoma cell line

CCK-8 Cell counting kit-8 cH2O Cold aqueous/water

CHS Chalcone synthase

CO2 Carbon dioxide d Day

D24 Human melanoma cell line

DAD Photodiode array detector dc Column internal diameter

DCM Dichloromethane

DMSO Dimethyl sulfoxide

DNA Deoxyribonucleic acid

DPPH 2,2-diphenyl-1-picrylhydrazyl dp Particle size

EC50 Half maximal effective concentration

ELSD Evaporating light scattering detector

Elv Elevation

ERK Extracellular signal-regulated kinase

EtBr Ethidium bromide

EtOH Ethanol

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FBS Foetal bovine serum

FITC Fluorescein isothiocyanate

FRAP Ferric reducing antioxidant power g Gram

GAE Gallic acid equivalent h Hour hH2O Hot aqueous/water

HPLC High Performance Liquid Chromatography

ITS Internal transcribed spacer kg Kilogram

L Litre l Length

M Molar m metre

MAPK Mitogen-activated protein kinase mAU Milli absorption unit

MCF7 Human breast carcinoma cell line

MEK Methyl ethyl ketone

MeOH Methanol mg Milligram

Min Minute

MIC Minimum inhibitory concentration mL Millilitre mm Millimetre mM Millimolar

MM418C1 Human melanoma cell line

MS Mass spectrometry

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MTT 3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyltetrazolium bromide

Na2CO3 Sodium carbonate ng Nanogram

NHDF Normal human dermal fibroblast cell line

NK Natural killer cell line nM Nano molar nm Nanometre

NMR Nuclear magnetic resonance

NS Not significant p53 Protein 53 gene

PAL Phenylalanine ammonia lyase

PBS Phosphate-buffered saline

PCA Principal component analysis

PCR Polymerase chain reaction

PI Propidium iodide

PO Peroxidase

PPO Polyphenoloxidase

PS Phosphatidylserine

PTFE Polytetrafluoroethylene

QE Quercetin equivalent r Pearson’s correlation coefficient

RAPD Random amplified polymorphic DNA rbcL Large subunit of the ribulose-bisphosphate carboxylase gene

RFLP Restriction fragment length polymorphism

Rn Rainfall rpm Revolutions per minute s Second

SAHN Sequential, agglomerative, hierarchic and non-overlapping

SD Standard deviation

SimQual Similarity for qualitative

SSR Simple sequence repeat

TBE Tris/borate/EDTA buffer

TEM Transmission electron microscopy

TFC Total flavonoid content

Teq Trolox equivalent

Tm Temperature

TPC Total phenolic content

UPGMA Unweighted pair group method with average mean

UV Ultraviolet

UV-Vis Ultraviolet-visible v/v Volume for volume

WST Water-soluble tetrazolium salts w/v Weight for volume w/w Weight for weight

°C Degree celcius

µg Microgram

µL Microlitre

µm Micrometre

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Abstract

There are approximately 300,000 higher plant species in the world, but only about 10% of these have been studied extensively for possible medical applications. Therefore, these plant species have significant potential for studies on their phytochemical constituents and bioactivities. These include Clinacanthus nutans (Burm. f.) Lindau (Acanthaceae), a popular medicinal plant native to Southeast Asia, traditionally used to treat infections and several diseases. The plant is normally propagated vegetatively by stem cuttings to meet consumer demand. C. nutans leaves are often prepared in dried form to prolong storage life and the dried leaves are commonly steeped in boiling water and then consumed as a herbal tea.

The level of phytochemicals in plants varies even amongst the members of the same species.

This variation can be influenced by genetics or environment alone, or as a combination of the two factors. Besides, postharvest processing of medicinal plants, such as drying and extraction, can also affect the level of phytochemicals in plants, and hence, their bioactivities.

However, there is still a lack of information on how and to what extend these factors affect the phytochemical constituents in C. nutans leaves, which may consequently influence its bioactivity. Therefore, this study aimed at addressing these knowledge gaps by investigating the genetic variation, total phytochemical content and anticancer activity of C. nutans using a multidisciplinary approach that integrates molecular genetics, analytical chemistry and cell biology.

The leaves of C. nutans grown at higher elevations with lower air temperatures (i.e. CT5 from Chiang Dao, Chiang Mai, Thailand) had higher total phenolic and flavonoid content as determined using the Folin-Ciocalteu, aluminium chloride and high performance liquid chromatography (HPLC) methods, compared to those grown at lower elevations with higher

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air temperatures. Likewise, the crude methanol extracts of C. nutans leaves from higher elevations and lower air temperatures exhibited higher cell cytotoxicity, with the leaf extract from Chiang Dao, Chiang Mai, Thailand, showed the highest activity (24 h EC50: 0.95 mg/mL and 72 h EC50: 0.77 mg/mL). This extract also showed selective cytotoxicty against cancer cells but not non-cancerous cells. The extract induced apoptosis in the D24 melanoma cells as determined by cytofluorimetry and microscopy (confocal and transmission electron).

Genetic study of C. nutans using molecular markers revealed high genetic similarity, with an average of 77.0% (RAPD) and 91.6% (microsatellite) among the vegetatively propagated C. nutans samples from Malaysia, Thailand and Vietnam, with identical genetic profiles even though they were geographically distant. These results suggest that the anticancer activity of the C. nutans leaf extract against D24 melanoma cells may be determined by its phytochemical constituents, which are influenced by environmental variables, particularly elevation and air temperature, rather than genetic factors.

The effect of postharvest treatments, including drying temperature and extraction solvent, on the phenolic and flavonoid content in C. nutans leaves was investigated using the same colourimetric and HPLC methods. An increase in the level of phytochemical constituents was observed with increasing drying temperatures, where leaves dried at 100°C and 80°C had the highest total phenolic and flavonoid content, respectively. HPLC analysis also showed the greatest amount of phytochemicals in the C. nutans leaves dried at 80°C. This event may be resulted from the formation of phenolic compounds through non-enzymatic reactions and/or a decrease in peroxidase and polyphenoloxidase enzymes activities. This suggests that low temperature drying may not be a suitable postharvest method for drying C. nutans leaves. All methods showed that the most polar solvent (water) resulted in the highest total phytochemical content being extracted from the leaves. The crude cold aqueous C. nutans

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leaf extract had the greatest quantities of both phenolics and flavonoids, suggesting that cold extraction (22°C) with water is the best method to prepare the crude extract of C. nutans leaves with maximum phenolics and flavonoids. Only the effect of extraction solvent on the viability of four human cancer cell lines (melanoma: D24 and MM418C1 cells and breast carcinoma: MCF7 and BT474 cells), was examined in the current study. The D24 melanoma cell line was the most sensitive to treatment and the crude aqueous (cold and hot) leaf extracts were the most active against all cancer cell lines, whereas the methanol, ethanol and dichloromethane extracts showed little or no cytotoxicity against these cell lines. The crude cold aqueous extract was the most cytotoxic against the D24 melanoma cells with a 72 h EC50 of 1.63 mg/mL. This is in agreement with that of the colourimetric methods, suggesting that the extraction with water at 22°C is the best method to extract the bioactive compound with anticancer property from C. nutans leaves, which remains to be isolated and identified. The extract also predominantly induced apoptosis in the D24 cells according to the cytofluorimteric, confocal and transmission electron microscopic analyses. This study provides important information on the optimisation of postharvest methods to prepare crude extracts that preserve the bioactive phytochemicals from dry C. nutans leaves. This may increase the potential of C. nutans as a valuable medicinal plant.

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CHAPTER 1

Introduction and review of literature

1.1 Importance of medicinal plants

The word “natural” is defined as something that is present in or produced by nature and not artificial or synthetic (Spainhour, 2005). Natural products originate from plants, animals and microbes (Nakanishi, 1999). The use of natural products as medicine is as ancient as human civilisation and the utilisation of natural products as sources of novel structures, but not necessarily the final drug entity, is still active and thriving (Newman and Cragg, 2012). In fact, up to 50% of the approved drugs during the last three decades are from natural products

(Cragg and Newman, 2005, Newman and Cragg, 2012).

The discovery of compounds from plants as drugs started at the beginning of the 19th century when morphine was first isolated from the opium poppy, Papaver somniferum, followed by other important drugs, including quinine from Cinchona officinalis, berberine from Berberis spp., aspirin from Salix spp., pilocarpine from Pilocarpus mycrophyllus and salicin from

Populus spp. (Johns, 1990). Since then, there has been an increased interest in botanical investigations to explore novel bioactive phytochemical compounds from plants for drug discovery. Higher plants and their derivatives represent no less than 25% of the total clinical drugs prescribed worldwide (Gurib-Fakim, 2006). There are approximately 300,000 higher plant species in the world (Kricher, 2011). However, only 25,000 plant species have been studied for possible medical applications (Efferth et al., 2008). Of these 25,000 species, only about 15% of them have been studied phytochemically, where less than 5% were screened for at least one biological activity (Ghosh and Jha, 2008). Therefore, plants still provide a largely

CHAPTER 1 unexploited resource for the discovery and development of new drugs, thus providing opportunities to new areas of research.

Moreover, the utilisation of medicinal plants to treat various ailments by people in developing nations is important, especially rural populations. Approximately 80% of citizens in Asia,

Africa and Middle East, still rely on traditional medicines, including herbal medicines, for their primary health care needs, due to restricted or no access to modern health facilities and limited funds to pay for high medical expenses (Ekor, 2013). While pharmaceutical medicines are commonly used in developed countries to treat a range of diseases, traditional medicines have grown popular in these countries in recent decades. For instance, between

70% and 90% of populations in Canada, France, Germany, and Italy have partaken traditional medicines in their lives (Robinson and Zhang, 2011).

The search for effective and novel therapeutic agents from plant entities for the treatment of various diseases or conditions, such as oxidative stress, inflammation, bacterial and viral infections, allergies, cancer and other pathological complications remains the pursuit for medicines (Chua, 2007). Huge efforts have been made in studying plants, especially those from diversity-rich regions, for their biological effects or characterising and isolating active components for the treatment of diseases. However, in some countries, scientific empirical- based evidence on biological actions of most traditional plants is still lacking due to limited resources, such as necessary equipments and expertise (Chua, 2007). This includes

Clinacanthus nutans (Burm. f.) Lindau, a member of the family Acanthaceae, which has been traditionally used in Asia for decades but its medicinal profile still remains unestablished because it is one of the least investigated medicinal plants in the continent (Pratoomsoot et al., 2015).

CHAPTER 1

1.2 Acanthaceae

1.2.1 Family characteristics

The Acanthaceae is a large dicotyledonous flowering plant family in the Lamiales order, which comprises approximately 220 genera and 4,000 species (Scotland and Vollesen, 2000,

Deng et al., 2011). Acanthaceae consists of four subfamilies, namely Nelsonioideae,

Acanthoideae, Thunbergioideae and Avicennioideae (Stevens, 2015). Most members are pantropical and subtropical species found in Indo-Malaysia, Africa, Madagascar, Brazil and

Central America, with a few species distributed in temperate regions, including the

Mediterranean, Australia, the United States and China (Mabberley, 1998, Deng et al., 2011).

Most of the species are shrubs, vines or herbaceous (annual or perennial) plants, although a few are tropical tree species (Meyer and Lavergne, 2004). Being a diverse family,

Acanthaceae has several common characteristics among its members. Most have simple leaves arranged in opposite pairs, exstipulate and usually entire-margined, with round to quadrangular stems. Cystoliths (inorganic concretions) are usually present in streaks on leaves or protuberances in the vegetative parts. The bisexual flowers are terminally or axillary spikes, racemes or cymes, with large and showy involucral bracts. Sepals and petals, which can be sub-actinomorphic or zygomorphic, are in five or four each, often fused into tubular structures. There are usually two or four stamens that extend beyond the mouth of the flower, often with the number of staminodes ranging between one and three. The pistil is superior and generally consists of two fused carpels, enclosing two locules, each of which has two to many ovules in two rows along the central axis of the ovary. Fruits are loculicidal, explosive dehiscent capsules, containing a few seeds, which are usually borne on hooklike retinacula

(Deng et al., 2011).

CHAPTER 1

1.2.2 Important members

Most members of Acanthaceae are used as ornamentals because of their numerous flowers or bracts with showy colours and bicolourous foliage (Meyer and Lavergne, 2004). In fact,

Acanthaceae is one of the most popular ornamental families along with Fabaceae (Whistler,

2000). Some of the species famous for its bicolorous foliage and showy bracts or flowers include Graptophyllum pictum, Hypoestes phyllostachya, Pachystachys lutea,

Pseuderanthemum carruthersii var. artropurpureum, Strobilanthes dyerianus and

Thunbergia mysorensis (Meyer and Lavergne, 2004).

Besides ornamental value or applications, several species are widely used by many ethnic communities as traditional medicine throughout the world, particularly in Africa and tropical

Asia (Fongod et al., 2013, Awan et al., 2014, Mahbubur Rahman et al., 2014). The most commonly used parts are the leaves to treat various ailments and diseases (Table 1.1).

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Table 1.1. List of medicinally important members of Acanthaceae and their traditional uses. Species Part(s) used Traditional uses Reference(s) Acanthus ilicifolius Leaf, root Aphrodisiac, rheumatism, neuralgia, pain-relief, blood Van Kiem et al. (2008) tonic, expectorant, leucorrhoea , cough, asthma Saranya et al. (2015) Acanthus montanus Leaf Cough, pneumonia, fever Fongod et al. (2013) Acanthus pubescens Leaf Dry cough, pneumonia, chronic asthma, cancer, Jeruto et al. (2008) tonsils, flu Adhatoda zeylanica Leaf Asthma, cough, cold, fever Ignacimuthu et al. (2006) Andrographis lineata Leaf Diabetes, scorpion stings, snake bites, fever, jaundice, liver ailments Renugadevi et al. (2013) Andrographis paniculata Leaf Snake bites, insect stings, diabetes, menstrual Ignacimuthu et al. (2006) disorders, malaria, filarial, fever, headache, vertigo Muthu et al. (2006) Mia et al. (2009) Renugadevi et al. (2013) Asystasia gangetica Leaf Burns, stimulate appetite Ignacimuthu et al. (2006) Muthu et al. (2006) Asystasia schimperi Leaf Cough, skin diseases Jeruto et al. (2008) Barleria grandiflora Whole plant Skin infections Kumari et al. (2015) Barleria lupulina Whole plant Mental stress, diabetes, rheumatoid, arthritis, snake Mazumder et al. (2012) bites Barleria prionitis Leaf, root, bark Fever, toothache, liver ailments, ulcers, piles, boils, Amoo et al. (2009) glandular swelling, cough, diuretic Jaiswal et al. (2014) Blepharis maderaspatensis Leaf Nervous disorders, cut wounds Ignacimuthu et al. (2006) Brillantaisia nitens Leaf Asthma Fongod et al. (2013) Clinacanthus siamensis Leaf Relieve painful swelling Kunsorn et al. (2013) Dipteracanthus prostratus Leaf Gonorrhoea, ear diseases Mahbubur Rahman et al. (2014) Dyschoriste radicans Leaf Skin diseases, wounds, eye infections Jeruto et al. (2008) Dyschoriste thumbergiiflora Leaf Skin diseases, wounds, eye infections Eremomastax speciosa Whole plant Convulsion, anaemia, headache, fever, malaria, herpes, Fongod et al. (2013) dysentery, diarrhoea, yellow fever, labour pains Elytraria acaulis Leaf Hip pain Ignacimuthu et al. (2008) Hemigraphis colorata Leaf Wounds Anitha et al. (2012)

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Table 1.1. List of medicinally important members of Acanthaceae and their traditional uses (continued). Species Part(s) used Traditional uses Reference(s) Hygrophila spinosa Leaf, root, seed Cancer Mumtaz et al. (2015) Hypoestes rosea Whole plant Typhoid Fongod et al. (2013) Nelsonia canescens Leaf Ear ache Mahbubur Rahman et al. (2014) Justicia adhatoda/Adhatoda Leaf, flower, fruit, root Cold, cough, chronic bronchitis, asthma Muthu et al. (2006) vasica Jha et al. (2012) Justicia betonica Leaf, flower Cough, diarrhoea, orchitis Jeruto et al. (2008) Justicia diffusa Leaf Skin diseases, toothache Ignacimuthu et al. (2008) Justicia flava Leaf Ulcers, pneumonia Jeruto et al. (2008) Justicia gendarussa Leaf, root Wounds, male contraceptive agent, chronic Mia et al. (2009) rheumatism, inflammation, bronchitis, headache, Ayob et al. (2014) arthritis, vaginal discharges, dyspepsia, eye diseases, fever Justicia tranquebariensis Leaf Poisonous bites and stings Sandhya et al. (2006) Lepidagathis scariosa Leaf Diarrhoea, wounds, oedema, pneumonia Jeruto et al. (2008) Phaulopsis falcisepala Whole plant Wounds, skin parasite, laxative, aphrodisiac Fongod et al. (2013) Phlebophyllus kunthianum Leaf, bark Nervous disorders Ignacimuthu et al. (2006) Rungia linifolia Whole plant Ulcer Ignacimuthu et al. (2008) Rungia pectinata Leaf, root Cooling, constipation, alleviate pain and swelling, Swain et al. (2008) as febrifuge Rungia repens Whole plant Cough, fever, vermifungal, diuretic, Tinea capitis Swain et al. (2008) Ruellia tuberosa Root Gonorrhoea, syphilis, renal infections, sore eyes Mahbubur Rahman et al. (2014) Thunbergia alata Leaf Cough, backache Jeruto et al. (2008) Thunbergia grandiflora Leaf, stem Stomach complaints, eye diseases, bladder stones, Mahbubur Rahman et al. (2014) elephantiasis Thunbergia laurifolia Leaf Menorrhagia, cuts, boils, antipyretic, antidote for Chan and Lim (2006) poisons

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1.3 Clinacanthus nutans (Burm. f.) Lindau

1.3.1 Vernacular names

Clinacanthus nutans (Burm. f.) Lindau is commonly known as Sabah snake grass in Malaysia, but the plant also has other vernacular names: belalai gajah (Malay), dandang gendis

(Javanese), phaya yo or phaya plongtong (Thai) and e zui hua or you dun cao (Chinese).

1.3.2 Geographical distribution/ecology

The medicinal plant is native to tropical and subtropical Asian countries, including Thailand,

Malaysia, Indonesia, Vietnam and China (Hu and Daniel, 2011, Chelyn et al., 2014). The plant is often cultivated by stem cuttings but wild populations are found in deciduous forests

(Mat Ali et al., 2010).

1.3.3 Botanical characteristics

1.3.3.1 Vegetative

The plant is a shrub reaching up to 1 m in height (Figure 1.1A). Stems are terete, striate and glabrescent (Figure 1.1B). Petioles are sulcate, bifariously pubescent and 0.3-2 cm long

(Figure 1.1B & C). Leaves are simple, opposite, with leaf blade having lanceolate-ovate, lanceolate or linear-lanceolate shapes, and a dimension of 7-12 cm length and 1-4 cm width

(Figure 1.1C & D). The leaf margin is subentire to sinuate-crenate. Leaf apex is acuminate, while the base is cuneate to rounded. Both leaf surfaces are pubescent when young, then becoming glabrescent when mature, except abaxially pilose along the veins. The number of secondary veins is 4-6 on each side of the midvein and abaxially elevated (Mat Ali et al.,

2010, Hu and Daniel, 2011).

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Figure 1.1. Vegetative characteristics of C. nutans. Shrub habit (A), terrete stem attached by petioles with leaves (B), simple lanceolate leaves (C) and opposite leaf arrangement (D).

1.3.3.2 Reproductive

Flowers of C. nutans are in dense cymes at or near the shoot apex (Figure 1.2A). The number of cymes can range from one to several, terminating with drooping horizontal stems but themselves erect, subsecund, and combined into a large lax, leafy panicle. Each flower has a

0.9-1.3 cm calyx with linear lobes and an acute apex, where the exterior is glandular- pubescent. The corolla is ~ 3.5 cm long, dull red with a green base and a basally cylindrical tube, ~ 3 mm in diameter, which gradually widens to ~ 6 mm at the opening. Flowers consist of a ~1.5 × 1 cm lower lip (turned upwards) with yellow streaks, apically yellow or greenish yellow, and an erect ~ 1.5 × 0.5 cm upper triangular lip with an emarginate apex (Figure

1.2B). The exterior is glandular-pubescent, whereas the interior is glabrous, except for a ring

CHAPTER 1 of trichomes ~ 5 mm above base. Each flower has two stamens, inserted in the throat and almost appressed against the upper lip with filaments 1-1.5 cm long and glabrous, while anthers are ovate-linear, muticous and have a dimension of 4-5 × 1 mm (Figure 1.2C). Style is filiform, 3-3.7 cm long with a sparsely pubescent base. Ovary is compressed, two-celled and consisting two ovules in each cell. Fruit is a capsule (~ 2 cm), pilose, oblong, basally contracted into a short, solid stalk and 4-seeded. Seeds are ~ 2 mm in diameter (Mat Ali et al.,

2010, Hu and Daniel, 2011).

† *

Figure 1.2. Floral characteristics of C. nutans. Flowers arranged in cymes (A) and flowers containing calyx (dash arrow), corolla with lower lip (asterisk) and upper triangular lip (dagger) (B), stamens (white arrow) and style (black arrow) (C).

1.3.4 Propagation

C. nutans is often vegetatively propagated by stem cuttings (Panyakom, 2006, Mat Ali et al.,

2010, Chen et al., 2015) because this method is easy, economical and produces a high multiplication rate. In addition, flowers are rare, which results in severely reduced sexual reproduction. The lack of flowers may be due to extended periods of vegetative propagation, selection and harvesting practices, where growers rapidly and constantly harvest the plant

CHAPTER 1 during the juvenile stage to meet consumer demand for this medicinal species. This may result in the inability of the plant to reach maturity for sexual reproduction.

Moreover, the popularity of C. nutans has prompted several investigators to perform in vitro tissue culture technique, which is considered to be a more rapid propagation method than stem cuttings in order to increase the supply. However, this technique is not easily achievable.

Ng (2013) and Gunasekaran (2014) showed success in callus induction of C. nutans but no plant regeneration was observed in both studies. Possible reasons include unsuitable experimental conditions, such as the concentration of growth regulators, suggesting the need for a better refined protocol to improve efficiency. Chen et al. (2015) reported the efficiency of shoot initiation to regenerate plantlet with Murashige and Skoog medium containing 1.0 mg/L benzyladenine and 0.02 mg/L 1-naphthaleneacetic acid, which showed about four times multiplication rate. Despite the high production rate of tissue culture, this technique may have negative long term impacts on the ecology of C. nutans, including inbreeding depression, disease resistance and inability to adapt to environmental changes, which may lead to increased risk of species extinction due to low genetic diversity.

1.3.5 Genetic diversity

The assessment of genetic diversity in plants is mainly based on morphological, biochemical and molecular traits (Govindaraj et al., 2015). However, molecular analysis is the preferred since morphological (e.g. floral structure, leaf, fruit and seed shape) and biochemical (e.g. allozymes and isozymes) characteristics are greatly influenced by environmental and growth conditions (Kumar et al., 2009, Lattoo et al., 2009). Thus, the consistency of results cannot be expected at all times for the same genotype (Kumar et al., 2009). Besides, results obtained from biochemical markers depends on the type of plant tissue (e.g. leaf vs. root) used for

CHAPTER 1 analysis (Kumar et al., 2009). This is because gene expression for a particular enzyme in one region may not be expressed in other regions (Kumar et al., 2009). In contrast, molecular markers can be selected based on the differences in the DNA sequences that are often not affected by environmental and growth factors. Also, molecular markers are able to detect more polymorphism than morphological and protein-based markers (Lattoo et al., 2009) and has been increasingly used in genetic diversity studies.

There are generally four types of DNA-based molecular marker techniques: i) restriction fragment length polymorphism (RFLP), ii) random amplified polymorphic DNA (RAPD), iii) microsatellite and iv) amplified fragment length polymorphism (AFLP) (Govindaraj et al.,

2015) (Table 1.2). These marker techniques are able to detect variations that arise from gene mutations, such as deletion, insertion, duplication and inversion in the chromosomes

(Paterson, 1996). Differences in the properties, such as the ability of these molecular makers to differentiate between homozygotes and heterozygotes, produce different estimates of genetic relationships among samples (Thormann et al., 1994). Therefore, careful thought in the selection of these techniques for genetic diversity studies is necessary to obtain more informative results. Molecular makers have been used to study the genetic diversity of a number of plant species, including members of Acanthaceae (Table 1.3).

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Table 1.2. A comparison of the different types of molecular marker techniques. Adapted from Kumar et al. (2009) and Govindaraj et al. (2015). Feature Molecular marker RFLP RAPD Microsatellite AFLP Principle of method Variation in the length of Variation in DNA The flanking regions of Combination of RFLPs DNA fragments produced fragments produced from microsatellites (1-10 bp) and PCR. DNA is digested by restriction PCR amplification of are used as primer binding by restriction enzymes endonucleases, which random segments of sites to amplify the DNA before the addition of identify particular genomic DNA with single region of interest. adapters that incorporate sequences and consistently primer or arbitrary Variation in the number of the primer sites for PCR. create double stranded cuts nucleotide sequence. microsatellite repeats The fragments are in the DNA. provides the difference selectively amplified by between individuals. extending the primers into the original fragment, and performing multiple PCR runs. Dominance Co-dominant Dominant (cannot Co-dominant Dominant (heterozygotes and distinguish heterozygotes homozygotes are from homozygotes) distinguishable) Prior sequence information No No Yes No Ease of use Difficult Easy Moderate Moderate Time required for detection Long Short Short Short Amount of DNA required High Low Low Low Level of polymorphism Moderate High High High Reproducibility High Low High High

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Table 1.3. Molecular markers used in genetic studies of Acanthaceae members. Taxon Marker(s) Genetic diversity level Reference(s) i) Genera Adathoda and Barleria RAPD Low Prasad (2014) Adathoda and Justicia RAPD Low to moderate Prasad (2014 Adathoda and Nigiranthus RAPD Low to moderate Prasad (2014 Adathoda and Hemigraphis RAPD Low to moderate Prasad (2014 Barleria and Justicia RAPD Low Prasad (2014 Barleria and Nigiranthus RAPD Low Prasad (2014 Barleria and Hemigraphis RAPD Low Prasad (2014 Clinacanthus and Andrographis RFLP, RAPD, microsatellite High Fong et al. (2014) Justicia and Nigiranthus RAPD Low Prasad (2014) Justicia and Hemigraphis RAPD Low Prasad (2014 Nigiranthus and Hemigraphis RAPD Low Prasad (2014 Suessenguthia and Sanchezia AFLP Low to high Schmidt-Lebuhn et al. (2005) ii) Interspecies Adathoda beddomei and A. zeylanica RAPD Low Prasad (2014) Clinacanthus nutans and C. siamensis RAPD, microsatellite Moderate Fong et al. (2014) RFLP No Fong et al. (2014) ITS sequencing Low Kunsorn et al. (2013) Sanchezia parvibracteata and Sa. oblonga AFLP Low Schmidt-Lebuhn et al. (2005) Sa. skutchii and Sa. parvibracteata AFLP Low Schmidt-Lebuhn et al. (2005) Sa. skutchii and Sa. oblonga AFLP Low Schmidt-Lebuhn et al. (2005) Suessenguthia barthleniana and Su. trochilophila AFLP High Schmidt-Lebuhn et al. (2005) Su. barthleniana and Su. koessleri AFLP High Schmidt-Lebuhn et al. (2005) Su. multisetosa and Su. wenzelii AFLP Low Schmidt-Lebuhn et al. (2005) Su. multisetosa and Su. vergasii AFLP Moderate Schmidt-Lebuhn et al. (2005) Su. multisetosa and Su. barthleniana AFLP Moderate Schmidt-Lebuhn et al. (2005) Su. multisetosa and Su. trochilophila AFLP High Schmidt-Lebuhn et al. (2005) Su. multisetosa and Su. koessleri AFLP High Schmidt-Lebuhn et al. (2005) Su. trochilophila and Su. koessleri AFLP Low to moderate Schmidt-Lebuhn et al. (2005)

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Table 1.3. Molecular markers used in genetic studies of Acanthaceae members (continued). Taxon Marker(s) Genetic diversity level Reference(s) Su. vergasii and Su. barthleniana AFLP Moderate Schmidt-Lebuhn et al. (2005) Su. vergasii and Su. trochilophila AFLP High Schmidt-Lebuhn et al. (2005) Su. vergasii and Su. koessleri AFLP High Schmidt-Lebuhn et al. (2005) Su. wenzelii and Su. vergasii AFLP Moderate Schmidt-Lebuhn et al. (2005) Su. wenzelii and Su. barthleniana AFLP Moderate Schmidt-Lebuhn et al. (2005) Su. wenzelii and Su. trochilophila AFLP High Schmidt-Lebuhn et al. (2005) Su. wenzelii and Su. koessleri AFLP High Schmidt-Lebuhn et al. (2005) iii) Intraspecies Acanthus volubilis RAPD Low Das et al. (2015) Andrographis paniculata RAPD Moderate Chua (2007) Minz et al. (2013) SRAP, SNP Low Chen et al. (2014b) RAPD, microsatellite No Sakuanrungsirikul et al. (2008) Avicennia germinans Microsatellite Variable degree Mori et al. (2010) Baphicacanthus cusia RAPD High Huang and Chen (2010) C. nutans RAPD, microsatellite No to low Fong et al. (2014) Hygrophila pogonocalyx RFLP Low Huang et al. (2005) Justicia areysiana RFLP, microsatellite High Meister et al. (2005) Justicia wrightii Microsatellite No genetic diversity Arbetan et al. (2005) within and among populations from the same location but there was genetic diversity between studied locations Ruellia nudiflora AFLP High Vargas-Mendoza et al. (2015) Su. multisetosa AFLP Low Schmidt-Lebuhn et al. (2005) Su. trochilophila AFLP Low to moderate Schmidt-Lebuhn et al. (2005)

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The genetic diversity of C. nutans at the intraspecies level was evaluated using RFLP, RAPD and microsatellite markers (Fong et al., 2014). However, RFLPs produced identical banding patterns among all C. nutans, suggesting that RFLP was unsuitable for the discrimination of

C. nutans at the intraspecies level. Low genetic diversity (or high genetic similarity) was found between C. nutans from Thailand, Malaysia and Vietnam using RAPDs and microsatellites even though they were geographically distant by land and water. The expected low genetic diversity in the species may be a consequence of the vegetative propagation method commonly employed to mass produce the plant. Vegetative or clonal propagation has reduced the genetic diversity in other plant families, such as Alternanthera philoxeroides

(Amaranthaceae) (Xu et al., 2003), Eichhornia crassipes (Pontederiaceae) (Li et al., 2006),

Allium sativum (Amaryllidaceae) (Paredes et al., 2008), Iris sibirica (Iridaceae)

(Kostrakiewicz and Wróblewska, 2008), Cypripedium calceolus (Orchidaceae) (Brzosko et al., 2011) and Chlorophytum borivilianum (Asparagaceae) (Tripathi et al., 2012).

C. nutans is often confused with Andrographis paniculata because of the similar common name i.e. Sabah snake grass vs. Indian snake grass. In Malaysia, C. nutans is commercially available as powdered or dried leaves and there is a possibility of contamination by other related plants. Misidentification and improper quality control of plants pre- and post- processing can lead to consumer problems, including adverse effects and reduced effectiveness due to their different biological activities (Ekor, 2013). Observation of plant morphological characteristics and taste are traditional identification methods (Li et al., 2011a), but these can be subjective to interpretation. Although both plants can be distinguished by the flowers, C. nutans rarely blooms. Therefore, other alternative identification methods are necessary. Molecular markers can offer an efficient alternative tool. For instance, RFLP,

RAPD and microsattelite markers have been used to differentiate C. nutans and A. paniculata,

CHAPTER 1 showing high genetic diversity between the two plants (Fong et al., 2014). However, not all molecular markers are useful for species discrimination, particularly for lower taxa levels.

For example, RFLP of the rbcL region could not discriminate between C. nutans and C. siamensis (Fong et al., 2014). The rbcL region is known to be highly conserved with low gene substitution rates in perennial angiosperm taxa (Bousquet et al., 1992, Frascaria et al.,

1993). This suggests that it is more suited for genetic studies of higher taxonomic levels

(intergeneric to subclass) but not lower levels (inter- to intraspecies) (Arca et al., 2012). Since there is only one study on the genetic diversity of C. nutans, further work is necessary to confirm the low genetic diversity found in the study. Besides, investigating the genetic diversity in larger populations may provide more information on the extent of diversity, allowing better understanding on the genetic profile of C. nutans.

1.3.6 Phytochemical constituents

Medicinal plants contain bioactive phytochemicals, defined as secondary metabolites that are produced to protect the plants. Since plants are stationary autotrophs, they need to develop survival strategies against a number of challenges, including engineering their own pollination and seed dispersal, nutrient deprivation, solar radiation and the coexistence of herbivores and pathogens in their immediate environment (Kennedy and Wightman, 2011).

Therefore, plants have evolved biochemical pathways for the production of secondary metabolites in vegetative (e.g. leaf, stem and root) and reproductive (e.g. flower, fruit and seed) regions in response to specific environmental stresses. They also serve as an attractant

(e.g. colour, pheromones) for pollinating insects or fruit-dispersing animals (Kennedy and

Wightman, 2011). These secondary metabolites are unique to specific plant species or genera and are not involved in the plants’ primary metabolic requirements (Harborne, 1993).

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Plant secondary metabolites are classified into three main groups based on their biosynthetic origin: i) terpenes, ii) nitrogen-containing alkaloids and iii) phenolic compounds (Croteau et al., 2002) (Figure 1.3). These secondary metabolites are synthesised from important building blocks (shikimic acid, acetyl coenzyme A, mevalonic acid and 1-deoxyxylulose-5-phosphate) via different pathways, such as shikimate, acetate, mevalonate and deoxyxylulose (Croteau et al., 2002) (Figure 1.3).

Figure 1.3. Biosynthetic pathways of secondary metabolites in plants. Adapted from Ribera and Zuñiga (2012)

Terpenes is the largest group of secondary metabolites, which consists of more than 30,000 lipid-soluble compounds, derived biosynthetically from units of isoprene (Kennedy and

Wightman, 2011). They possess at least one 5-carbon isoprene units. The number of attached isoprene units determine the classification of terpenes: 1 = hemiterpenes, 2 = monoterpenes, 3

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= sesquiterpenes, 4 = diterpenes, 5 = sesterpenes, 6 = triterpenes and 8 = tetraterpenes

(Kennedy and Wightman, 2011).

The second largest secondary metabolite group is the alkaloids, which consists of more than

12, 000 cyclic nitrogen-containing compounds that can be found in over 20% of plant species

(Kennedy and Wightman, 2011). This medicinally important group has no single classification, but alkaloids are commonly distinguished based on a structural similarity (e.g. indole alkaloids) or a common precursor (e.g. benzylisoquinoline, tropane, pyrrolizidine or purine alkaloids) (Kennedy and Wightman, 2011).

The third largest group is the phenolic compounds, which are synthesised via the phenylalanine precursor in the shikimate pathway (Figure 1.4). Phenolic compounds are characterised by having at least one aromatic ring bearing one or more hydroxyl groups, which can undergo esterification, methylation, etherification or glycosylation reactions

(Fresco et al., 2006). As a result, ~ 10, 000 structural variants, ranging from a simple molecule to a complex high-molecular polymer have been identified thus far (Kennedy and

Wightman, 2011). These compounds are generally categorised into different classes such as phenolic acids, flavonoids, stilbenes, coumarins, quinones, lignans, curcuminoids and tannins

(Figure 1.5). The categorisation is based on i) the nature and complexity of the basic carbonaceous skeleton; ii) the degree of skeletal modification (e.g. oxidation, hydroxylation, methylation, etc.); and iii) the link between the base unit and other molecules, including primary and secondary metabolites (Ewane et al., 2012).

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Figure 1.4. Simplified pathway of the synthesis of phenolic compounds. Adapted from Ewane et al. (2012).

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Figure 1.5. Classification of phenolic compounds.

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Of these phenolic compounds, flavonoids represent the largest and most diverse group, with ~

6, 000 identified compounds (Kennedy and Wightman, 2011). All flavonoids have the basic skeleton of a phenylbenzopyrone structure (C6-C3-C6) consisting of two aromatic (benzene) rings linked by a heterocyclic pyrane ring (Kumar and Pandey, 2013) (Figure 1.5). These compounds are further subdivided into flavones, flavonols, flavanones, flavanonols, isoflavones, flavan-3-ols, glycosylflavonoids, chalcones, aurones and anthocyanins, according to modifications of the basic skeleton (Bowsher and Tobin, 2008). Although not rare, C-glycosyl flavonoids are a special class of flavonoids due to their chemical structures and presumed different biosynthetic origin (Du et al., 2010). More than 300 of these compounds have been identified in plants (Bruneton, 1995). They are characterised by having a bond established between the asymmetric carbon on the sugar (often glucose, galactose or pentose) and the C6 or C8 of the aglycone, which is often a flavone and sometimes a flavonol or a chalcone (Bruneton, 1995).

Although phenolic compounds have gained momentum as a natural product pharmaceutical potential, terpenoids and alkaloids are more recognised and established as therapeutic agents

(Kinghorn, 1992). The most recognised drugs used in pharmaceuticals are terpenoids, including the anticancer drug, taxol, isolated from the bark of Taxus brevifolia (Taxaceae) and the antimalarial drug, artemisinin isolated from the leaves of Artemisia annua

(Asteraceae) (Mbaveng et al., 2014). Bioactive alkaloids isolated from plants include the antineoplastic agents, vinblastine and vincristine, isolated from the leaves of Catharanthus roseus (Apocynaceae) (Noble, 1990) and antipsychotic and antihypertensive drugs, reserpine, isolated from the roots of Rauvolfia serpentina (Apocynaceae) (Kinghorn, 1992).

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Rpresentatives of Acanthaceae, particularly A. paniculata and C. nutans, posseses a wide range of phytochemicals including terpenoids, alkaloids, flavonoids, glycosides, steroids, saponins, phenolic acids and tannins (Kaliswamy et al., 2015) (Table 1.4). Andrographolide, a bitter water soluble diterpene lactone, was isolated from the leaves of A. paniculata (Gorter,

1911). Two other diterpenes, deoxyandrographolide and neoandrographolide were later isolated from the A. paniculata leaves, but until this day, the bioactivity of these compounds is still unknown (Bright et al., 2001). C. nutans contains terpenoids, phenolics and other potentially bioactive compounds, including sulphur-containing glucosides, lipids, chlorophyll derivatives and benzonoids (Table 1.4). However, to date, no alkaloids have been discovered in the C. nutans.

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Table 1.4. Phytochemical compounds present in different parts of C.nutans. Compound class Compound Plant part(s) Extract(s) Reference(s) Steroids Stigmasterol Stem Petroleum Dampawan (1976) β-sitosterol Leaf Dampawan et al. (1977) Root Lin et al. (1983) Triterpenoids Lupeol Stem Petroleum Dampawan et al. (1977) Betulin Root Lin et al. (1983)

Phenolic acids Caffeic acid Leaf Methanol Ghasemzadeh et al. (2014) Gallic acid Leaf buds Flavonoids Catechin Leaf Methanol Ghasemzadeh et al. (2014) Kaempferol Leaf buds Luteolin Quercetin C-glycosyl flavones Vitexin Stem Butanol and water soluble Teshima et al. (1997) Isovitexin Leaf portions of methanol Chelyn et al. (2014) Shaftoside Ethanol Mustapa et al. (2015) Isomollupentin 7-O-ϐ-glucopyranoside Orientin Isoorientin Sulphur-containing Clinacoside A Stem Butanol and water soluble Teshima et al. (1998) glucosides Clinacoside B Leaf portions of methanol Tu et al. (2014) Clinacoside C Ethanol Cycloclinacoside A1 Cycloclinacoside A2 Clinamide A Clinamide B Clinamide C 2-cis-entadamide A Glycerides Digalactosyl diglycerides Leaf Unknown Janwitayanuchit et al. Trigalactosyl diglycerides (2003)

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Table 1.4. Phytochemical compounds present in different parts of C.nutans (continued). Compound class Compounds Plant part Extract(s) Reference(s) Cerebrosides 1-O-ϐ-D-glucosides of phytosphingosines Leaf Ethyl acetate soluble Tuntiwachwuttikul et al. Monoacylmonogalactosyl- (2S)-1-O-linolenoyl-3-O-ϐ-D- fraction of ethanol (2004) glycerol galactopyranosylglycerol Chlorophyll derivatives 132-hydroxy-(132-R)-phaeophytin a Leaf Chloroform Shuyprom (2004) (phaeophytins) – related to 132-hydroxy-(132-R)-phaeophytin b Hexane Sakdarat et al. (2008) chlorophyll a and b 132-hydroxy-(132-S)-phaeophytin a Sakdarat et al. (2009) 132-hydroxy-(132-S)-phaeophytin b Sittiso et al. (2010) 132-hydroxy-(132-R)-chlorophyll b 132-hydroxy-(132-S)-chlorophyll b purpurin 18 phytyl ester phaeophorbide a Benzenoids 1,2-benzenedicarboxylic acid, mono(2- Leaf Chloroform Yong et al. (2013) ethylhexyl) ester

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1.3.6.1 Influence of genetics

The quality and quantity of phenolic compounds are affected by genetics (Manach et al.,

2004). The pattern and intensity of the biosynthesis of these compounds are regulated by the expression of genes that are involved, such as the F3’H and ANR genes that regulate flavonoid synthesis (Gerats and Martin, 1992). In some plant species, genetics is a major determinant of phenolic concentration rather than environmental factors, such as climate and sunlight (Goldberg et al., 1998, Bruni and Sacchetti, 2009, Schlag and McIntosh, 2013).

Conversely, this is not the case for other plant species, where the variation in phenolic compounds is mainly controlled or regulated by environmental factors (Grass et al., 2006,

Šamec et al., 2015). However, the combination of both genetic and environmental factors may synergistically influence phenolic synthesis and accumulation in plants, such as that seen on Fragaria × ananassa (Rosaceae) (Carbone et al., 2009). Currently, no studies have investigated the influence of genetics on the phenolic content in C. nutans. This is particularly important in the identification and selection of superior varieties for optimising pharmacological properties and the medicinal value of the plant for end consumer products.

1.3.6.2 Influence of environmental factors

The habitat or cultivation site of medicinal plants is important in the production of phenolic compounds. This is because the content of these compounds in the plants varies with changing environmental conditions (Fonseca et al., 2006), resulting in differences in their medicinal values. Environmental factors, including solar radiation, temperature, rainfall

(water availability), soil pH and salinity can affect the quality and quantity of phenolic compounds (Ewane et al., 2012). Certain compounds are only synthesised or significantly increased under specific environments (Liu et al., 2015). Medicinal plants of the same species grown in different environments (e.g. altitude, air temperature and sunlight) have been shown

CHAPTER 1 to differ in their phenolic contents. For instance, plants grown at higher altitudes, which is often associated with an increased intensity of ultraviolet radiation (Blumthaler et al., 1997) and lower air temperatures (Whiteman, 2000), had higher amounts of phenolics compounds

(Kishore et al., 2010, Dong et al., 2011, Guo et al., 2011, Bernal et al., 2013). However, higher altitudes do not always result in higher levels of phenolic compounds (Umek et al.,

1999, Bahukhandi et al., 2014, Yang et al., 2011), such as that seen in Justicia pectoralis

(Acanthaceae) (Méndez Arteaga et al., 2011), where no significant correlation was observed between the two parameters. The disparity in results suggests that the influence of altitude variability on the phenolic content is different for each plant and species or it could be due to other factors, such as soil nutrients. Currently, the influence of altitude and air temperature on

C. nutans is unknown. Given that the phenolic content in medicinal plants is significantly impacted by the environment, it is crucial to investigate how and to what extent environmental factors affect the bioactive compounds in C. nutans, which may determine its efficacy. The effects of ultraviolet radiation and air temperature on the phytochemical content will be further discussed.

1.3.6.2.1 Ultraviolet radiation

Ultraviolet (UV) light is an electromagnetic radiation, which comprises of just over 8% of the total solar radiation (Hollosy, 2002). UV is mainly divided into three types of rays based on their wavelength ranges:

i) UVA (320-400 nm), which represents ~ 6% of the incoming solar radiation and is

the least harmful among the other UV rays (Hollosy, 2002);

ii) UVB (280-320 nm), although this represents only ~ 1.5% of the total solar

spectrum, it can induce various damaging effects on plants and hence, this ray

gains the most attention (Hollosy, 2002); and

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iii) UVC (200-280 nm), which is the most hazardous to living organisms, but is not

relevant under natural conditions because it is blocked by the ozone layer (Hollosy,

2002).

UV radiation can affect the morphological, physiological and biochemical processes in plants

(Hollosy, 2002). Morphological alterations such as increased leaf thickness, decreased leaf area and stomatal frequencies are mechanisms of plants to cope with UV exposure (Barnes et al., 1990) to reduce the penetration of the radiation (Hollosy, 2002). The epidermis plays a key role as a protective barrier against harmful effects of UV radiation, since it is the first cell layer to intercept radiation (Hollosy, 2002). The epidermis often comprises of UV-absorbing compounds, such as phenolics, particularly flavonoids, to prevent further penetration of the radiation into inner cell layers (Carbonell-Bejerano et al., 2014). Flavonoids have absorption wavelengths in the range of 240-545 nm (Zhang et al., 2011), which fall in the absorbance of

UVA and UVB wavelength bands. Therefore, these compounds provide protection for the inner leaf cells, acting as sunscreens in the leaf epidermis against UV radiation, while transmitting photosynthetically active radiation (PAR) to the mesophyll cells (Morales et al.,

2010). Plants exposed to increased UV radiation produce higher flavonoid content (Ibdah et al., 2002, Ryan et al., 2002, Garcia-Macias et al., 2007, Zhang et al., 2011) through the increase in activity of enzymes, including phenylalanine ammonia lyase (PAL) and chalcone synthase (CHS), involved in the flavonoid biosynthesis (Hollosy, 2002).

1.3.6.2.2 Air temperature

Low air temperatures cause oxidative stress in plants, resulting in increased production of reactive oxygen species (ROS) (Aroca et al., 2001). ROS reacts with plant lipids, proteins and

DNA, causing cellular damage (Shohael et al., 2006). Therefore, plants develop protective

CHAPTER 1 mechanisms to combat the damaging effect by producing antioxidant substances, such as phenolic compounds, including flavonoids (Brewer, 2011). Low temperatures affect the biochemical processes in plants through changes in the expression of genes involved in the biosynthesis of phenolic compounds, such as PAL and CHS genes (Lattanzio et al., 2012).

Low temperatures enhance PAL and CHS enzyme activities, leading to increased in phenolic compounds and their easter- and glycoside-bound forms in plants (Dufoo-Hurtado et al.,

2013). Low temperature treatments stimulated phenolic metabolism, resulting in significantly higher phenolic content in Siberian ginseng (~ 10-fold, 24 vs. 30°C) (Shohael et al., 2006) and comparatively higher in olive leaves (-20 vs. 4°C) (Cansev et al., 2012) and in sweetpotato leaves (20 vs. 30°C) (Islam et al., 2003).

1.3.6.3 Influence of plant age and part

The quality and quantity of bioactive phenolic compounds are dependent on the plant age

(Inderjit, 1996). Young and developing leaves contain an abundance of photosynthases and serve as a major site for phytohormone production, where they are nutrient-rich and tender in physical structure compared to mature leaves (Liu et al., 1998). Consequently, they are susceptible to herbivory and pathogenic attacks. Most plants defend their juvenile developmental stages by producing higher phytochemical concentration as a chemical defense mechanism (Liu et al., 1998). In constrast, de novo biosynthesis of phytochemicals typically proceeds at lower rate in mature leaves, resulting in decrased levels of phytochemical content (Zagrobelny et al., 2008). This was observed in C. nutans, where the

CHS activity, hence, the total phenolic and flavonoid content was comparatively higher in 6- month old leaves than 12-month old leaves (Ghasemzadeh et al., 2014). Similar patterns were also observed in other species, such as Ilex paraguariensis (Aquifoliaceae) (Blum-Silva et al.,

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2015), Olea europaea (Oleaceae) (Achakzai et al., 2009) and A. paniculata (Chua et al.,

2013).

The concentration of phytochemical constituents varies with different plant parts. Parts that are exposed to environmental stimuli, such as the presence of light, generally have greater amounts of phenolic compounds (Achakzai et al., 2009). For example, the leaves of Larrea tridentate (Zygophyllaceae) contained higher total phenolic content with 36.2 mg/100 g than the roots with 28.6 mg/100 g (Hyder et al., 2002). Comparatively higher total phenolic content was also observed in the leaves of O. europaea than in the stems (Achakzai et al.,

2009). However, this effect is still unknown on C. nutans.

1.3.6.4 Influence of postharvest processes

Postharvest is defined as the transition of a product from harvest to its ready-to-use state.

Postharvest processes of medicinal plants generally focus on optimising drying, storage and extraction to retain bioactive compounds. These processes have been described in several reviews (Tanko et al., 2005, Naczk and Shahidi, 2006, Rocha et al., 2011), including drying

(Section 1.3.6.4.1) and extraction (Section 1.3.6.4.2) treatments.

1.3.6.4.1 Drying

Drying is necessary to prolong the storage shelf life of medicinal plants by reducing the moisture content to prevent deterioration and infestation (Müller, 2007). Convective drying is a popular method to dry plants because it offers a faster drying rate and controllable drying temperatures (Yi and Wetzstein, 2011, Lusia Barek et al., 2015). The quality of phytochemical constituents and their concentrations in plant materials are influenced by drying temperatures (Chen et al., 2011, Yi and Wetzstein, 2011, Srinivasan and

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Sivasubramaniam, 2014, Periche et al., 2015). Hence, the selection of proper drying temperature is crucial for the retainment of bioactive compounds in plant materials. This is to ensure that the desired chemical and medicinal properties of target compounds are achieved by maintaining their structure during different stages of drying (Volf et al., 2014). Given the different physiochemical properties of phenolic compounds, they can vary in their thermal stability. Phenolic compounds with a higher number of hydroxyl groups are more susceptible to thermal degradation (Buchner et al., 2006). For instance, rosmarinic acid and eupatorin from Orthosiphon stamineus leaves showed higher degradation (54.2 and 39.8%, respectively) due to the presence of hydroxyl groups compared to sinensetin (21.7%), which has no hydroxyl group on its molecular structure (Pang et al., 2014). This was also observed with catechin in vegetal extracts of grape seeds and spruce barks, which showed 5% higher degradation than gallic acid when exposed to 100°C for 250 min (Volf et al., 2014). However, the effect of drying temperatures on the phenolic compounds of C. nutans remains unknown.

1.3.6.4.2 Extraction

The aromatic ring of a phenolic compound is nonpolar, whereas the hydroxyl group attached to the ring is polar in nature (Galanakis et al., 2013). Therefore, the polarity of phenolic compounds varies significantly, depending on the number of aromatic rings and hydroxyl groups they possess (Naczk and Shahidi, 2006). Besides, phenolic compounds may have multiple hydroxyl groups that can be conjugated to sugars, acids or alkyl groups, altering their polarities (Garcia-Salas et al., 2010). Less polar phenolic compounds (e.g. chlorinated phenols, flavonols, isoflavones, flavanones and methylated flavones) are extracted with hexane, chloroform, dichloromethane, diethyl ether or ethyl acetate, while more polar phenolic compounds (e.g. ferullic acids, vanillic acids, flavonoid glycosides and aglycones) are extracted with alcohols, such as methanol and ethanol (Marston and Hostettmann, 2006).

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Some phenolic compounds (e.g. benzoic, cinnamic acids) cannot be completely extracted with pure organic solvents or water alone because these compounds distribute between mutually saturated solvent phases (Marcus, 1990). Hence, alcohol-aqueous mixtures are recommended (Stalikas, 2007). The presence of water in these mixtures increases the process of solvation (Boeing et al., 2014), while organic solvents, such as methanol and ethanol, act as hydrogen bond donors and acceptors (Nedic et al., 2011). Consequently, this synergistic effect increases the solubility of compounds, resulting in better extraction capacity. For instance, extraction using 50% (v/v) ethanol yielded higher total phenolic (~ 1.6-fold) and flavonoid (~ 2-fold) contents from leaves and stems of C. nutans than with 86% (v/v) ethanol

(Mustapa et al., 2015), suggesting the former ethanol concentration is the best solvent combination to extract polyphenols from the plant. Extraction of phenolic compounds from A. paniculata leaves with 60% (v/v) ethanol also showed ~ 2.5-fold higher level of total phenolic content than using 100% (v/v) ethanol (Thoo et al., 2013).

Solvent temperature also affects the amount of phenolic compounds being extracted from plants (Naczk and Shahidi, 2006, Stalikas, 2007), but this effect is unknown on C. nutans phenolics. An increase in solvent temperatures can promote higher analyte solubility by increasing mass transfer rate through interruption of the physical chemistry interactions in the plant material (Ong et al., 2006). Additionally, the viscosity and the surface tension of the solvents are decreased at higher temperatures, allowing the solvents to penetrate the plant matrix (Dai and Mumper, 2010). For instance, extraction of A. paniculata leaves with ethanol at 65°C resulted in ~ 14% higher in total phenolic content than 25°C (Thoo et al., 2013). A similar trend was also observed with leaves of Camellia sinensis (Theaceae), where extraction with aqueous at 90°C had 25% higher amount of phenolic compounds than those extracted at ambient temperature (25°C) (Venditti et al., 2010). However, too high a

CHAPTER 1 temperature can increase the chance of compound degradation by hydrolysis, internal redox or polymerization, resulting in decresed yield of phenolic compounds (Alonso-Salces et al.,

2001). For example, anthocyanins are best extracted at temperatures ranging from 20 to 50°C because temperatures over 70°C cause rapid degradation of these compounds (Dai and

Mumper, 2010).

1.3.6.5 Analysis and quantification

Selection of suitable analytical and quantification methods for studying bioactive phenolic compounds in plant materials depends on the compounds or components of interest (Naczk and Shahidi, 2006). Several methods have been used to qualify and quantify phenolic compounds from plant extracts (Naczk and Shahidi, 2006, Stalikas, 2007, Khoddami et al.,

2013). Chemical procedures, such as Shinoda’s, ferric chloride, lead ethanoate and sodium hydroxide tests are used to detect the presence of phenolic compounds qualitatively, while spectrophotometric and chromatographic techniques are used to separate, quantify and identify them (Khoddami et al., 2013).

1.3.6.5.1 Spectrophotometry

A number of UV spectrophotometric methods have been developed for the quantification of plant phenolics. Reactions forming coloured complexes are commonly used due to its affordability, simplicity, fast measurement and applicability in most laboratories (Pelozo et al., 2008). The Folin-Ciocalteu is the most widely used method for the quantification of total phenolic content in plant materials (Khoddami et al., 2013). The method is based on the oxidation of phenolic compounds in plant extracts by the Folin-Ciocalteu reagent, containing a mixture of phosphotungstic acid and phosphomolybdic acid. The redox reaction results in the formation of blue chromophores consisting of phosphotungstic-phosphomolybdenum

CHAPTER 1 complexes, which has a maximum absorption of 765 nm, and is proportional to the total quantity of phenolic compounds (Ainsworth and Gillespie, 2007). The total phenolic content of C. nutans determined using the Folin-Ciocalteu method ranged from 0.78 to 20 mg gallic acid equivalent/g dry extract (Yuann et al., 2012, Ghasemzadeh et al., 2014, Tiew et al., 2014,

Wong et al., 2014, Lusia Barek et al., 2015, Mustapa et al., 2015).

The determination of total flavonoid content is based on the detection of coloured complexes formed from the reaction between aluminium chloride (AlCl3) and the carbonyl and hydroxyl groups of flavonoids in an alkaline solution (Magalhães et al., 2012). The total flavonoid content of C. nutans ranged from 0.21 to 6.32 mg quercetin equivalent/g dry extract

(Ghasemzadeh et al., 2014, Tiew et al., 2014, Wong et al., 2014, Mustapa et al., 2015).

Different flavonoid classes react differently with AlCl3, resulting in dissimilar absorption maxima, which leads to variations in experimental conditions and universal parameters

(Mammen and Daniel, 2012). Therefore, several compromises and resolutions have been recommended based on the different flavonoid classes of interest. The AlCl3 method was initially proposed by Christ and Mueller (1960) for the analysis of O-glycoside flavonoids but unsuitable for C-glycosides because this class of flavonoid involves a different chemical reaction (Fernandes et al., 2012). Subsequently, Woisky and Salatino (1998) described the use of AlCl3 to develop deep yellow-coloured complexes of the flavonoids detected at 420 nm, using quercetin as the standard but this method only estimated flavones and flavonols.

Further modifications, including the post-addition of potassium acetate and measurement of absorbance at 415 nm produced better detection and worked well for flavonols kaempferol, quercetin and myricetin (Mammen and Daniel, 2012).

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1.3.6.5.2 Chromatographic

High performance liquid chromatography (HPLC) is one the most frequently used chromatographic techniques for the separation, identification and quantification of phenolic compounds in plants, including C. nutans (Table 1.5). HPLC can detect and separate all analysed compounds simultaneously, regardless of the chemical nature of phenolics and flavonoids, including hydrophobic aglycones to hydrophilic flavonoid glycosides, derivatives and degraded products (Stalikas, 2007). Moreover, it can detect low concentrations of compounds ranging from nanogram (ng) to picogram (pg) (Dong, 2006) in the presence of many interfering and co-eluting components (Stalikas, 2007). The main disadvantages are its high initial capital cost, complex procedure that is time consuming and results are affected by various factors, including sample purification, mobile phase, column types and detectors

(Khoddami et al., 2013).

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Table 1.5. HPLC parameters used for the detection and quantification of compounds in C. nutans. Stationary phase Column description Mobile phase Elution Detector and Compounds Reference (l × dc, dp) wavelength Pentafluoro- 250 × 4.6 mm, 5 µm Water with 0.8% (v/v) Gradient UV-Vis/DAD Flavone C-glycosides Chelyn et al. (2014) phenyl glacial acetic acid and 330 nm acetonitrile

C18 250 × 4.6 mm, 5 µm Ortho-phosphoric acid Gradient UV-Vis Phenolic acids and Ghasemzadeh et al. and methanol 260-360 nm Flavonoids (2014)

C18 150 × 4.6 mm, 5µm Methanol and Isocratic ELSD β-sitosterols Mustapa et al. (2015) 2-propanol 210 nm

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The separation of phenolic compounds is usually performed using a reversed-phase system, which involves the adsorption of hydrophobic (nonpolar) compounds onto a hydrophobic stationary phase in a polar mobile phase, resulting in more polar compounds eluting first

(Dobes et al., 2013). A typical stationary phase is made up of silica-based particles, where the surface is attached with hydrophobic alkyl chains that interact with the analyte. The three common carbon chain lengths are C4, C8 and C18. The most popular column for the separation of different classes of compounds is the reversed-phase octadecyl carbon chain (C18)

(Proestos et al., 2006, Chua, 2007, Mitra, 2014, Uddin et al., 2014). This is because C18 columns offer retention and selectivity for a wider range of compounds containing different polar and nonpolar groups on their surface (Sagar et al., 2012), whilst C4 and C8 columns are preferentially used to separate proteins (Sagar et al., 2012). Most reversed-phase C18 columns have lengths ranging from 100-250 mm, with an internal diameter between 3.9 and 4.6 mm and particle sizes between 3-10 µm (Stalikas, 2007). A pentafluorophenyl stationary is preferred for the separation of compounds with closely related structures through the unique interactions between the fluorine groups of the stationary phase and the compounds (Joseph et al., 2010). For example, pentafluorophenyl column was used to separate vitexin and isovitexin extracted from C. nutans leaves (Chelyn et al., 2014).

Acetonitrile and methanol, or their aqueous forms, are the dominant mobile phases used in

HPLC quantification of phenolic compounds (Vagiri et al., 2012, Ni et al., 2013, Zhang et al.,

2013, Mehta et al., 2014). Maintaining the pH of the mobile phase in the range of 2-4 is recommended to prevent ionisation of phenolic compounds during analysis. Therefore, aqueous acidified mobile phases usually contain acids, such as acetic, formic and phosphoric, or buffers like phosphate, citrate and ammonium acetate (Lee et al., 2008, Diagone et al.,

2012, Porel et al., 2014, Aranda-Gonzalez et al., 2015). Additionally, a gradient elution

CHAPTER 1 system is preferred (Khoddami et al., 2013) over isocratic elution (constant mobile phase composition throughout the chromatographic run), when plant extracts contain phenolic compounds with variable polarity (Rounds and Gregory). The gradual changes of the mobile phase composition during the chromatographic run allow a faster separation without a loss of resolution in the earlier peaks or unnecessary broadening of later peaks.

The HPLC system consists of a detector, which measures the concentration of eluting compounds based on one of their inherent properties, such as UV absorbance (Dong, 2006).

Most plant phenolics can be detected at UV-vis wavelengths of 210-370 nm (Brandsteterova and Ziakova-Caniova, 2005), with flavonoids at 240-545 nm (Zhang et al., 2011). The UV-

Vis detector has reasonable sensitivity, wide linear dynamic ranges, stability and compatibility with gradient solvent system (Burgard and Kuznicki, 1990). However, the use of photodiodide array detectors (DAD) is gaining popularity for the detection of phenolic compounds in plant extracts (Luo et al., 2013, Ni et al., 2013, Riethmuller et al., 2013, Chen et al., 2014a). This is because it allows the selection of wavelength range (variable wavelength) for each peak during an analysis, which UV-Vis cannot (single wavelength)

(Burgard and Kuznicki, 1990).

1.3.7 Traditional medicinal uses of C. nutans

C. nutans is a popular medicinal plant endemic to Asia and recognised as one of Thailand’s

National List of Essential Medicines (2011) by the National Drug Committee (Pratoomsoot et al., 2015). This implies that C. nutans satisfies the priority health care needs of the Thai population of Thailand with public health relevance, evidence of efficacy and safety as well as comparative cost-effectiveness (WHO, 2015).

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In Thailand, C. nutans leaves have been used for the treatment of skin rashes, burns, mucositis and lesions of herpes simplex virus (HSV) and varicella-zoster virus (VZV)

(Putwatana et al., 2009, Sakdarat et al., 2009). The leaves are also used as a remedy against venomous snake bites, scorpion and insect stings in Malaysia and Thailand (Uawonggul et al.,

2006, Globinmed, 2015), resulting in the vernacular name, Sabah snake grass. Leaf extracts of C. nutans are widely used on cancer patients throughout Malaysia (Shim et al., 2013). In

Indonesia, the leaves are used to treat diabetes and dysentery (Tiew et al., 2014, Lusia Barek et al., 2015).

1.3.8 Pharmacological activities of C. nutans

C. nutans possesses a range of pharmacological activities (Table 1.6). Clinical trials have been carried out to investigate the efficacy of C. nutans extract in treating herpes (HSV and

VZV) patients (Jayavasu et al., 1992, Sangkitporn et al., 1993, Sangkitporn et al., 1995,

Charuwichitaratana et al., 1996). Findings from these studies revealed faster recovery

(crusting time within 3 d and healing time within 7 d) in patients treated with the extract compared to the placebo group (crusting time 4-7 d and healing time 7-14 d). Additionally, the extract exhibited similar efficacy as the antiviral drug, Acyclovir, which is chiefly prescribed to treat herpes infections. Besides, patients receiving the extract as treatment described minimal pain and no side effects.

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Table 1.6. Pharmacological activities of C. nutans. Activity Plant part Extract Effective dose Effect Reference Anticancer Leaf Chloroform 47.70 µg/mL 50% growth inhibition of human Yong et al. (2013) erythroleukemia (K-562) cells at 72 h Leaf Chloroform 47.31 µg/mL 50% growth inhibition of Burkitt’s Yong et al. (2013) lymphoma (Raji) cells at 72 h Leaf Petroleum ether 18.00 µg/mL 50% growth inhibition of cervical Arullappan et al. carcinoma (HeLa) cells at 72 h (2014) Leaf Petroleum ether 20.00 µg/mL 50% growth inhibition of K-562 cells Arullappan et al. at 72 h (2014) Antidiabetic Whole Aqueous 50 mg/mL 88% α-glucosidase inhibition Wong et al. (2014) Antiinflammatory Whole Methanol 9 mg/ear 79% ethyl phenylpropriolate-induced Wanikiat et al. (2008) ear oedema inhibition in rats at 15 min Whole Methanol 9 mg/ear 44% myeloperoxidase inhibition Wanikiat et al. (2008) Whole Methanol 200 mg/kg 59% carrageenan-induced hindpaw Wanikiat et al. (2008) oedema inhibition in rats at 3 h Aerial Ethanol 0.01 mg/mL 68% elastase release inhibition in Tu et al. (2014) human neutrophil cells Antimicrobial Leaf Ethyl acetate 6.31 mg/mL Inhibition of Bacillus cereus growth Arullappan et al. (2014) Leaf Ethyl acetate 6.31 mg/mL Inhibition of Candida albicans growth Arullappan et al. (2014) Antioxidant Leaf Chloroform 7.85 mg Teq/g extract Against DPPH radicals Yong et al. (2013) Leaf Chloroform 12.24 mg Teq/g extract Against galvinoxyl radicals Yong et al. (2013) Leaf Methanol 0.10 mg/mL 34% hydrogen peroxide inhibition Yong et al. (2013) Leaf Aqueous 0.10 mg/mL 32% nitric oxide scavenging activity Yong et al. (2013) Leaf Water-soluble 0.11 mg/mL 50% DPPH scavenging activity Pannangpetch et al. portion of ethanol (2007) Leaf Water-soluble 0.36 mg/mL 50% haemolysis inhibition in rats Pannangpetch et al. portion of ethanol (2007) Leaf Methanol 1.33 mg/mL 50% DPPH scavenging activity Tiew et al. (2014)

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Table 1.6. Pharmacological activities of C. nutans (continued). Activity Plant part Extract Effective dose Effect Reference Antioxidant Leaf Petroleum ether 4.00 mg/mL 82% DPPH scavenging activity Arullappan et al. (2014) Stem Methanol 10.00 mg/mL 70% DPPH scavenging activity Arullappan et al. (2014) Whole Aqueous 16.00 mg/mL 60% DPPH scavenging activity Wong et al. (2014) Whole Aqueous 10.00 mg/mL 95% nitric oxide scavenging activity Wong et al. (2014) Whole Aqueous 10.00 mg/mL 90% metal chelating activity Wong et al. (2014) Antiviral Aerial Ethanol 31. 04 µg/mL 50% anti-dengue inhibition at 72 h Tu et al. (2014) Leaf Ethy acetate 19.00 µg/mL 100% inhibition of HSV-type 1F plaque Thongchai et al. (2008) formation Mode of action: pre-infection stage Leaf Phaeophytins* 5.00 µg/mL 100% inhibition of HSV-type 1F plaque Sakdarat et al. (2009) formation Mode of action: pre-infection stage Leaf CL03* 200.00 µg/mL 87% DNA intensity reduction of HSV- Vachirayonstien et al. type 2 (2010) Leaf CL21* 80.00 µg/mL 92% DNA intensity reduction of HSV- Vachirayonstien et al. type 2 (2010) Leaf Hexane 32.05 µg/mL 50% HSV-type 1 plaque reduction Kunsorn et al. (2013) Leaf Dichloromethane 44.50 µg/mL 50% HSV-type 1 plaque reduction Kunsorn et al. (2013) Leaf Methanol 64.93 µg/mL 50% HSV-type 1 plaque reduction Kunsorn et al. (2013) Leaf Hexane 72.62 µg/mL 50% HSV-type 2 plaque reduction Kunsorn et al. (2013) Leaf Dichloromethane 65.19 µg/mL 50% HSV-type 2 plaque reduction Kunsorn et al. (2013) Leaf Methanol 65.13 µg/mL 50% HSV-type 2 plaque reduction Kunsorn et al. (2013) Leaf Unknown 1:9600 weight per 50% VZV plaque reduction Thawaranantha et al. volume dilution ratio (1992) Leaf 136B* 3.70 µg/mL 74% inhibition of human Sookmai et al. (2011) papillomavirus (HPV) infection Leaf 136C* 7.60 µg/mL 85% inhibition of HPV infection Sookmai et al. (2011) Leaf 136D* 19.40 µg/mL 96% inhibition of HPV infection Sookmai et al. (2011) Note: * pure compounds

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Table 1.6. Pharmacological activities of C. nutans (continued). Activity Plant part Extract Effective dose Effect Reference Cholinergic Leaf Methanol 0.25 g/kg 1.2-fold increased in acetylcholinesterase Lau et al. (2014) neurotransmission activity in mouse brain compared to modulatory distilled water-treated control after 14 d Leaf Methanol 0.25 g/kg 2.5-fold increased in acetylcholinesterase Lau et al. (2014) activity in mouse heart compared to distilled water-treated control after 14 d Leaf Methanol 0.25 g/kg 1.2-fold increased in acetylcholinesterase Lau et al. (2014) activity in mouse kidney compared to distilled water-treated control after 14 d Leaf Methanol 0.25 g/kg 1.6-fold increased in acetylcholinesterase Lau et al. (2014) activity in mouse liver compared to distilled water-treated control after 14 d DNA-protective Leaf Ethanol 10.00 mg/mL Reduced plasmid DNA cleavages and Yuann et al. (2012) retained DNA integrity after riboflavin exposure Immune- Aerial Ethanol 0.10 µg/mL 78% up-regulated interferon gamma Tu et al. (2014) stimulating (IFN-ɣ) compared to media control Aerial Ethanol 100.00 µg/mL 70% down-regulated interferon gamma Tu et al. (2014) (IFN-ɣ) compared to media control Leaf Water-soluble 1.00 mg/mL 60% reduction in natural killer (NK) cell Sriwanthana et al. (1996) portion of ethanol activity compared to non-treated control

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1.3.9 Cancer

The cell cycle (Figure 1.6) is divided into two basic parts: mitosis and interphase, where the latter consists of G1, G2 and S phases (Cooper, 2000). The mitosis (M) phase corresponds to the separation of daughter chromosomes (nuclear division) and ends with cell division. This phase is followed by the G1 phase, which is the interval between M phase and initiation of

DNA replication. During G1, cells are metabolically active and continue to grow but do not replicate its DNA. G1 is followed by synthesis (S) phase, during which DNA replication occurs. The completion of DNA synthesis is followed by the G2 phase, during which cell growth continues and proteins are synthesized in preparation for mitosis (Cooper, 2000). If extracellular conditions are unfavourable, cells enter a specialised resting phase (G0), in which they remain for days, weeks or even years before resuming proliferation (Alberts et al.,

2002).

Figure 1.6. The phases of the cell cycle. Checkpoints (red bars) are control mechanisms to ensure proper cell growth and division. Oncogenes (A) allow for checkpoints to be overcome (stop lines) and tumour suppresors (B) act to maintain checkpoints (arrows). Adapted from Calvert and Frucht (2002) and Chow (2010).

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In normal cells, growth is controlled in a balanced manner between growth-promoting and growth-inhibiting signals such that proliferation occurs only when required (Macdonald et al.,

2004). On the contrary, cancer is characterised by unregulated cell growth and inappropriate proliferation (Pecorino, 2008). Proto-oncogenes code for normal proteins that mediate the growth-promoting signal in a controlled manner (Chow, 2010). However, once they are altered to become oncogenes through chromosomal translocation, gene mutation or amplification (Croce, 2008), they code for abnormal proteins that exhibit increased activity, which leads to tumour formation (Chow, 2010). Instead of stopping within a G phase as it normally should, oncogenes interfere with the cell cycle checkpoints by overcoming them and progressing through subsequent phases, resulting in uncontrolled cell division (Figure

1.6A) (Chow, 2010). Tumour suppressor genes on the other hand, are normal genes, which code for proteins that maintain cell cycle checkpoints to ensure proper cell growth and division (Figure 1.6B) (Chow, 2010). These proteins mediate the growth-inhibiting signal that restricts unnecessary cell growth and division or activates programmed cell death

(apoptosis) as well as DNA repair (Nicot, 2015). However, mutated tumour suppressor genes lead to tumourigenesis by preventing regulatory proteins from inhibiting cell proliferation

(Chow, 2010) and promoting cell death (Croce, 2008). Examples of oncogenes, tumour suppressor genes and the cancer types that have been identified ranging from localised to systemic cancers (Table 1.7).

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Table 1.7. Oncogenes and tumour suppressor genes associated with cancers (Weinstein and Joe, 2006, Pecorino, 2008). Gene category Gene Cancer associated with mutation Oncogene c-Myc T-cell and acute myeloid leukaemia, pancreas β-cell, osteogenic sarcoma, breast BCR/ABL Leukaemia HRAS Melanoma, breast KRAS Melanoma, pancreas ERBB2 Breast WNT1 Breast CCND1 Oesophagus, colon, pancreas, squamous, nasopharynx, breast CTNNB1 Colon CCNE Liver BRAF Melanoma, kidney MITF Melanoma c-Kit Gastrointestinal EFGR Head and neck, colorectal, pancreas, non-small-cell lung VEGF Breast, colorectal, kidney Tumour suppressor gene RB1 Retina, bone, breast WT1 Kidney p53 Sarcoma, breast, brain APC Colon CDKN2A Melanoma, pancreas BRCA 1 & 2 Breast, ovary FHIT Lung, stomach, kidney, cervix PTEN Brain, prostate, breast TSC2 Renal, brain CDH1 Breast, colon, skin, lung MSH2 & 6 Colorectal PMS1 & 2 Colorectal

Cancer is considered as a major cause of mortality worldwide. The International Agency for

Research on Cancer (IARC) estimates that approximately 14 million new cancer cases are diagnosed each year worldwide and the number is predicted to increase by about 70% over the next two decades (Forman and Ferlay, 2014).

Cancers are characterised based on the tissue of origin. Cancers from epithelial cells (e.g. skin) are classified as carcinomas, while those derived from mesoderm cells (e.g. bone,

CHAPTER 1 muscle) and glandular tissue (e.g. breast) are sarcomas and adenocarcinomas, respectively.

Cancer tumours can be benign or malignant, where the former is not cancerous and it does not spread (metastasize) to other parts of the body. Malignant tumours do not remain confined, they invade and metastasize to other organs (Pecorino, 2008).

1.3.9.1 Breast cancer

Breast cancer is the most commonly diagnosed cancer and cause of death among women

(Schnitt and Lakhani, 2014). In 2012, there were an estimated 1.7 million new cases (25% of all cancers in women) and 0.5 million cancer deaths (15% of all cancer deaths in women) worldwide (Schnitt and Lakhani, 2014). Breast cancers have been studied more than any other cancer types with regard to oncogene expression (Macdonald et al., 2004) but there are still knowledge gaps on the molecular mechanisms leading to breast cancer development.

Over 60% of cases were found to express MYC, ERBB2 or one of the RAS family genes

(Macdonald et al., 2004). Meanwhile, some of the mutated tumour suppressor genes that are frequently found in breast cancers are BRCA1, BRCA2, TP53 and RB1 (Macdonald et al.,

2004). The two human breast carcinoma cell lines used in this study were MCF7 and BT474.

MCF7 cells have the functional wild-type p53 gene, whereas BT474 cells carry a temperature-sensitive mutation in the p53 gene (E285K), which has been suggested to adopt a mutant conformation at 37°C but has a wild-type conformation at 32°C (Müller et al., 2005).

In addition to mutations, a second mechanism is believed to inactivate the wild-type p53 gene.

In some types of breast cancer, there is a possibility of the wild-type p53 protein losing its tumour-suppressive functions by being sequestered in the cytoplasm and prevented from entering the nucleus, its normal site of action (Moll et al., 1992). This may be an explanation to why some breast cancers do not show any p53 gene mutations (Macdonald et al., 2004).

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1.3.9.2 Melanoma

Melanoma is a type of skin cancer, characterised as a neoplastic disorder of the pigment- producing cells in the skin known as melanocytes (Uong and Zon, 2010). Melanoma is the least common but most fatal form of skin cancer and it has increased two-fold in the past 20 years (Cifola et al., 2013). Globally, there were more than 232,000 new cases of melanoma and about 55, 000 deaths estimated in 2012 (Guo and Mihm Jr., 2014). In Australia, it is the fourth most commonly diagnosed cancer, with more than 11,000 cases reported in 2010

(Australian Institute of Health and Welfare, 2014). Although the development of melanoma occurs via different signalling pathways, approximately 50% of melanomas bear mutations in the BRAF gene (Johnson and Sosman, 2014). Among the BRAF mutations, over 90% are at codon 600, and among these, more than 90% are a single nucleotide mutation resulting in the substitution of glutamic acid for valine (BRAF V600E) (Ascierto et al., 2012). BRAF is a serine/threonine kinase that functions downstream of RAS protein in the mitogen-activated protein (MAP) kinase signalling pathway (Lee and Fakharzadeh, 2008). Mutated BRAF gene causes a constitutional activation of the methyl ethyl ketone (MEK)/ extracellular signal- regulated kinase (ERK) effectors, which are involved in the MAP kinase pathway, leading to enhanced cellular proliferation and neoplastic transformation (Laurini et al., 2012). The two melanoma cell lines used in this study were D24 and MM418C1. The former has the wild- type BRAF gene, whereas the latter harbours the BRAF V600E mutation.

1.3.9.3 Classification of cell death

Generally, cell death can be classified into two major forms: apoptosis, which is programmed cell death, and necrosis, defined as non-programmed or uncontrolled cell death (Messner et al., 2012). In the last decade, programmed cell death has expanded to include autophagy and necroptosis (programmed necrosis) (Su et al., 2015). Autophagy is a catabolic process, where

CHAPTER 1 a double-membraned vesicle is formed to encapsulate cytoplasm and organelles, which then fuses with lysosomes to degrade the contents in the vesicle (Hippert et al., 2006). Necroptosis, a newly characterised mechanism, is a form of regulated necrosis, which is activated by ligands of death receptors through the kinase activity of receptor-interacting protein 1 (RIP1)

(Zhou and Yuan, 2014).

Apoptosis differs from necrosis in a number of ways as described by Macdonald et al. (2004).

First, it is an active process as opposed to an unplanned process induced by cell injury.

Second, apoptotic cells are recognised by phagocytes and removed before they disintegrate.

As a result, there is no surrounding tissue damage of induction of inflammatory responses, which can result in unwanted side effects, such as autoimmune diseases. In contrast, necrotic cells are leaky due to plasma membrane disruption, releasing macromolecules and rapidly disintegrate, thereby inducing inflammation and damage to cellular environment. This is why the induction of apoptosis has been described as a desirable and more efficient strategy in anticancer therapy (Kelly and Strasser, 2011, Strasser et al., 2011). Plants with apoptosis induction properties included several species of Acanthaceae (Table 1.8) but it is still unknown whether C. nutans extracts are able to induce apoptotic cell death in cancer cells.

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Table 1.8. Representatives of Acanthaceae able to induce apoptosis in human cancer cells. Species Plant part Compound Cell line Reference Justicia spicigera Leaf Unknown Cervical carcinoma (HeLa) Alonso-Castro et al. (2012)

Rhinacanthus nasutus Root Rhinacanthone HeLa Siripong et al. (2009)

Ruellia tuberosa Leaf Unknown Hepatocellular carcinoma (HepG2) Dey et al. (2013)

Andrographis paniculata Stem & leaf Unknown Oropharyngeal epidermoid Saengkhae and Meewassanasuk carcinoma (KB) (2009) Andrographolide Breast (TD-47) Sukardiman et al. (2007)

Acanthus ilicifolius Leaf Unknown HepG2 Rajamanickam et al. (2014)

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Apoptotic cells display a very characteristic morphology (Figure 1.7A). Cells undergoing early apoptosis show shrinkage, pyknosis (chromatin condensation) (Kerr et al., 1972) and rippled nuclear contours (Chen et al., 2001). With cell shrinkage, the cells are smaller in size due to cellular volume reduction, the cytoplasm is dense and the organelles are more tightly packed (Elmore, 2007). The most distinct characteristic feature of apoptosis is pyknosis and during the early phase of this event, the nuclear material aggregates peripherally at the nuclear membrane although there can also be uniformly dense nuclei (Elmore, 2007).

Extensive plasma membrane blebbing, followed by karyorrhexis (fragmentation of the nucleus) happen during the latter stage of apoptosis. Eventually, separation of cell fragments into apoptotic bodies occurs during a process called budding. The plasma membrane bound apoptotic bodies consist of cytoplasm with tightly packed organelles with or without nuclear fragments, which are subsequently phagocytosed by macrophages, parenchymal or neoplastic cells and degraded within phagolysosomes (Elmore, 2007).

In necrotic cells, some major morphological changes include swelling; formation of cytoplasmic vacuoles and blebs; distended endoplasmic reticulum; condensed, swollen or ruptured mitochondria; disaggregation and detachment of ribosomes; swollen and ruptured lysosomes; and eventually disruption of the plasma membrane, resulting in the loss of cell membrane integrity (Figure 1.7B) (Kerr et al., 1972, Elmore, 2007).

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Figure 1.7. Morphological changes in apoptotic (A) and necrotic (B) cells. Adapted from Gewies (2003).

1.3.9.3.1 Cellular pathways of apoptosis

Apoptosis was first described as a series of genetically controlled events, which plays a role in controlling cell proliferation and removing damaged cells (Kerr et al., 1972). Any disruption of apoptotic activity leads to abnormal cell growth, resulting in tumourigenesis.

Cells may be induced to undergo apoptosis by extracellular stimuli e.g. death factors, activating the extrinsic signalling pathway (Figure 1.8A) or by internal physical/chemical stressors, such as DNA damage and oxidative stress, activating the intrinsic signalling pathway (Figure 1.8B). The extrinsic pathway requires the ligation of death receptors by death ligands outside the cell, resulting in the assembly of adaptor molecules and pro- caspase-8 activation. This leads to the activation of effector caspases-3, -6 and -7. The intrinsic pathway requires stimuli from inside the cell leading to the activation of pro- apoptotic proteins, such as BAX, BAD and BAK. This then induces disruption of

CHAPTER 1 mitochondrial polarity, resulting in the release of cytochrome c, which binds to APAF-1 in the apoptosome causing cleavage and activation of caspase-9. Consequently, apoptosis occurs through the activation of downstream caspases. Sometimes, both pathways may be activated by cross-talking from the extrinsic pathway via the activation of BID, which disrupts the mitochondrial polarity (White, 2011). Oncogenes (e.g. MYC and RAS), mutated tumour suppressor genes (e.g. PTEN and TP53) as well as anti-apoptotic proteins, including Bcl-2,

Bcl-xL, XIAP, cIAP-1 and -2, disrupt the regulation of apoptosis by impinging the activation of caspases.

Figure 1.8. The extrinsic (A) and intrinsic (B) caspase activation cascades, leading to apoptosis. Adapted from White (2011).

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1.3.9.4 Detection of cell death

1.3.9.4.1 Cell surface markers

Cell surface markers (phosphatidylserine (PS) exposure) are commonly employed to detect cell death induced by plant extracts using cytofluorometry (Engel et al., 2014, Jiang et al.,

2014, Nair et al., 2014) and confocal microscopy (Ismail et al., 2013, Issara-Amphorn and T-

Thienprasert, 2014). Both techniques have high sensitivity and can detect single cells (Krysko et al., 2008). PS is an important aminophospholipid membrane component that is held in the inner leaflet of the plasma membrane of living cells. During apoptosis, PS is actively externalised to the outer surface of the plasma membrane, signalling the requirement for removal (Krysko et al., 2008). A possible mechanism for PS externalisation may be due to a coordinated increase in phospholipid turnabout through the activation of aminophospholipid translocase (Williamson and Schlegel, 2002). PS can be used as an early apoptotic marker through the binding of Annexin V, a Ca2+-dependent phospholipid-binding protein, conjugated with various fluorochromes, such as FITC and green fluorescent Annexin V.

Combining Annexin V and propidium iodide (PI) or 7- aminoactinomycin D (7-AAD), a fluorescent intercalator that has a strong affinity for DNA, can help to distinguish between early apoptosis and necrosis (Krysko et al., 2008). During early apoptosis, there is a lag period between PS positivity and PI/7-AAD positivity, whereas during necrosis, both events coincide (Krysko et al., 2008). However, necrotic cells can have PS externalisation, while remaining PI/7-AAD negative (Krysko et al., 2004), resulting in false positive events. This may be due to the weaker necrotic signalling favouring the detection of PS exposure with the absence of rapid membrane permeabilisation (Krysko et al., 2008), but the exact reason is not clear. Moroever, late apoptotic cells have high affinity for PI/7AAD but not Annexin V, which is similar to the uptake of stains by necrotic cells (Brauchle et al., 2014). Therefore, staining with Annexin V and PI/7-AAD may not discriminate between apoptosis, especially

CHAPTER 1 the late stage, and necrosis (Brauchle et al., 2014). Other techniques, such as transmission electron microscopy should be used to further characterise and confirm cell death types.

1.3.9.4.2 Transmission electron microscopy

Electron microscopy can identify the ultrastructural changes that occur during apoptosis and it is able to discriminate apoptosis from traumatic cell death (necrosis) (Kerr et al., 1972,

Wyllie et al., 1980). Transmission electron microscopy (TEM) is considered the “gold standard” as the most accurate technique for the identification of apoptotic cells (Krysko et al., 2008). It offers better definition of cell morphology with higher resolving power, thus providing much more detailed information at cellular and subcellular levels (Elmore, 2007).

TEM has been used to investigate the mode of cell death in cancer cells induced by plant extracts (Tan et al., 2005, Ait-Mohamed et al., 2011, Echiburú-Chau et al., 2014). However, the high capital, operating and maintenance costs as well as the requirement for highly specialised skills to operate, hinder its used in most laboratories for the detection of cell death.

1.3.9.5 Current cancer therapy

The main approaches for cancer treatment are surgery, chemotherapy, radiation, hormones and immunotherapy, often supplemented by complementary and alternative therapies, such as herbal medicines (Monteiro et al., 2014). Among these approaches, chemotherapy is the most common approach used but most chemotherapeutic drugs create significant problems (Tan et al., 2011). These problems include severe toxicity due to unselective activity of the cytotoxic drugs against tumour cells, affecting healthy cells as well (Chua, 2007); limited efficacy and multidrug resistance due to a surge of survival signals stimulated by cytotoxic drugs (Wu and

Singh, 2011). Therefore, there is a strong emphasis for the discovery and development of selective chemotherapeutic drugs.

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1.3.9.6 Plant sources of anticancer agents

Natural plant products been considered to produce lesser side effects in general than synthetic drugs and therefore, they are one of the main focuses in anticancer drug discovery research

(Fennell et al., 2004). For instance, curcumin from Curcuma longa (Zingiberaceae) reduces toxicity induced by anticancer agents and sensitizes chemo-resistant cancer cells through synergistic effects (Tan et al., 2011). Furthermore, plant-derived natural products containing bioactive phytochemicals still contribute to the overall total number of new chemical entities

(NCE) that continue to be introduced as prescription drugs (Table 1.9) (Salim et al., 2008).

Approximately 48% of the anticancer drugs are either natural products or directly derived therefrom, with plants being the main source of effective anticancer agents (Cragg and

Newman, 2005, Newman and Cragg, 2012). Plants have been long used for cancer treatment and local and traditional knowledge have been the initiator for many drug development studies (Heinrich and Bremner, 2006). For example, Hygrophila spinosa (Acanthaceae) is one of the medicinal plants used by the indigenous villagers in Mahabubnagar, India for the prevention and treatment of cancer (Mumtaz et al., 2015). An in vivo study on H. spinosa found that the crude petroleum root extract showed antitumour activity in Ehrlich ascites carcinoma and sarcoma-180 bearing mice by suppressing the tumour fluid volume (~ 50% reduction in cell voume) and increasing life span of the mice (Mazumdar et al., 1997). The bioactive compounds of H. spinosa still remain to be identified and may be developed into functional anticancer drugs.

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Table 1.9. Plant-derived drugs currently used in cancer treatment. Plant species Compound Generic Trade name Cancer treated Clinical trial Reference class name phase Camptotheca acuminata Alkaloids Irinotecan Camptosar ® Colorectal II/III Fuchs et al. (2006)

Topotecan Hycamtin ® Ovarian, cervical, small II/III Robati et al. (2008) cell lung

Catharanthus roseus Vinca alkaloids Vinblastine Alkaban-AQ ® Germ cell tumours, III/IV Tsao and Stewart Velban ® Hodgkin’s lymphoma, (2009) breast Vincristine Oncovin ® Paedetric and adult III/IV Tsao and Stewart Vincasar Pfs ® acute leukaemias, (2009) Hodgkin’s and non- Hodgkin’s lymphoma, Wilm’s tumour, Ewing’s sarcoma, neuroblastoma, rhabdomyosarcoma

Dysoxylum binectariferum Flavoalkaloid Flavopiridol Experimental Experimental I/II Jain et al. (2012)

Taxus brevifolia Diterpenes Paclitaxel Taxol ® Ovarian, breast, lung, III/IV Weaver (2014) Onxal ® Kaposi’s sarcoma, Docetaxel Taxotere ® Breast, head and neck, III/IV Cline et al. (2013) gastric, prostate, non- small cell lung

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1.3.9.7 Screening of medicinal plants

1.3.9.7.1 Random approach

A random approach is usually used for the screening of plants that are randomly selected for the discovery of new anticancer agents and it has of two types of screening methods: i) Qualitative screening of class of compounds using chemical tests (e.g. Shinoda’s for

flavonoids, ferric chloride for phenols): Although this method is simple to perform, it

provides no information on the biological activity (Katiyar et al., 2012). Most

researchers used this method as a preliminary screening of plant compounds, which can

provide essential information for future bioactivity studies (Vaghasiya et al., 2011,

Yadav and Agarwala, 2011, Ndam et al., 2014). ii) Screening of randomly selected plants for selected bioassays: This method screens for

phytochemicals and the biological activity of the selected plant species, including that

of Acanthaceae (Lindholm, 2005, Mothana et al., 2009, Falodun et al., 2011, Khatun et

al., 2014, Moustafa et al., 2014). The US National Cancer Institute (NCI) studied about

35,000 plant species for anticancer activity using this method, which resulted in two

successful discoveries of clinical-used paclitaxel and camptothecin isolated from T.

brevifolia and C. acuminate, respectively (Katiyar et al., 2012). This method provides

more details on the bioactivity of the plant species (Katiyar et al., 2012).

1.3.9.7.2 Ethnopharmacological approach

An ethnopharmacological approach highly depends on empirical experiences related to the use of botanical drugs for the discovery of biologically active NCE (Katiyar et al., 2012). It involves the observation, description and experimental investigations of indigenous drugs, based on a wide range of disciplines, including enthobotany, medical anthropology and pharmacology (Ganesan, 2008). Studies on traditionally used medicinal plants have

CHAPTER 1 employed this approach to confirm bioactivity (Savigni, 2010, Fadeyi et al., 2013, Hernández et al., 2013). This approach has resulted in the isolation of andrographolide from A. paniculata (Acanthaceae), morphine from Papaver somniferum (Papaveraceae), berberine from Berberis aristata (Berberidaceae) and picroside from Picrorrhiza kurroa

(Plantaginaceae) (Katiyar et al., 2012).

1.3.9.7.3 Traditional medicine system approach

Codified traditional medicine systems, such as traditional Chinese medicine (TCM) and ayurveda, use botanicals as main sources for medicines (Katiyar et al., 2012). These systems are considered different from ethnomedicine in that the later is based on empirical experiences, whereas the former are based on strong conceptual foundations of human physiology and pharmacology gathered throughout history (Katiyar et al., 2012). Therefore, the traditional medicine system is standardised and widely institutionalised, whereas ethnomedicinal practices are localised to small communities (Katiyar et al., 2012). Several studies (Gacche et al., 2011 , Zhang, 2012) have used this approach to screen for anticancer activity, with some reported the discovery of novel compounds, which can be potential anticancer agents. For instance, compounds 33-hydroxyepigambogic acid and 35- hydroxyepigambogic acid isolated from Garcinia species (Clusiaceae) inhibited the cell growth of human lung cancer cells (NCI-H1650) by targeting the signal transducers and activators of transcription (STAT)/ Janus kinase (JAK) signalling pathway (Xu et al., 2015).

1.3.9.8 Anticancer activity of C. nutans

The leaf extracts of C. nutans have been shown to have anticancer activity against several cancer cell lines. The crude chloroform leaf extract at 100 µg/mL showed significant antiproliferative activity against the K-562 and Raji cells at 72 h with 91.3% (IC50: 47.70

CHAPTER 1

µg/mL) and 89.0% (IC50: 47.31 µg/mL) inhibition, respectively (Yong et al., 2013).

Furthermore, the same concentration of the extract also showed > 50% antiproliferative activity against the human lung (NCI-32) (55.8%) and HeLa (56.7%) cancer cells at 72 h

(Yong et al., 2013). The crude petroleum ether C. nutans leaf extract showed cytotoxicity against the HeLa and K-562 cells with 72 h IC50 values of 18 and 20 µg/mL, respectively

(Arullappan et al., 2014). However, the leaves extracted with more polar solvents, such as methanol exhibited lower cytotoxicity against the K-562 cells with a 72 h IC50 value > 100

µg/mL. This was also observed with the crude aqueous leaf extract, where the antiproliferative activity against the K-562 and Raji cells was < 50% (Yong et al., 2013).

These results suggest that the crude chloroform and petroleum ether C. nutans leaf extracts may possess more potent anticancer compounds than that of methanol, aqueous and ethyl acetate extracts. Moreover, the bioactive compounds that acted against the tested cell lines may be non- to mid-polar in nature, according to the polarity of solvents used. However, further studies on other cancer cell lines are needed to confirm these findings. Additionally, these studies also did not examine the mechanism of actions of the extracts against the cancer cells. Studying on the underlying molecular mechanisms and cellular actions (e.g. cell death) of C. nutans extracts is important and helps to discover more therapeutic values of the plant.

1.4 Overview and aims of study

Despite the long traditional use of C. nutans by communities throughout Asia, little is known about the genetic diversity and to what extent it may influence the phytochemical constituents and the bioactivity of C. nutans. Besides, the plant is often confused with A. paniculata because of similar common name. Although both plants have different floral characteristics,

C. nutans rarely blooms. The lack of flowers may be due to extended periods of vegetative propagation by stem cuttings, where growers rapidly and continually harvest the plant during

CHAPTER 1 the juvenile stage to meet the consumer demand, resulting in the inability of the plant to reach maturity (i.e. developing reproductive features). Without having the reproductive features, such as flowers, discrimination between C. nutans and A. paniculata is difficult. Therefore, other alternative identification methods, such as the use of molecular markers are necessary.

The concentration of active compounds in a medicinal plant species is the most important aspect of medicinal plant phytochemistry as low levels of bioactive secondary metabolites can reduce the therapeutic efficacy of the plant. The environment influences the concentration and composition of phytochemicals, but this effect is still unknown on the phytochemical content, particularly phenolics and flavonoids in C. nutans leaves. In addition,

C. nutans leaves are often dried to prolong storage life and the dried leaves are steeped in hot water before being consumed as a herbal tea. This processes can affect the level of phytochemicals in C. nutans leaves and hence, changing its medicinal value. Currently, little is known about the effect of drying temperatures and extraction solvents on the total phenolic and flavonoid content in C. nutans. It is important to optimise postharvest methods to preserve the active phytochemical content in C. nutans dried leaves.

Therefore, this study aimed at addressing these knowledge gaps by investigating the genetic variation, effect of different factors on the phytochemical content and anticancer activity of C. nutans using different interdisciplinary approaches. The specific aims were to:

1) assess the genetic similarity among C. nutans and genetic differences between C.

nutans, C. siamensis and Andrographis paniculata (Chapter 2).

2) compare the total phenolic and flavonoid content in C. nutans leaf samples collected

from different locations with varying environmental characteristics (Chapter 3).

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3) examine the effect of postharvest factors, including drying temperature and extraction

solvent on the total phenolic and flavonoid content in C. nutans leaves (Chapter 4).

4) investigate the effect of extraction solvent on the the viability of selected human

cancer cell lines and evaluate the anticancer activity (including the mode of cell death

induced) of the most active crude C. nutans leaf extract against the most sensitive cell

line (Chapter 5).

5) compare the cytotoxicity of crude methanol C. nutans extratcs of leaves from

different locations with varying environmental characteristics against D24 melanoma

cells and evaluate the anticancer activity (including the mode of cell death induced) of

the most active C. nutans sample against D24 melanoma cells (Chapter 6).

CHAPTER 2

Genetic homogeneity of vegetatively propagated Clinacanthus nutans (Burm. f.) Lindau

(Acanthaceae)

2.1 Introduction

Angiosperms can reproduce either by sexual or asexual reproduction, and most plant species are capable of both. In sexual propagation, reproduction is through fertilisation of the pollen and ovaries to produce fruits containing seeds. Sexual reproduction increases offspring genetic diversity and dispersal potential for seeds and pollens through various dispersal mechanisms (Wilk et al., 2009), which reduces overcrowding and competition compared with ramets. Vegetative propagation involves asexual reproduction, which includes grafting, micropropagation, budding, reproduction from storage organs (e.g. bulbs, corms, tubers, and rhizomes), and cuttings from stems and roots, which results in a clonal population consisting of genetically uniform clones derived from a single individual. Clonal propagation limits gene flow through pollen and seed (Wilk et al., 2009). Therefore, clonality is often viewed to reduce genetic diversity (Hamrick and Godt, 1989), which may lead to decreased resistance to disease and adaptability to the changing environment but is favourable in horticultural industries because agriculturists, breeders and farmers desire consistent superior plant characteristics (genotype and phenotype). The reduction of genetic diversity of clonal populations is a generalisation, which need not always be true.

Clinacanthus nutans (Burm. f.) Lindau is a highly sought-after medicinal plant with potent pharmacological activities (Table 1.6, Section 1.3.8). Growers vegetatively propagate the plant by stem cuttings to meet consumer demand. However, little is known about the genetic diversity of C. nutans and whether propagation method influences its genetic characteristics.

CHAPTER 2

C. nutans is marketed under the trade name “Sabah Snake Grass” and is generally sold as powdered or whole dried leaves in Malaysian herbal markets. The plant is often misidentified with the closely related C. siemensis and confused with another locally available medicinal plant in the same family, Andrographis paniculata (Burm. f.) Nees, which has the common name “Indian Snake Grass”. All have similar growth habits (Figure 2.1 A, D and G) and leaf appearance (Figure 2.1B, E and I) but are distinguished by the flowers (Figure 2.1C and F), stems (Deng et al., 2011) and bitter taste (Chandrasekaran et al., 2009) with different medicinal purposes.

Morphological and taste characteristics can be subjective and do not differentiate between closely related individuals, particularly for populations of clonal plants consisting of genets and ramets that often occur with vegetative propagation. Traditional markers based on morphological or allozyme variation have limited abilities to differentiate between genetically similar individuals and have been resolved using DNA-based markers that are efficient in detecting genotypic distribution in clonal populations (Esselman et al., 1999,

Keller, 2000). PCR-based DNA markers have been used to assess genetic homogeneity in sexually propagated Acanthaceae representatives (Chua, 2007, Mori et al., 2010, Behera et al.,

2012, Suwanchaikasem et al., 2012, Wee et al., 2013) but not on C. nutans that is commonly vegetatively propagated.

Little information is available not only on the genetic characteristics of C. nutans, but also on other species of Acanthaceae that are vegetatively propagated; and the genetic differences between C. nutans and A. paniculata. Therefore, the current study aimed to i) assess the genetic characteristics, particularly the similarity of C. nutans propagated by stem cuttings as

CHAPTER 2 a representative of Acanthaceae from different regions between and within countries, ii) evaluate the genetic differences between C. nutans. C. siemensis and A. paniculata as an alternative identification method, and iii) compare it to sexually propagated A. paniculata using RFLP, RAPD and microsatellite markers.

Figure 2.1. Macromorphology of plants investigated in this study. C. nutans growth habit (A), leaves (B) and flower (C); A. paniculata growth habit (D), leaf (E) and flower (F); and C. siamensis leaves (G). Comparisons between C. nutans (indicated with black asterisk) leaves with A. paniculata (indicated with red asterisk) (H) and C. siamensis (indicated with blue asterisk) (I).

CHAPTER 2

2.2 Materials and methods

2.2.1 Plant materials

Fresh leaves of 12 samples of C. nutans grown under different environmental conditions

(Table 2.1) were collected from Peninsular Malaysia (CP), East Malaysia (CE), Thailand

(CT), and Vietnam (CV) (Figure 2.2) between November, 2012 and January, 2013. Leaves of two samples of A. paniculata were collected from Seremban, Negeri Sembilan, Malaysia

(AP1 and AP2). C. siamensis from Thailand and Momordica cochinchinensis from Australia were used as outgroups to compare C. nutans genotypes and C. nutans and A. paniculata, respectively. All leaves were thoroughly washed using cold tap water. All samples, except samples CP1 and AP2 were air dried for at least seven days.

Figure 2.2. Sampling sites of C. nutans, C. siamensis and A. paniculata in Malaysia, Thailand and Vietnam.

CHAPTER 2

Table 2.1. The taxon, sample codes, collection sites, environmental and growth conditions of collected samples. Taxon Sample Country State/Province Region Environmental conditions Tm (°C) Elv (m) H L Rn (mm) C. nutans CP1 Malaysia Negeri Sembilan Seremban 66.6 31.3 23.0 2124.0 CP2 Malaysia Negeri Sembilan Seremban 83.4 31.2 22.4 2010.0 CE1 Malaysia Sabah Sandakan 158.7 30.8 22.9 2973.0 CE2 Malaysia Sabah Sandakan 74.9 30.8 22.9 2973.0 CE3 Malaysia Sabah Tawau 6.8 30.7 23.2 1975.0 CE4 Malaysia Sabah Kota Kinabalu 9.7 30.7 23.3 2818.0 CT1 Thailand Nakhon Pathom Map Khae 8.1 32.7 22.7 1237.0 CT2 Thailand Nakhon Pathom Map Khae 8.1 32.7 22.7 1237.0 CT3 Thailand Nakhon Pathom Salaya 4.3 32.5 23.4 1334.0 CT4 Thailand Chiang Mai San Sai 309.5 31.5 19.6 1191.0 CT5 Thailand Chiang Mai Chiang Dao 439.4 30.6 18.7 1261.0 CV1 Vietnam Ho Chi Minh Ho Chi Minh 2.2 31.9 23.1 1873.0 C. siamensis CS Thailand Nakhon Pathom Salaya 4.3 32.5 23.4 1334.0 A. paniculata AP1 Malaysia Negeri Sembilan Seremban 83.8 31.2 22.4 2010.0 AP2 Malaysia Negeri Sembilan Seremban 83.8 31.2 22.4 2010.0 M. cochinchinensis MC Australia New South Wales Newcastle 79.0 25.0* 25.0* NA Abbreviation: Elv = elevation; Tm = mean annual temperature; H = highest; L= lowest; Rn = mean annual rainfall; * = greenhouse conditions.

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2.2.2 DNA extraction

Total DNA from samples CP1 and AP2 was extracted in fresh form, using the GF-1 Plant

DNA Extraction Kit (Vivantis Technologies, Malaysia) according to the manufacturer’s protocol. The FavorPrep Plant Genomic DNA Extraction Mini Kit (Favorgen Biotech Corp.,

Taiwan) was used to extract the total DNA from all the other dried samples with slight modifications of the extraction protocol provided by the manufacturer to improve the DNA yield. Here, 20 mg of dried sample was used instead of 100 mg and then DNA extraction was carried out as outlined in the protocol. Finally, 50 µL of preheated elution buffer was added to the column matrix and centrifuged at 6,600 x g for 2 min to elute the purified DNA.

2.2.3 PCR-RFLP

Primer pair rbcL1F-724R (Applied Biosystems, Australia) was used to amplify the ribulose-

1,5-biphosphate carboxylase/oxygenase large subunit (rbcL) region (ca. 700 bp) (Olmstead et al., 1992). A total volume of 25 µL PCR mixture contained the following: 12.5 µL 2X GoTaq

Green Master Mix (Promega Corporation, Australia), 1.0 µL 0.4 mM forward primer, 1.0 µL

0.4 mM reverse primer, 1.0 µL DNA, and 9.5 µL nuclease-free water. The control included all of the PCR reagents except the DNA template. Amplification was performed in a G-Storm

GS1 thermal cycler (Gene Technologies, England), where the reaction consisted of an initial denaturation step at 95˚C for 2 min, 30 cycles of denaturation at 94˚C for 1 min, annealing at

55˚C for 30 s, elongation at 72˚C for 1 min, followed by a final elongation step at 72˚C for 7 min and held at 4˚C. Amplicons were analysed by gel electrophoresis with a 1.4% (w/v) agarose gel in 1X TBE buffer and stained with ethidium bromide (0.5 µg/mL) for visualisation using the Discovery Series Quantity One 1-D Analysis Software (Bio-Rad

Laboratories, USA).

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Aliquots (5 µL) of the PCR products from the rbcL gene were digested with seven restriction enzymes (Table 2.2). The restriction fragments were separated by 2.0% (w/v) agarose gel electrophoresis in 1X TBE buffer, and stained with ethidium bromide (EtBr) (0.5 µg/mL) for visualisation. PCR amplification and restriction of PCR products were repeated twice for all samples for reproducibility.

Table 2.2. Restriction endonuclease digestions of the rbcL 1F and 724R PCR products. Restriction Enzyme Buffer (10X) Sequence 5’ – 3’ Reaction Conditions TaqI (Fermentas) TaqI Buffer with BSA T-CGA 65˚C for 3 hr AGC-T MspI (Fermentas) Buffer Tango with BSA C-CGG 37˚C for 3 hr GGC-C BamHI (Promega) Buffer E G-GATCC 37˚C for 3 hr CCTAG-G HaeIII (Promega) Buffer C GG-CC 37˚C for 3 hr CC-GG HindIII (Promega) Buffer E A-AGCTT 37˚C for 3 hr TTCGA-A XbaI (Biotech) XbaI Reaction Buffer T-CTAGA 37˚C for 3 hr AGATC-T XhoI (Promega) Buffer D C-TCGAG 37˚C for 3 hr GAGCT-C

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2.2.4 RAPD amplification

Of the 22 RAPD primers (Operon Technologies, Australia) screened, three (OPA-09, 11 and

18) that produced polymorphic banding patterns were selected (Table 2.3). RAPD reactions were performed in a total volume of 25 µL containing 12.5 µL 2X GoTaq Green Master Mix

(Promega Corporation), 1.0 µL 0.4 µM primers, 1.0 µL 5-10 ng of genomic DNA, and 10.5

µL nuclease-free water. RAPD amplifications were performed in a G-Storm GS1 thermal cycler (Gene Technologies) programmed for initial denaturation at 94˚C for 2 min, 35 cycles of denaturation at 94˚C for 1 min, annealing at 37˚C for 1 min, elongation at 72˚C for 2 min, followed by a final elongation step at 72˚C for 8 min and held at 4˚C. The amplicons were then subjected to electrophoresis in 1.5% (w/v) agarose gel in 1X TBE buffer and stained with EtBr for visualisation.

2.2.5 Microsatellite amplification

Two microsatellite primers (GTG5 and GACA4) (Operon Technologies) were used to evaluate the genetic relationships in this study (Table 2.3). Microsatellite reactions were performed in a total volume of 25 µL containing 12.5 µL 2X GoTaq Green Master Mix

(Promega Corporation), 1.0 µL 0.4 µM primers, 1.0 µL 5-10 ng of genomic DNA, and 10.5

µL nuclease-free water. Microsatellite amplifications were performed in a G-Storm GS1 thermal cycler (Gene Technologies) programmed for initial denaturation at 94˚C for 5 min,

40 cycles of denaturation at 94˚C for 1 min, annealing at 36˚C for 1 min, elongation at 72˚C for 1 min, followed by a final elongation step at 72˚C for 7 min and held at 4˚C. The amplicons were then separated by electrophoresis in 1.5% (w/v) agarose gel in 1X TBE buffer and stained with EtBr for visualisation.

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Table 2.3. The five primers and the number of corresponding fragments used in the RAPD and microsatellite analyses. Primer Sequence (5’ – 3’) No. of polymorphic Percentage of polymorphic fragments fragments (%) OPA-09 GGGTAACGCC 11 100 OPA-11 CAATCGCCGT 14 100 OPA-18 AGGTGACCGT 11 100 GTG5 GTGGTGGTGGTGGTG 25 100 GACA4 GACAGACAGACAGACA 25 100

2.2.6 Data collection and analysis

Each gel was analysed by manually scoring the presence (1) or absence (0) of bands in individual lanes, generating a binary data matrix. The molecular weights of the bands were estimated based on the DNA marker GeneRuler DNA Ladder Mix (Fermentas, Australia).

Bands of equal molecular weight generated by similar primers were considered to be identical locus. Similarity indices were calculated using the similarity for qualitative data

(SimQual) computer algorithm in NTSYS-pc ver. 2.10e (Rohlf, 2000) using Dice’s similarity coefficient equation (Dice, 1945): Sij = 2a/(2a+b+c), where, Sij = the similarity between two samples, i and j; a = the number of bands present in both i and j; b = the number of bands present in i and absent in j; c = the number of bands present in j and absent in i.

From these similarity indices, the sequential, agglomerative, hierarchic and non-overlapping

(SAHN) clustering method was performed using the unweighted pair group method with average mean (UPGMA) to generate the dendrograms in the same software. The genetic similarity coefficient value between samples ranged from 0 to 1, where 0 indicated no similarity and 1 indicated that the two samples were genetically identical. Binary data from

RAPD and microsatellites were combined for principal component analysis (PCA) using the

Minitab statistical software (Minitab, 2013). Score plots were generated and genetically similar samples were clustered together.

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2.3 Results

2.3.1 PCR-RFLP analysis

PCR amplification using primer pair rbcL 1F-724R produced a ca. 700 bp amplicon. Of the seven restriction endonucleases used, only two (TaqI and MspI) cleaved the 700 bp amplicon

(Table 2.4). Both TaqI and MspI digestions revealed that the restriction fragment patterns were different between C. nutans and A. paniculata but were identical between C. nutans and

C. siamenesis. TaqI endonuclease cut the amplified rbcL gene of C. nutans and C. siamemsis at two restriction sites, producing three fragments, but only one site for A. paniculata, producing two fragments (Figure 2.3A). All fragments when combined totalled the original template size i.e. ~700 bp.

MspI endonuclease cut the amplified rbcL gene of C. nutans, C. siamensis and A. paniculata at only one restriction site and produced the same number of fragments in each species but at different sites and sizes (Figure 2.3B). Restriction digestion of the amplified rbcL gene of C. nutans and C. siamensis generated 200 and 500 bp, whereas that of A. paniculata generated

150 and 550 bp fragments. RFLP marker revealed an identical banding pattern among all samples of C. nutans as well as A. paniculata.

Table 2.4. Restriction endonuclease digestions of rbcL 1F-724R PCR products. Restriction Enzyme Fragment Size (bp) C. nutans C. siamensis A. paniculata Taq I 150 150 250 250 250 450 300 300 Msp I 200 200 150 500 500 550 BamH I No restriction No restriction No restriction Hae III No restriction No restriction No restriction Hind III No restriction No restriction No restriction Xba I No restriction No restriction No restriction Xho I No restriction No restriction No restriction

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Figure 2.3. PCR-RFLP patterns of Taq I (A) and MspI (B) digested rbcL gene. Lane M = 100 bp DNA ladder; lanes CP1, CP2, CE1, CE2, CE3, CE4, CT1, CT2, CT3, CT4, CT5, and CV1 = C. nutans; lanes AP1, AP2 = A. paniculata; lane CS = C. siamensis; lane MC = M. cochinchinensis; lane Ctrl = no template control.

2.3.2 RAPD analysis

Three primers, OPA-09 (Figure 2.4A), OPA-11 (Figure 2.4B), and OPA-18 (Figure 2.4C), produced amplification patterns in all C. nutans, A. paniculata and M. cochinchinensis samples, except C. siamensis (OPA-18 only). Pooled-data revealed that C. nutans was genetically different from A. paniculata and the outgroups (C. siamensis and M. cochinchinensis).

Within species, C. nutans samples were genetically similar but not identical between countries. All the Malaysian C. nutans samples were identical while the four Thai samples were identical and one was not. The Vietnamese sample was separated from other C. nutans.

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Overall, of the 29 loci scored, 100.0% were polymorphic. Eight loci were scored for C. nutans and four loci (50.0%), which were produced from OPA-11 and -18 primers amplifications were polymorphic. All the Thai samples revealed an absence of bands at 950 and 1400 bp. All samples showed an absence of bands at 550 and 1000 bp, except CV1

(Vietnam) and CT5 (Thailand), respectively. In contrast, A. paniculata samples obtained from the same farm but different sampling sites showed genetic variation. A higher number of polymorphic loci was revealed in A. paniculata, where of the 11 loci scored, seven (63.6%) were polymorphic. Fragments produced from RAPD amplification ranged in size from 250 to

2600 bp.

The genetic similarity calculated using Dice’s coefficient in NTSYS software showed no to low genetic similarity between C. nutans and A. paniculata samples, where coefficient values ranged from 0.0000 to 0.2857. Meanwhile, RAPD analysis revealed high genetic similarities between C. nutans samples. The Malaysian samples although from different regions were identical (1.0000). All the Thai samples were also identical except for CT5 with a coefficient value of 0.8889. The genetic similarity coefficient values between the Malaysian and the Thai samples ranged from 0.7273 to 0.8000. The Vietnamese sample showed a higher similarity to the Malaysian C. nutans (0.7692) compared to the four Thai samples (0.7273).

Based on the UPGMA clustering analysis using Dice’s coefficient on pooled-RAPD data, the

14 samples (12 C. nutans and two A. paniculata) were divided into four major divisions at a

0.52 coefficient level (Figure 2.6A). Division I comprised of all the Malaysian samples, division II comprised of all the Thai samples, which further separated into two sub-divisions

(a:CT1-4; b:CT5), and cluster III comprised of the Vietnamese sample. Cluster IV comprised

CHAPTER 2 of the two A. paniculata samples, which were genetically different from each other, with a low similarity coefficient of 0.5333.

Figure 2.4. RAPD amplification products of OPA-09 (A), OPA-11 (B) and OPA-18 (C). Lane M=100 bp DNA ladder; lanes CP1, CP2, CE1, CE2, CE3, CE4, CT1, CT2, CT3, CT4, CT5, and CV1=C. nutans; lanes AP1, AP2=A. paniculata; lane CS=C. siamensis; lane MC=M. cochinchinensis; lane Ctrl=no template control.

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2.3.3 Microsatellite analysis

Two microsatellite primers, GACA4 (Figure 2.5A) and GTG5 (Figure 2.5B), were selected and pooled-microsatellite data was used for genetic analysis. The outcome of the microsatellite analysis was the same as the RAPD analysis in which, (a) C. nutans was genetically different from A. paniculata, C. siamensis and the outgroup; (b) C. nutans samples were genetically diverse and different between countries; and (c) some genetic homogeneity existed for the Malaysian and the Thai C. nutans samples but not all.

Overall, of the 36 loci scored, 100.0% were polymorphic. From these samples, 17 loci were scored for C. nutans and five loci (29.4%), which were produced from both primer amplifications, were polymorphic. Bands 800 and 1350 were absent only in the Malaysian and CT5 samples, respectively. Band 1300 was absent in both the Malaysian and the

Vietnamese samples but present in all the Thai samples. Band 1700 was absent in all samples except the Malaysian samples and band 1800 was absent in the Vietnamese sample as well as in all the Thai samples, except CT5. For A. paniculata, of the 13 loci scored, three (23.1%) were polymorphic. Fragments produced from microsatellite amplification ranged in size from

450 to 2500 bp.

The Dice’s coefficient of similarity showed that there was a low genetic similarity between C. nutans and A. paniculata samples, where the coefficient values ranged from 0.2308 to 0.3077.

Meanwhile, microsatellite analysis revealed a high genetic similarity between the C. nutans samples. All the Malaysian samples were identical (1.0000) but were different from the Thai and the Vietnamese samples. The Thai samples were also identical, except for CT5 with a coefficient value of 0.9333. The genetic similarity coefficient value between Malaysia and

Thailand was high at 0.8667. The Vietnamese sample showed a higher similarity to the Thai

CHAPTER 2 samples (except CT5, 0.8966), with a coefficient value of 0.9655 compared to the Malaysian samples (0.8966).

The dendrogram based on UPGMA cluster analysis using Dice’s coefficient on pooled- microsatellite data showed a different topology from the dendrogram constructed using

RAPD data. The 14 samples (12 C. nutans and two A. paniculata) were divided into three major divisions at a 0.86 coefficient level (Figure 2.6B). Division I comprised all the

Malaysian samples and division II comprised all the Thai and the Vietnamese samples, which was further divided into three sub-divisions (a:CT1-4; b:CV1; c:CT5). Cluster III comprised of the two A. paniculata samples, which were genetically different from each other, with a similarity coefficient of 0.8696.

Combined banding profiles of RAPD and microsatellite markers from PCA clustered all C. nutans samples together with low genetic diversity between countries and homogeneity from

Malaysia (6 samples) and Thailand (4 samples) (Figure 2.6C). Interestingly, A. paniculata samples were also clustered closely together but were not identical and well separated from C. nutans, C. siamensis and M. cochinchinensis. All DNA markers were able to detect genetic difference between genera, but only RAPD and microsatellite markers were able to detect genetic variation within and between species. RAPD and microsatellite markers distinguished differences between and within countries of C. nutans and A. paniculata including distant, closely related and identical samples.

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Figure 2.5. Microsatellite amplification products of GACA4 (A) and GTG5 (B). Lane M=100 bp DNA ladder; lanes CP1, CP2, CE1, CE2, CE3, CE4, CT1, CT2, CT3, CT4, CT5, and CV1=C. nutans; lanes AP1, AP2=A. paniculata; lane CS=C. siamensis; lane MC=M. cochinchinensis; lane Ctrl=no template control.

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Figure 2.6. The relationships among 12 C. nutans, two A. paniculata, one C. siamensis, and one M. cochinchinensis samples according to UPGMA cluster analysis of the RAPD (A) and microsatellite (B) data, and PCA analysis (C) of the combined RAPD and microsatellite data.

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2.4 Discussion

2.4.1 Genetic homogeneity in C. nutans

This is the first study to assess the genetic characteristics of C. nutans samples between and within countries. Low variation was observed between countries and most samples within each country were genetically identical and highly likely to be clones. It is uncertain how these clones could exist from geographically distant sites, borders and terrains such as peninsular Malaysia and Sabah that were separated by the South China Sea. Some possibilities for the genetic homogeneity include: 1) all plants originated from the same parents and have stable persistent genomes that do not evolve rapidly; 2) the genetic diversity of the species may have been low originally and in the long-term domestication process, vegetative reproduction may have contributed to the genetic uniformity or monoculture of the species (Gepts, 2004); 3) inferior or different varieties no longer exist due to agricultural and economic pressures in rural communities; and 4) stem cuttings of the plant were shared among growers and farms.

C. nutans is vegetatively propagated through stem cuttings (Panyakom, 2006) because the method is easy, economical and produces a high multiplication rate. In addition, flowers are rare, which results in severely reduced sexual reproduction and difficulty in crossing genotypes. The lack of flowers may be due to extended periods of vegetative propagation, selection and harvesting practices, where growers rapidly and constantly harvest the plant during the juvenile stage to meet the consumer demand. This may result in the inability of the plant to reach maturity for sexual reproduction but is unlikely as some plants selected in this study were at mature stages (CT2 sample) and had not flowered in its history (Mr

Rattanapong, personal communication, January 27, 2013). Most members of Acanthaceae, especially weedy species produce fruits, or reproduce both by seeds and tubers, for example

CHAPTER 2

Ruellia tuberosa and some are easily propagated by seeds, which are produced in abundance, for example Thunbergia fragans (Meyer and Lavergne, 2004). However, according to

Whistler (2000), most ornamental species in Acanthaceae are propagated by stem cuttings or layers in horticulture, and their fruits are rarely formed in cultivation.

Genetic homogeneity is consistent with other studies that have shown no genetic variation in clonal plants in other plant families but not Acanthaceae, for instance Alternanthera philoxeroides (Amaranthaceae) (Xu et al., 2003) and Eichhornia crassipes (Pontederiaceae)

(Li et al., 2006). Other studies have also revealed low levels of genetic diversity in clonal species or in populations of such species with more intense vegetative reproduction such as

Microtis parviflora (Orchidaceae) (Peakall and Beattie, 1991), Haloragodendron lucasii

(Haloragaceae) (Sydes and Peakall, 1998), Allium sativum (Amaryllidaceae) (Paredes et al.,

2008), Iris sibirica (Iridaceae) (Kostrakiewicz and Wróblewska, 2008), Cypripedium calceolus (Orchidaceae) (Brzosko et al., 2013) and Chlorophytum borivilianum

(Asparagaceae) (Tripathi et al., 2012).

In contrast, a study on Poikilacanthus macranthus (Acanthaceae) revealed high clonal diversity in the species (Bush and Mulcahy, 1999), but this is not the case for C. nutans.

Other studies have also shown unexpected high levels of genetic diversity in clonal plants such as Anemone nemorosa (Ranunculaceae) (Stehlik and Holderegger, 2000), Vaccinium stamineum L. (Ericaceae) (Kreher et al., 2000), V. myrtillus L. (Albert et al., 2003),

Potamogeton maackianus (Potamogetonaceae) (Li et al., 2004) and Goodyera repens

(Orchidaceae) (Brzosko et al., 2013). A hypothesis for the high levels of genetic diversity in clonal species is the presence of sexual reproduction in the founder populations, although it is sporadic and poor (Brzosko et al., 2013). Many populations of clonal species are founded by

CHAPTER 2 seeds, which is the most common means of recruitment of individuals to an existing population, but show increasing vegetative reproduction over time, likely due to limited resource availability or changing environmental conditions (Eriksson, 1993). Such populations can be genetically diverse by preserving genetic diversity of the initial population through clonal reproduction to dominate an area (Yeh et al., 1995). In addition, somatic mutations accumulated from constant division of mitotic cells can occur in clonal plants, as observed in Grevillea rhizomatosa (Proteaceae) (Gross et al., 2012), which would contribute to genetic diversity, but this has not been well studied in Acanthaceae.

2.4.2 Genetic differences between C. nutans, C. siamensis and A. paniculata

The genetic differences between C. nutans, C. siamensis and A. paniculata were investigated and this is the first study to report the genetic diversity between C. nutans and A. panicultata.

At the interspecies level, PCR-RFLP analysis on the rbcL region could not distinguish between the C. nutans and C. siamensis, where results showed identical profiles. This suggests that this technique is unsuitable for the study of the genus Clinacanthus. It has been documented that the rbcL region is highly conserved with low gene substitution rates in perennial angiosperm taxa (Bousquet et al., 1992, Frascaria et al., 1993), which makes it more suited for genetic studies of higher taxonomic levels (intergeneric to subclass) but not lower levels (inter- to intraspecies) (Arca et al., 2012). Likewise, Kunsorn et al. (2013) also examined the genetic diversity between the two species by genome sequencing of the ITS region. The authors revealed 97-99% genetic similarity between C. nutans and C. siamensis, and suggested the region might not be useful for distinguishing the two species. Consequently,

RAPD and microsatellite marker techniques were used to further assess the genetic difference between C. nutans and C. siamensis in this study. The two techniques were able to discriminate the two species, showing considerable genetic diversity between them. This

CHAPTER 2 suggests that both marker techniques are more suitable for the analysis of genetic diversity in the genus Clinacanthus than RFLP. On the other hand, all molecular markers used in this study showed unambiguous differentiation between C. nutans and A. paniculata based on

DNA fingerprinting. Results imply that these markers can be considered as an alternative method to support traditional identification, such as vegetative characteristics, reproductive morphology and taste (Leyew, 2011) that can be subjective to interpretation. This could be a useful validation tool for ambiguous samples in situ and is independent of visual characteristics that may be compromised due to environmental factors. The disadvantage of molecular differentiation is its limited availability to growers particularly from developing nations compounded by impoverished rural regions where these medicinal plants are most popular. Even the collection of plants used in this study was with great difficulty since farmers only spoke native Asian languages and had minimal literacy skills so the technology may not be embraced widely.

2.4.3 Genetic variation in A. paniculata

By comparison, genotypes of A. paniculata, a member of the same family Acanthaceae, which reproduces sexually, were genetically different, although collected from the same farm.

Previous studies on the genetic diversity in A. paniculata also showed considerable genetic variation within the species using RAPD, SSR and AFLP techniques (Maison et al., 2005,

Chua, 2007, Wijarat et al., 2012). White flowers of A. paniculata are frequently observed and the plant is commonly propagated by seeds obtained from mature pods (Talei et al., 2011).

Cross-pollination and sexual reproduction may contribute to the greater genetic variation within the species compared with vegetative reproduction in C. nutans.

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2.5 Conclusion

In conclusion, the current study improved our understanding of the distribution and extent of genetic variation within and between C. nutans, C. siamensis and A. paniculata using different molecular marker techniques. RAPD and microsatellite markers were proven to be reliable alternative methods for plant identification, but not RLFP, particularly at the interspecies level.

In the current situation, genetics should not play a major role in the designing of conservation management plans for C. nutans since there is a lack of genetic diversity in the species even though they were geographically distant. The most appropriate measure is to minimise continuous vegetative propagation of the plant and maintaining it through mature stage with the development of reproductive features, where possible. With these features, C. nutans can be multiplied by sexual reproduction, thus, increasing its genetic diversity, which in turn, may aid in long-term survival and evolution of the species.

CHAPTER 3

Variation in the total phenolic and flavonoid content of Clinacanthus nutans (Burm. f.)

Lindau (Acanthaceae) leaves obtained from different locations

3.1 Introduction

Bioactive secondary metabolites can be produced from either the vegetative (bark, leaves and roots) or the reproductive (flowers, fruits and seeds) components in response to environmental stimuli, such as herbivore attack, pathogen infections or nutrient deprivation

(Kennedy and Wightman, 2011). These secondary metabolites are often unique to specific plants where they are not involved in the plants’ primary metabolic requirements. Instead these compounds enhance the overall ability of the plant to survive through ecological interactions with the environment (Harborne, 1993). Environmental factors can therefore substantially influence the concentration of plant secondary metabolites (Radušienė et al.,

2012) and is strongly dependent on the growing conditions (Akula and Ravishankar, 2011). It has been suggested that plants of the same species occurring in different environments may differ significantly in their secondary metabolite content of particular (Szakiel et al., 2011).

For instance, the variation in the levels of secondary metabolite production in Fagopyrum tartaricum (Polygonaceae) from different locations in China was observed (Guo et al., 2011).

It is also suggested that a wide range of environmental factors changes are observed as the elevation of the natural growing site changes. This includes the air temperature, rainfall, daily thermal amplitudes, soil fecundity, wind speed, atmospheric pressure and radiation intensities

(Zidorn, 2010). For instance, studies on Arnica montana (Asteraceae) (Spitaler et al., 2006,

Spitaler et al., 2008) and Matricaria chamomilla (Asteraceae) (Ganzera et al., 2008) have shown a higher concentration of secondary metabolites in extracts obtained from plants grown at high altitude compared to those grown at low altitude.

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One of the general classes of plant secondary metabolites are the phenolics, which generally possess hydroxyl groups attached to aromatic rings (Dai and Mumper, 2010). The phenolics class can be further divided into two subclasses, namely the non-flavonoids and the flavonoids. Approximately 50% of the estimated 8000 phenolics currently known are flavonoids (Harborne et al., 1999). Non-flavonoids include simple phenols, benzoic acids, hydrolysable tannins, acetophenones, phenylacetic acids, cinnamic acids, lignans, coumarins, benzophenones, xanthones, stilbenes and secoiridoids (Khadem and Marles, 2010). Flavonoid compounds, which are characterised by a phenylbenzopyran chemical moiety, possess a basic structure consisting of a C15 (C6-C3-C6) skeleton linked to a chroman ring (Pereira et al.,

2009). The flavonoids include flavanones, flavones, dihydroflavonols, flavonols, flavan-3-ols, isoflavones, anthocyanidins, proanthocyanidins and chalcones (Khadem and Marles, 2010).

In recent years, there has been increased interest in edible naturally occurring plant-based sources rich in phenolics due to the health associated with these compounds (Hooper and

Cassidy, 2006) including their antioxidant, anti-inflammatory (Huang et al., 2009) oestrogenic, enzyme inhibition, antimicrobial, antiallergic and anticancer activities (Cushnie and Lamb, 2005). This has led to many studies focusing on the identification and quantification of various classes of bioactive phytochemicals. In addition, numerous studies have also focussed on the plant postharvest processing methods on plants to retain maximum levels of bioactive compounds. However, this remains an understudied area in terms of the enhancement of these valuable compounds, such as exploring the effects of breeding and selection or the influence of preharvest growth conditions on the concentrations (Joubert et al.,

2008). Rapid screening and determination of the concentration of phenolics is necessary and therefore, selection of methods to perform these tasks is an important strategy not only for researchers, but, also for breeders, to minimise time and costs.

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The commonly used colorimetric methods to determine the total contents of phenolics and flavonoids using a UV-vis spectrophotometer include the Folin-Ciocalteu and the AlCl3 methods, respectively. The Folin-Ciocalteu method is based on the oxidation of the phenolic compounds with phosphoric molibdotungstate (Singleton and Rossi, 1965), while the AlCl3 method is based on the formation of complexes between the AlCl3 compound and the keto and hydroxyl groups of flavonoids (Joubert et al., 2008). Other instrumental chromatography methods have also been used to separate and determine phenolics in plants. HPLC is one of the most popular methods used to analyse phenolic composition. In general, HPLC separations are based on C-18 reverse-phased chromatography (stationary phase) and a binary solvent gradient (mobile phase), which usually consists of an aqueous solution of acid and an organic solvent (Mišan et al., 2011). Although HPLC methods have been widely used to study the bioactive compounds in various plants, the methodology requires a high initial capital cost, involves complicated operating procedures and is usually time-consuming. These factors could be restrictive to breeders and some manufacturers. In contrast, UV-vis spectrophotometers are not cost- and labour-intensive and they are accessible in most laboratories, so colorimetric assays could potentially offer plant breeders and industries a simple alternative.

The medicinal plants of Acanthaceae are rich in secondary metabolites, such as flavonoids, glycosides, alkaloids, phenols, steroids, saponins and tannins (Kaliswamy et al., 2015).

Previous chemical studies report the presence of several phytochemical compounds in the different parts and extracts of C. nutans (Table 1.4, Section 1.3.6). Despite the known phytochemicals and pharmacological activities (Table 1.6, Section 1.3.8) of C. nutans, it remains unclear whether collection sites with different environmental characteristics have an effect on the concentration of secondary metabolites, especially phenolics and flavonoids in

CHAPTER 3 the leaves. Therefore, the aims of the present study were to: i) determine and compare the total phenolic (TPC) and flavonoid content (TFC) in C. nutans leaves collected from different regions between and within countries with varying environmental characteristics and ii) examine the correlation between the colorimetric and HPLC methods to investigate the TPC and TFC.

3.2 Materials and methods

3.2.1 Plant materials

Fresh leaves of 11 C. nutans samples grown under different environmental conditions (Table

2.1, Section 2.2.1) were collected from Peninsular Malaysia (CP), East Malaysia (CE),

Thailand (CT), and Vietnam (CV1) (Figure 3.1). Geographic data, including elevation, annual temperature (high, low and mean) and rainfall of sampling sites was obtained from

DIVA-GIS version 7.5 software (Hijmans et al., 2012). Prior to sample extraction, all leaf pieces were thoroughly washed using cold tap water. All samples were air dried in the shade for seven days at 22°C and stored as the whole leaf in air-tight bags in darkness at 22°C until further analysis.

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Figure 3.1. Collection sites of C. nutans samples from Peninsular and East Malaysia, Thailand and Vietnam.

3.2.2 Preparation of crude extracts

One gram of dried powdered C. nutans leaves from each location was extracted with 50 mL of methanol (MeOH) (Merck, Germany), on an orbital shaker at a speed of 200 rpm at 22°C for seven days. The extracted solution was decanted, filtered with Whatman No. 1 filter paper and concentrated under reduced pressure using a rotary evaporator (Buchi, Switzerland) to produce the crude MeOH dried extract, which was then weighed and stored at -20˚C for the

TPC, TFC and HPLC analyses.

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3.2.3 Determination of the TPC

The TPC of all samples was determined from a gallic acid standard curve developed according to the Folin-Ciocalteu method described by Ozkok et al. (2010). In order to generate a standard curve, standard solutions of five concentrations (0, 0.025, 0.05, 0.075, and 0.1 mg/mL) were prepared in 70% MeOH from a stock solution of 25% (w/v) gallic acid

(Sigma, USA) in 70% MeOH. Then, 1 mL of each standard solution was mixed with 5 mL of

10% aqueous dilution of Follin-Ciocalteu reagent (Sigma) for 1 min and allowed to stand for another 8 min, followed by the addition of 4 mL of 75 g/L anhydrous Na2CO3 (BDH, London) solution. The solution was then mixed thoroughly for 1 min and incubated in a water bath at

40°C for 30 min. After incubation, absorbance was measured at 765 nm using a Cary 60 UV-

Vis spectrophotometer (Agilent Technologies, USA) and results were expressed as gallic acid equivalent (mg GAE/g of dry extract). Samples were prepared in MeOH (1 mg/mL) and measured following the same procedure as the preparation of standards. TPC of samples was calculated using Equation 3.1.

C = (c × V)/ m (3.1)

where, C = TPC (mg GAE/g dry extract), c = concentration of gallic acid established from the calibration curve (mg/mL), V = volume of extract (mL), and m = weight of dry plant extract

(g).

3.2.4 Determination of the TFC

The TFC of all samples was determined from a quercetin standard curve developed according to the AlCl3 colorimetric method (Ozkok et al., 2010). In order to generate a standard curve, standard solutions of five concentrations (0, 0.025, 0.05, 0.075, and 0.1 mg/mL) were

CHAPTER 3 prepared in 80% MeOH from a stock solution of 25% (w/v) quercetin (Sigma) in 80% MeOH.

Then, 1 mL of each standard was mixed thoroughly with 3 mL of 80% MeOH, 0.2 mL of 10% aqueous dilution of AlCl3 (Sigma), 0.2 mL of 1 M potassium acetate (Sigma) and 5.6 mL of distilled water for 30 s and allowed to stand at 22°C for 30 min. The absorbance was measured at 415 nm using the same spectrophotometer and results were expressed as quercetin equivalent (mg QE/g of dry extract). Samples were prepared in MeOH (1 mg/mL) and measured following the same procedure as the preparation of standards. The TFC of samples was calculated using Equation 3.2.

C = (c × V)/ m (3.2)

where, C = TFC (mg QE/g dry extract), c = concentration of quercetin established from the calibration curve (mg/mL), V = volume of extract (mL), and m = weight of dry plant extract

(g).

3.2.5 HPLC analysis

All the C. nutans dried crude MeOH leaf extracts were re-dissolved in absolute HPLC grade

MeOH and filtered through 0.45µm PTFE filters before being analysed by HPLC.

Analysis was performed on a Dionex (USA) P680 solvent delivery system equipped with a

PDA100UV detector and operated using Chromeleon software (version 6.7). The separation was carried out on an Alltech Alltima (Grace, USA) C-18 column (250 × 4.6 mm, 5 µm) with a flow rate of 1 mL/min and an injection volume of 20 µL. Gradient elution was performed with solvent A: water and solvent B: HPLC grade acetonitrile (Ajax Finechem, Australia) for

40 min in the following order: 0-2 min (10% B), 14-24 min (75% B), 26-30 min (100% B),

CHAPTER 3 and 32-40 min (10% B). For preliminary phytochemical screening, the extracts were scanned at wavelengths ranging from 190 – 600 nm. Maximum absorption of the compounds in the extract was detected between 275 and 340 nm, where the greatest number of peaks was detected at 332 nm. Hence, this wavelength was selected for comparison of total peak area (≥

0.5 mAU*min) to estimate the amount of phytochemical compounds present.

3.2.6 Statistical analysis

Assays were performed in triplicate and data was expressed as the mean ± standard deviation

(SD). Analysis was performed using statistical software Minitab 17 (Minitab, 2013). The correlations were calculated according to Pearson’s correlation coefficient (r).

3.3 Results

3.3.1 Comparison of the TPC and TFC in leaf samples of C. nutans obtained from different locations within and between countries

Country and region of sampling had no significant effect on the TPC and TFC found in the crude MeOH leaf extracts of C. nutans collected from nine regions between and within three countries. Overall, there were variations in the total phenolic and flavonoid concentrations in the samples tested (Table 3.1). The TPC and TFC of each sample were calculated according to the equations obtained from the standard curves of GAE (Equation 3.3) and QE (Equation

3.4), respectively:

y = 10.094x + 0.0463 (r2 = 0.9957) (3.3)

y = 8.5958x + 0.0094 (r2 = 0.9992) (3.4)

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The TPC of all samples varied from 8.29 to 72.16 mg GAE/g dry extract, with a mean of

33.16 mg GAE/g dry extract. Higher variation was found between the Thai samples compared to the Malaysian samples, where the latter revealed consistent mid-range concentrations of TPC. Phenolic content variation within Vietnam could not be examined because only one sample was collected due to the rarity of native and cultivated C. nutans in the country. For the Malaysian samples, the highest TPC was detected in CE1 from Sandakan, which was almost twice that of the sample with the lowest TPC i.e. CE3 from Tawau. For the

Thai samples, CT5 from Chiang Dao had the highest TPC, which was nearly nine times greater than that of CT2 from Map Khae, which had the lowest levels.

The TFC of all samples ranged from 2.96 to 58.38 mg QE/g dry extract, with a mean of 17.71 mg QE/g dry extract. Like TPC, higher variation was also found between the Thai samples compared to the Malaysian samples. For the Malaysian samples, both the highest and the lowest TFC levels were detected in samples from the same region, Sandakan, where CE1 had levels approximately three times greater than that of CE2. For the Thai samples, CT5 from

Chiang Dao had TFC levels nearly 20 times greater than that of CT2 from Map Khae, which had the lowest amounts.

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Table 3.1. The TPC and TFC (mean mg GAE or QE/g dry extract ± SD, n = 3) of 11 C. nutans samples. Country Region Sample code TPC TFC

Malaysia Seremban CP2 45.04 ± 0.06 12.17 ± 4.52 Sandakan CE1 51.19 ± 0.24 34.13 ± 0.05 Sandakan CE2 31.54 ± 0.13 10.35 ± 0.00 Tawau CE3 27.96 ± 0.17 14.43 ± 0.01 Kota Kinabalu CE4 36.55 ± 0.24 18.24 ± 0.01 Thailand Map Khae CT1 10.52 ± 0.05 3.61 ± 0.03 Map Khae CT2 8.29 ± 0.22 2.96 ± 0.03 Salaya CT3 15.69 ± 0.08 8.11 ± 0.04 San Sai CT4 44.28 ± 0.19 23.45 ± 0.05 Chiang Dao CT5 72.16 ± 0.56 58.38 ± 0.19 Vietman Ho Chi Minh CV1 21.58 ± 0.12 8.96 ± 0.08 Mean 33.16 ± 19.29 17.71 ± 16.24

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3.3.2 Correlation between the colorimetric and HPLC methods to investigate the TPC and TFC in C. nutans leaves

Pearson’s correlation revealed that the colorimetric and HPLC methods were significantly (p

≤ 0.01) correlated and the trends among the samples were similar. Stronger correlation was observed between the TFC and the total HPLC peak areas (r = 0.914) than with the TPC (r =

0.893) (Figure 3.2). Detection at 332 nm in HPLC revealed that compounds in the C. nutans crude MeOH leaf extract eluted between 1.0 and 11.0 min, where major peaks were seen between 8.0 to 10.0 min (Figure 3.3). The mean total peak area of the 11 C. nutans samples was 92.49 ± 71.44 mAU*min. The maximum total peak area was 235.12 ± 17.81 mAU*min of CT5, almost 13 times that of minimum total peak area detected in CT2 (18.40 ± 0.34 mAU*min) (Figure 3.4), suggesting that there were higher concentrations and/or number of phytochemical compounds present in the former.

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Figure 3.2. Scatter plots showing correlations between HPLC peak area and the TPC (A) and the TFC (B) in C. nutans leaves. ** p ≤ 0.01.

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Figure 3.3. HPLC chromatograms of crude MeOH leaf extracts of C. nutans samples CE1 (A) and CE2 (B) detected at 332 nm.

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Figure 3.4. Total peak areas of 11 C. nutans samples from HPLC method detected at 332 nm. Data represent mean ± SD, n = 3.

3.4 Discussion

3.4.1 The TPC varies in C. nutans leaves collected from different locations

This is the first study to compare the TPC in C. nutans leaves collected from 11 different locations of Malaysia, Thailand and Vietnam, with diverse geographical and climatic conditions, to find the most suitable variety for commercialisation. Although these 11 C. nutans samples were very similar to each other genetically, with some showing identical genetic profiles (Fong et al., 2014), when grown under different environmental conditions even from the same region, they exhibited variable concentrations of TPC. The Thai samples revealed higher variations in the TPC (highest and lowest) than the Malaysian samples, although the mean TPC of the latter was higher than the former. In this study, the mean TPC of the 11 samples was higher (averagely ~ 2.4-fold) than that seen in previous studies (Yuann et al., 2012, Ghasemzadeh et al., 2014, Tiew et al., 2014, Wong et al., 2014, Mustapa et al.,

2015). This may be due to the different extraction methods used, for instance, different solvents and the duration of the extraction process.

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A notable observation is that, overall, C. nutans samples grown at higher elevations with cooler air temperatures exhibited higher TPC than those grown at lower elevations with warmer air temperatures. This was shown in CE1 and CE2, which were collected from the same region but with different elevations, where they possessed different concentrations of total phenolics. These results are in agreement with previous studies on other species that have shown interaction between phenolic content and environmental conditions, suggesting that environmental factors may play a role in the production of these compounds (Kishore et al., 2010, Guo et al., 2011, Dufoo-Hurtado et al., 2013).

The results of this study showing a significant positive correlation between elevation and the

TPC (Figure A2A, Appendix 4) are supported by one of the main theories, which suggest that elevation variations may be interpreted as adaptive responses or a defence mechanism of the plant to the increased level of harmful UVB radiation present at higher elevations

(Blumthaler et al., 1997). Meanwhile, a significant negative correlation was shown between the annual mean temperature and the TPC (Figure A2B). A number of studies have reported that photoperiod and low temperature could affect the biosynthesis of phenolics (Leyva et al.,

1995, Chalker-Scott, 1999, Azuma et al., 2012). Low temperature (cold stress) may enhance

PAL activity, an enzyme that is involved in the polyphenol biosynthesis, as well as increase phenolic content and their ester- and glycoside-bound forms in plants (Dufoo-Hurtado et al.,

2013). Moreover, Sanders (1982) reported that an increase in unsaturated fatty acids is generally associated with cooler climates, resulting in the production of antioxidants for a self-defence system against environmental stress. Phenolics also play an important role in the hydrogen peroxide scavenging system in plants, which besides phenolics, contains peroxidase, ascorbic acid and glutathione. This system functions less efficiently at low temperatures, and therefore, more of these compounds have to be produced to prevent damage to plants grown

CHAPTER 3 at lower temperatures (Bilger et al., 2007). Interestingly, a study by Albert et al. (2009) on

Arnica montana (Asteraceae) reported that lower air temperature was the main factor influencing altitudinal variation of phenolics in the plant rather than increased UVB radiation.

In contrast, Chen et al. (2013) described UV radiation as the primary factor that caused the variation in leaf phenolics across Chinese grasslands. The inconsistency in results suggests that further studies at the taxonomic level are needed to understand the main functional roles of phenolics involved in chemical defence mechanism of plants.

Low rainfall may also affect the production of phenolics, as seen in tartary buckwheat

(Suzuki et al., 2005) and carrots (Leja et al., 2013). Like any other environmental stresses, desiccation could also enhance the defence mechanism of plants, leading to an increase in the production of secondary metabolites. However, this phenomenon was not observed in the current study (Figure A2C).

3.4.2 The TFC varies in C. nutans leaves collected from different locations

This study is also the first to compare the TFC in C. nutans leaves collected from 11 different locations in the three countries, with different environmental conditions. This study has revealed that higher significant variations in the TFC among the Thai samples compared to the Malaysian samples extracted. It also showed a higher mean TFC of all the Thai samples compared to the Malaysian samples. In this study, the mean TFC of the 11 C. nutans samples was higher (averagely ~ 5-fold) than that seen previously (Ghasemzadeh et al., 2014, Tiew et al., 2014, Wong et al., 2014, Mustapa et al., 2015). This difference may be attributed to the different extraction method used as described earlier. Moreover, the variation in the TFC shown in this study is in accordance with that of Chelyn et al. (2014), where C. nutans leaves

CHAPTER 3 of different origins in Malaysia showed variation in flavonoids, particularly flavones C- glycosides, including shaftoside, orientin, isovitexin and vitexin.

The effects of elevation (Figure A2A) and annual mean temperature (Figure A2B) on the

TFC were found to be similar to that observed for the TPC, where samples from regions at higher elevations and with cooler air temperatures exhibited higher flavonoid levels. The results are in agreement with studies performed on maize seedlings (Christie et al., 1994),

Veronica chamaedrys (Plantaginaceae) (Nikolova and Gevrenova, 2006), red-wine grapes

(Mori et al., 2007) and ginkgo leaves (Wang et al., 2014), where higher elevations and lower air temperatures were shown to be beneficial to the biosynthesis and accumulation of flavonoids. However, studies have also reported the opposite effect, where altitudinal variation did not show a positive correlation with the TFC (Wang and Zheng, 2001, Kishore et al., 2010, Bradish et al., 2012). This suggests that the plants might have developed other adaptive traits other than the production of flavonoids to cope with environmental stresses or other environmental factors, besides altitude, might induce flavonoid synthesis in the plants.

A number of reports have discussed the effect of UV radiation on the metabolism of flavonoids in plants (Jaakola and Hohtola, 2010). Increased level of UVB radiation leading to enhanced activity of CSH, which is a key enzyme that involves in the initial step of flavonoid biosynthesis, has been well documented in a number of plant species (Jenkins, 2009). The light-absorbing flavonoids, flavonols and anthocyanins, in particular, act as a solar screen by accumulation of these compounds in epidermal cells to protect internal tissues of the leaves and the stems from the damaging UVB radiation (Jaakola and Hohtola, 2010). Meanwhile, low temperature exposure has also shown to increase the expression of PAL and CSH genes,

CHAPTER 3 stimulating the specific enzymes involved in the biosynthesis of flavonoids (Leyva et al.,

1995).

In addition, a significant positive correlation was found between the TPC and TFC (Figure

A2D), which has also been observed in studies on Cuphea spp. (Lythraceae) (Cardenas-

Sandoval et al., 2012) and hawthorn fruits (García-Mateos et al., 2013). However, some studies have shown that a high phenolic content is not always associated with a high flavonoid content (Spitaler et al., 2008, Lizcano et al., 2010), suggesting that other phenolic compounds may be present.

3.4.3 Relationship between the colorimetric and HPLC methods

The correlation between the two methods to investigate the TPC and TFC in C. nutans leaves was highly significant, where similar trends among all extracts were observed. However, the results from HPLC analysis cannot be directly compared with the results from Folin-

Ciocalteu or AlCl3 methods due to several reasons. In HPLC analysis, quantifications of phenolics and flavonoids using external standards were not performed and the total phytochemical content of each crude methanol leaf extract of C. nutans sample was based on the sum of all integrated peak areas at 332 nm, which was expressed as mAU*min. Therefore, the data was expressed differently from the data obtained from both colorimetric methods.

Besides, general screening of phytochemicals by HPLC in the present study revealed not only phenolics and flavonoids, but also a wide range of other compounds, which may contribute to the sum of the peak areas. It is suggested that future studies using standards are needed for comparison of the two methods based on a specific class of compounds.

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Several studies have shown significant correlation between the colorimetric and HPLC methods to determine phenolic compounds in extracts of Ipomoea batatas (Convolvulaceae)

(Truong et al., 2007), Aspalathus linearis (Fabaceae) and Cyclopia genistoides (Fabaceae)

(Joubert et al., 2008). Although, the correlation between the two methods was highly significant, the concentrations determined by the colorimetric method were greater than the

HPLC values. These differences were reported in several studies, where colorimetric results were ten times higher than that of HPLC in red wine (Burns et al., 2000), three to six times in champagnes (Chamkha et al., 2003), four to six times in peanuts (Talcott et al., 2005) and two to five times in sweet potato (Truong et al., 2007). The low values obtained from HPLC method can be attributed to the fact that not all the compounds were isolated, identified and quantified. In contrast, the colorimetric method can overestimate the total content, since it also reacts with hydroxyl groups in amino acids and sugars in addition to phenolic groups

(Singleton et al., 1999).

The most frequently used detection method for HPLC is UV spectrophotometry. Detection in

HPLC is typically based on the measurement of UV absorption, or visible absorption. No single wavelength is ideal to detect phenolics and especially all classes of flavonoids since they display absorbance maxima at distinctly different wavelengths (Marston and

Hostettmann, 2006). The range of wavelengths for the detection of these compounds is wide, ranging from 210 to 370 nm for phenolics (Brandsteterova and Ziakova-Caniova, 2005) and

210 to 520 nm for flavonoids (Marston and Hostettmann, 2006). Generally, the majority of phenolic acid and flavonoid derivatives demonstrate UV absorption maximum at 270-290 and 317-340 nm, respectively (Kassim et al., 2010, Caunii et al., 2012). In this study, UV spectrum of C. nutans methanolic leaf extract revealed absorption maxima at 275 and 340 nm.

There was also a strong correlation between the total HPLC peak area detected at 332 nm in

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CHAPTER 3 both the TPC and TFC, suggesting the compounds may be phenolics and flavonoid derivatives. Further studies using other analytical techniques, such as HPLC-MS or NMR, are required for the identification of these compounds in C. nutans leaves.

3.5 Conclusion

In conclusion, understanding the effect of environmental factors on the biosynthesis of secondary metabolites, which possess valuable medicinal benefits, is important in assisting us to obtain a higher medicinal content of constituents from the leaves of C. nutans. This study showed differences in the TPC and TFC among leaf samples collected from different regions and countries with varying environmental characteristics. Results demonstrated that the TPC and TFC in C. nutans leaves may be associated with growing conditions, where higher elevation and lower temperature are beneficial to the overall production of phenolics and flavonoids. This study presents essential insights into how collection sites and the growing environment may influence the TPC and TFC of C. nutans. Further studies are required to gain a better understanding on these interactions. In addition, according to the correlation results of this present study, colorimetric measurements of the crude MeOH leaf extracts of C. nutans can be used to predict their TPC and TFC to an acceptable accuracy as an alternative to HPLC analysis. The latter has the advantage over the colorimetric methods in that it provides more specific information about individual phytochemical compounds or groups at the same time.

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Effect of drying temperature and extraction solvent on the total phenolic and flavonoid content of Clinacanthus nutans (Burm. f.) Lindau (Acanthaceae) leaves

4.1 Introduction

Despite the progress in synthetic organic chemistry and modern biotechnology, medicinal plants are still an essential source of therapeutic drugs, especially in developing countries, where about 80% of the population still rely on herbal medicine as their primary form of healthcare (Ekor, 2013). Moreover, medicinal plants are used worldwide either as an individual treatment method or as part of a comprehensive treatment plan together with traditional methods of diagnosis and treatment (Tanko et al., 2005). However, the collection of wild plants has many disadvantages, such as decreasing availability, irregular supply and sometimes unreliable botanical identification. Therefore, there has been an increased interest in applying methods to cultivate popular medicinal plants. The possibility of cultivating these plants under preferable environmental conditions and most likely with genetically improved breeds enables the acquisition of plants, which are rich in their production of biologically active compounds (Palevitch, 1991). The concentration of active phytochemicals in a medicinal plant species is the most important aspect of medicinal plant phytochemistry since low levels of bioactive secondary metabolites hinder the therapeutic efficacy of the plant.

Besides environmental and genetic factors, postharvesting processes can affect the level of phytochemicals in a plant, and in so doing, change its nutritional value. In order to preserve the quality and the amount of valuable bioactive phytochemicals, it is important to consider the postharvest handling and processing of the plants, including storage, drying and extraction, which can affect the final phytochemical composition of a preparation.

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Water is essential to living organisms because it is required in many metabolic reactions. So, reducing the water content in freshly harvested medicinal plants is necessary to prevent enzymatic and microbial activities, which can negatively affect the shelf life and quality of the plants. There are many methods available for the removal of the water content from plants but generally, drying is performed using natural and mechanical means, which is based on a heat source and different forms of energy (e.g. solar, electromagnetic) (Cai et al., 2004).

Regardless, different drying methods can affect the composition of compounds present in plants. Previous studies have shown discrepancy in the amount of phytochemicals present in plant materials that were dried using different methods. Capecka et al. (2005) showed that air-drying in the shade resulted in a considerable increase of total phenolics in oregano and peppermint leaves, whereas no significant difference was seen between fresh and dried leaves of lemon balm. In addition, Periche et al. (2015) reported that stevia leaves dried with hot air at 180°C had the highest phenol, flavonoid and antioxidant levels compared to that of hot air- dried at 100°C, freeze-dried, shade-dried and fresh leaves. Meanwhile, Chan et al. (2009)

Pinela et al. (2011) found that freeze-drying was the most efficient method to preserve the quality of ginger leaves and flowers of Cytisus and Pterospartum spp. respectively.

Extraction is an important step not only for end users who would concoct their own herbal products, but also for the phytochemical analysis and biological activity of medicinal plants, because it is necessary to extract the desired components from the plant materials. There are various types of extraction methods for phytochemical preparation, such as solid-liquid, supercritical fluid, pressurised liquid and pressurised hot water extractions. The most commonly used method for isolating phytochemical compounds from plant materials is solid- liquid extraction (Boeing et al., 2014), where it involves the transport of solvent onto particles; desorption of compounds from plant matrices; solubilisation of solutes in the

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CHAPTER 4 solvent; and diffusion of solute molecules to the solvent (Tanko et al., 2005). The yield of phenolics may be affected by several parameters, which include extraction time, temperature, solvent-to-sample ratio, the number of repeat extractions of the sample and the type of solvent used. In addition, varying samples will have a different optimum recovery of phenolics and it depends on the type of plant and its active compounds. The selection of the extraction solvent system mainly relies on the specific polarity and chemical nature of the target solutes or phytochemicals and is based on the principle of “like dissolves like”.

Therefore, extraction of lipophilic (hydrophobic) compounds (e.g. alkanes, fatty acids, chlorophylls, sterols, some terpenoids, alkaloids and coumarins) uses non-polar solvents, such as hexane and dichloromethane. Medium polarity solvents, including ethyl acetate and chloroform, are used for extraction of compounds of intermediate polarity (e.g. some alkaloids and flavonoids), while the extraction of more polar or hydrophilic compounds (e.g. flavonoids, glycosides, tannins and some alkaloids) uses polar solvents like ethanol, methanol or water (Mandal et al., 2015). Different plant species and even their materials contain different classes or groups of phenolics, which are selectively soluble in different solvents.

Also, there is a possibility of interaction of phenolics with other plant components, such as carbohydrate and proteins, which may lead to the formation of complexes with different solubility (Naczk and Shahidi, 2006). Therefore, solvent polarity plays a key role in the extraction of phenolic compounds and no single standard extraction method is used to extract all plant phenolics. The most widely used solvents for extracting phenolic compounds are water, methanol, ethanol, acetone and their water mixtures, with or without acid (Naczk and

Shahidi, 2006, Rababah et al., 2010, Bunea et al., 2012, Michiels et al., 2012).

Due to the increasing interest in Clinacanthus nutans (Burm. f.) Lindau, the plant is often exported and sold in herbal markets as powdered or whole dried leaves. Extracts of C. nutans

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CHAPTER 4 can be prepared by a number of methods but the most common preparation by consumers is by steeping the dried leaves in boiled water and the infusion is then taken as a herbal tea.

Little information is available on the effect of postharvest processing, including drying treatment and solvent extraction, and on the quality and the amount of phytochemicals present in the leaves. Therefore, the present study aimed to investigate the effect of using different drying temperatures (40, 50, 60, 80 and 100°) and five different extraction solvents on the phytochemical content, especially the TPC and TFC, in C. nutans leaves using colorimetric and HPLC methods.

4.2 Materials and methods

4.2.1 Plant materials

Fresh leaves of C. nutans were collected from TKC Herbal Nursery, Seremban, Negeri

Sembilan, Malaysia and labelled as CP2 (Figure 4.1). Prior to sample treatment and extraction, all leaf pieces were thoroughly washed using cold tap water. All leaves, except those used for the postharvest drying study, were air dried in the shade for seven days at 22°C.

All dried leaves were stored as the whole leaf in air-tight bags in the darkness at 22°C until further analysis.

Figure 4.1. Collection site of CP2 from Peninsular Malaysia.

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4.2.2 Postharvest drying treatment

4.2.2.1 Drying procedure

Approximately 20 g of fresh leaves were weighed, placed on a flat tray and dried in a hot air convection oven (Model FM27MIX, Ariston, Italy) at temperatures of 40, 50, 60, 80, and

100°C, respectively. About 5 g of fresh leaves dried at 22°C were used as a control. During treatment, leaves were weighed every 15 or 30 min until a constant weight was obtained to ensure complete drying. Dried powdered C. nutans leaves of each post-drying treatment group were extracted following the procedure used to prepare the crude dichloromethane and

MeOH extracts as described in Section 4.2.3.1. However, only the MeOH extract was used for the TPC, TFC and HPLC analyses.

4.2.3 Postharvest crude extraction

4.2.3.1 Dichloromethane (DCM) and MeOH

One gram of dried powdered C. nutans leaves was extracted with 50 mL of a mixture of

DCM (Merck, Germany) and MeOH (Merck) in a ratio of 1:3 on an orbital shaker at a speed of 200 rpm at 22°C for seven days. The extracted solution was decanted, filtered with

Whatman No. 1 filter paper and concentrated under reduced pressure using a rotary evaporator (Buchi, Switzerland) to produce the crude DCM-MeOH dried extract, which was then weighed and used for further DCM and MeOH extractions.

The crude DCM-MeOH dried extract was first dissolved in DCM, filtered with glass wool and concentrated under reduced pressure to give the crude DCM dried extract. The remaining extract in the round bottom flask was dissolved in MeOH, filtered with glass wool and concentrated in vacuo to produce the crude MeOH dried extract. Both extracts were weighed and stored at -20˚C for the TPC, TFC and HPLC analyses.

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4.2.3.2 Ethanol (EtOH)

One gram of dried powdered C. nutans leaves was extracted with 50 mL of EtOH (Sigma,

USA) on an orbital shaker at a speed of 200 rpm at 22°C for 24 h. The extracted solution was decanted, filtered with Whatman No. 1 filter paper and concentrated in vacuo to produce the crude EtOH dried extract, which was then weighed and stored at -20˚C for the TPC, TFC and

HPLC analyses.

4.2.3.3 Cold aqueous (cH2O)

One gram of dried powdered C. nutans leaves was extracted with 50 mL of distilled water on an orbital shaker at a speed of 200 rpm at 22°C for 24 h. The extracted solution was decanted, filtered with Whatman No. 1 filter paper and freeze dried using a freeze dryer (Operon, Korea) at -60°C for 24 h to produce the crude cH2O dried extract, which was then weighed and stored at -20˚C for the TPC, TFC and HPLC analyses.

4.2.3.4 Hot aqueous (hH2O)

One gram of dried powdered C. nutans leaves was extracted with 50 mL of distilled water at

100°C for 5 min. The extracted solution was decanted, filtered with Whatman No. 1 filter paper and freeze dried using a freeze dryer (Operon) at -60°C for 24 h to produce the crude hH2O dried extract, which was then weighed and stored at -20˚C for the TPC, TFC and

HPLC analyses.

4.2.4 Determination of the TPC

The TPC was determined following the procedure as described in Section 3.2.3.

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4.2.5 Determination of the TFC

The TFC was determined following the procedure as described in Section 3.2.4.

4.2.6 HPLC analysis

HPLC analysis was performed following the procedure as described in Section 3.2.5.

4.2.7 Statistical analysis

Assays were performed in triplicate and data was expressed as the mean ± SD. Data was analysed using statistical software Minitab 17(Minitab, 2013). Polarity of solvents was ranked according to Snyder’s Polarity Index (Snyder, 1974) in increasing order: dichloromethane (3.4) < ethanol (5.2) < methanol (6.6) < water (9.0). The correlations were calculated based on the Pearson’s correlation coefficient (r) and significant difference was determined using ANOVA Fisher’s test at p ≤ 0.05 significance level.

4.3 Results

The TPC and TFC of all C. nutans samples treated with respective treatments were measured and calculated according to the equation obtained from the calibration curves of GAE

(Equation 4.1) and QE (Equation 4.2), respectively.

y = 10.094x + 0.0463 (r2 = 0.9957) (4.1)

y = 8.5958x + 0.0094 (r2 = 0.9992) (4.2)

4.3.1 Effect of drying temperature on the TPC and TFC in C. nutans leaves

Drying temperatures had a significant impact on the TPC and TFC found in the crude MeOH dried leaf extracts of C. nutans. The TPC levels decreased with low drying temperatures (40,

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50 and 60 °C) and increased with high drying temperatures (80 and 100 °C), compared to the air dried sample, which was used as the control. In contrast, the TFC showed a different trend, where it increased as drying temperature increased. However, leaves dried at 100 °C showed a decrease in the level of TFC (Table 4.1A).

Table 4.1. The TPC and TFC (mean mg GAE or QE/g dry extract ± SD, n = 3) in C. nutans leaves dried at different drying temperatures and extracted with different solvents. Samples that do not share a letter are significantly different at p ≤ 0.05 level Sample TPC TFC A. Temperature (°C): Control (air dried) 45.04 ± 0.06 12.17 ± 4.52 40 22.44 ± 0.03 14.02 ± 1.68 50 38.40 ± 0.04 18.23 ± 2.08 60 40.92 ± 0.05 24.79 ± 0.77 80 45.70 ± 0.03 27.72 ± 0.14 100 63.31 ± 0.03 25.21 ± 0.03 Mean 42.64 ± 13.20 20.36 ± 6.47 B. Solvent: Crude extract yield (% w/w) DCM 19.16 36.68 ± 0.04 8.48 ± 3.14 MeOH 7.87 45.04 ± 0.06 12.17 ± 4.52 EtOH 6.53 24.57 ± 0.07 7.09 ± 2.98 cH2O 23.27 48.08 ± 0.04 14.66 ± 1.71 hH2O 24.00 41.61 ± 0.06 12.93 ± 0.68 Mean 39.19 ± 9.21 11.07 ± 3.17

The TPC of all samples varied from 22.44 to 63.31 mg GAE/g dry extract, with a mean of

42.64 mg GAE/g dry extract. Leaves dried at 100°C had the highest TPC, which was almost triple that of the leaves dried at 40°C with the lowest TPC. The TPC of the treated samples decreased in the following order 100°C > 80°C > 60°C > 50°C > 40°C. Statistical analysis on the TPC data showed that all treated groups were significantly (p ≤ 0.05) different from the control as well as between them.

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Meanwhile, the TFC of all samples ranged from 27.72 to 12.17 mg QE/g dry extract, with a mean of 20.36 mg QE/g dry extract. Unlike TPC, leaves dried at 80°C exhibited the highest level of TFC, approximately twice that of air dried leaves, which had the lowest amounts.

The TFC of the samples decreased in the following order 80°C > 100°C > 60°C > 50°C >

40°C. ANOVA Fisher’s test revealed significant (p ≤ 0.05) difference between all treated groups and the control, except for 40°C.

In addition, correlation analysis showed significant (p ≤ 0.01) correlations between drying temperatures and both contents with TPC having a coefficient value (r) of 0.643 and TFC with 0.791 (Figure 4.2A). This suggests that drying temperatures have a considerable impact on the total phenolics in C. nutans leaves, particularly on flavonoids. However, TPC of this postharvest processing factor was not significantly correlated to TFC (r = 0.420) (Figure

4.2C).

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Figure 4.2. Scatter plots showing correlations between the contents (TPC and TFC) in C. nutans leaves and postharvest factors including drying temperatures (A) and extraction solvents (B) as well as between the two contents of each factor (C). ** p ≤ 0.01 and NS not significant.

4.3.2 Effect of extraction solvents on the TPC and TFC in C. nutans leaves.

Extraction solvents had significant effects on the TPC but not the TFC found in the five different crude dried leaf extracts of C. nutans. There were variations in the total phenolic concentrations in the samples tested. However, little variation was observed with the total flavonoid concentrations (Table 4.1B). The aqueous solvent, particularly hH2O (24%),

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CHAPTER 4 produced higher yield of dried crude extracts than organic solvents, with EtOH extracted the least amount (6.53%) (Table 4.1B).

The TPC of all extracts varied from 24.57 to 48.08 mg GAE/g dry extract, with a mean of

39.19 mg GAE/g dry extract. Among the five solvent extracts tested, the cH2O extract had the highest TPC, which was almost twice that of the extract with the lowest TPC i.e. EtOH. The

TPC of the extracts decreased in the following order cH2O > MeOH > hH2O > DCM > EtOH.

Statistical analysis on the TPC data showed significant (p ≤ 0.05) difference in concentrations between all extracts, including the cH2O and hH2O extracts, indicating that lower temperature of solvent (i.e. water) extracted significantly greater amounts of total phenolics from C. nutans leaves than higher temperature.

Meanwhile, the TFC of all extracts ranged from 14.66 to 7.09 mg QE/g dry extract, with a mean of 11.07 mg QE/g dry extract. Similar to TPC, the cH2O extract had the highest level, twice that of the EtOH extract, which had the lowest amounts. However, the TFC of the crude extracts decreased in a different order than TPC: cH2O > hH2O > MeOH > DCM >

EtOH. ANOVA test using Fisher method revealed that water temperature had no significant effect on the extraction of total flavonoids from the leaves, although the level of TFC was higher in the cH2O extract than that of the hH2O extract.

Moreover, correlation analysis revealed significant (p ≤ 0.01) correlations between the concentrations of both groups of secondary metabolites and extraction solvents according to

Snyder’s polarity index (Snyder, 1974) (Figure 4.2B). The TPC was also found to be significantly (p ≤ 0.01) correlated to the TFC with a coefficient value of 0.716 (Figure 4.2C).

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4.3.3 HPLC analysis and its correlation with the colorimetric methods

Detection at 332 nm in the HPLC analysis revealed that compounds in the crude extracts of all C. nutans leaf samples were eluted between 1.0 and 10.0 min, where major peaks were detected between 8.0 and 10.0 min (Figure 4.3).

Figure 4.3. HPLC chromatograms of C. nutans crude MeOH extract of the leaves dried at 40 and 80°C (A) and the crude EtOH and hH2O extracts (B) detected at 332 nm.

The mean total peak area of all C. nutans samples subjected to drying treatment at different temperatures was 134.40 ± 58.65 mAU*min. Leaves dried at 80°C had the maximum total peak area with 218.47 ± 1.73 mAU*min, approximately four times that of the minimum total peak area detected in the leaves dried at the lowest temperature i.e. 40°C (53.60 ± 0.39 mAU*min) (Figure 4.4A). According to ANOVA Fisher’s test, 60, 80 and 100°C treated groups had a significantly (p ≤ 0.05) higher content than the control, whereas no significant difference was observed for 40 and 50°C treated groups. The total HPLC peak areas of the

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CHAPTER 4 drying temperatures was found to be significantly correlated to the TPC (r = 0.565, p ≤ 0.05) and the TFC (r = 0.830, p ≤ 0.01) determined by the two colorimetric methods (Figure 4.5A).

The mean total peak area of the five crude dried extracts of C. nutans leaves detected at 332 nm was 80.42 ± 38.50 mAU*min. The maximum total peak area detected was 125.93 ± 1.82 mAU*min of the hH2O extract, almost four times that of the minimum total peak area found in the EtOH extract (31.97 ± 0.23 mAU*min) (Figure 4.4B), suggesting that there were higher concentrations and/or number of phytochemical compounds present in the former.

Statistical analysis on the HPLC data showed significant (p ≤ 0.05) differences between all the crude extracts. The total HPLC peak areas of extraction solvents was found to be significantly (p ≤ 0.01) correlated to the results determined by the Follin-Ciocalteu and AlCl3 methods (TPC: r = 0.745 and TFC: r = 0.685) (Figure 4.5B).

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Figure 4.4. Total peak areas of leaf samples dried at different temperatures (A) and extracted with five different solvents (B) obtained from HPLC with UV detection at 332 nm. Data represent mean ± SD, n = 3. * p ≤ 0.05, significantly different from control.

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Figure 4.5. Scatter plots showing correlations between HPLC peak area and the TPC and TFC in C. nutans leaves treated with different drying temperatures (A) and using different solvent extracts (B). ** p ≤ 0.01, * p ≤ 0.05.

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4.4 Discussion

4.4.1 Changes in the TPC and TFC in C. nutans leaves at different drying temperatures

To date, this is the first study to report the changes in the TPC and TFC concentrations in C. nutans leaves at different drying temperatures (40, 50, 60, 80 and 100°C), where the total contents of both phenolics and flavonoids increased with increasing temperatures, although the TFC slightly decreased at 100°C. In addition, results obtained from HPLC also showed a decrease in the amount of phytochemical compounds at 100°C. In most studies on the effects of heat treatment on the phenolic and flavonoid contents, the results are contradicting. Some studies reported an increase in the phenolic and flavonoid contents with increasing drying temperatures, including citrus peels (Chen et al., 2011), capsicum (Shotorbani et al., 2013), goldenberry (Lopez et al., 2013) and stevia leaves (Periche et al., 2015). On the contrary, others reported a decrease, such as that seen in sea buckthorn (Guan et al., 2005), motherwort and peppermint (Yi and Wetzstein, 2011) leaves or, in other cases, no significant changes, such as in the study of rosemary leaves (Yi and Wetzstein, 2011) and murta berries

(Rodriguez et al., 2014).

According to Mrad et al. (2012), a decrease in polyphenols during drying may be caused by the binding of these compounds with other compounds, such as proteins, or alterations in the chemical structure of polyphenols that cannot be extracted or determined by existing methods.

In this study, the reduction of TFC as well as the quantity of phytochemical compounds in general (analysed by HPLC) at the highest drying temperature i.e. 100°C may be caused by thermal degradation of heat-sensitive compounds, as suggested by Alonzo-Macias et al.

(2013). An increase in contents may be attributed by the formation of phenolic compounds at high temperatures e.g. 80 and 100°C, due to the availability of precursors of phenolic resulting in non-enzymatic interconversion between the molecules (Vega-Gálvez et al., 2012).

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Also, a decrease in peroxidase (PO) and polyphenoloxidase (PPO) enzymes activities, which are involved in phenolic oxidation, may lead to an increase in the level of phenolics (Ramani and Kant, 1989). Mizobutsi et al. (2010) reported that both PO and PPO enzymes were inactivated at high temperatures (i.e. 60 – 100°C).

4.4.2 The TPC and TFC vary among the different crude extracts of C. nutans leaves

The results of this study revealed that the most polar solvent (water) extracted the highest concentrations of both total phenolics and flavonoids. Among all the extracts, the cH2O contained the greatest amounts of both TPC and TFC, suggesting that cold water is the best solvent for the extraction of phenolics and flavonoids from C. nutans leaves.

This result is in agreement with previous studies on Ficus carica (Moraceae) (Konyahoglu et al., 2005), Leea indica (Vitaceae) (Reddy et al., 2012), Mentha spp. (Lamiaceae) (Barchan et al., 2014) and Psidium guajava (Myrtaceae) (Seo et al., 2014) leaves, suggesting that the type of solvent used, its polarity and the solubility of phytochemical compounds in the extraction solvents significantly influences the extraction efficiencies and recovery of the compounds.

These studies showed that the aqueous extract had greater content of phenolics and flavonoids than the organic extracts, including methanol, ethanol, ethyl acetate, hexane and dichloromethane. Nyirenda et al. (2012) reported that polar compounds like phenolics and flavonoids were more soluble in aqueous solvents than in organic solvents. Furthermore, several studies have also shown higher phenolic and flavonoid contents obtained using aqueous organic solvents compared to respective pure organic solvents, signifying that the amount of water in aqueous organic solvent mixtures has a significant impact on the extraction of phenolics than the solvent itself (Sultana et al., 2007, Sultana et al., 2009, Dent et al., 2013). However, the effect of mixed aqueous-organic solvent systems on the

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CHAPTER 4 phytochemical content in C. nutans leaves was not examined in this study and this should be investigated in future research. In contrast, other studies have also shown contradicting results, where a higher TPC and TFC content was found in the organic extracts compared to the aqueous extract (Aktumsek et al., 2013, Do et al., 2014). This suggests that different phenolic compounds possess a slightly different chemical nature e.g. polarity and hence, the selection of the extraction solvent system greatly depends on the compound of interest.

Aglycones of flavonoids are readily soluble in organic solvents but less soluble in water, while flavonoid glycosides are easily dissolved in hot water, methanol and other polar solvents, but they are insoluble in benzene and chloroform (Liu, 2011). The number of sugars attached to the aglycone is directly proportional to the solubility of the glycoside in water

(Liu, 2011). The higher amounts of total phenolics and flavonoids in C. nutans crude leaf solvent extracts of higher polarity (i.e. cH2O, hH2O and MeOH) implies that there were more polar compounds present in the leaves, such as C-glycosides, than less polar ones. This study also showed MeOH was more efficient than EtOH for the extraction of phenolics and flavonoids from C. nutans leaves, albeit their similar polarities. This suggests a lower solvation in EtOH, leading to a lower dissolution of molecules (Boeing et al., 2014).

A number of studies have shown that higher extraction temperatures increased the quantity of polyphenols extracted from plant materials, including Cymbopogon citratus (Poaceae) (Oboh et al., 2010), Camellia sinensis (Theaceae) (Venditti et al., 2010) and Thymus vulgaris

(Lamiaceae) (Vergara-Salinas et al., 2012). Ong et al. (2006) explained that increased temperatures can disrupt the physical chemistry interactions in the matrix, affecting the desorption process and will also decrease the viscosity of the liquid used, thus allowing better penetration of matrix particles and enhancing extraction. This can be seen from the results

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CHAPTER 4 obtained by HPLC, where leaves extracted with hot water had greater amounts of phytochemical compounds than those extracted with cold water. Interestingly, the results acquired by the colorimetric methods showed the opposite effect, where cold water had better extraction capacity than hot water in extracting total phenolics and flavonoids from C. nutans leaves. An explanation for the difference is that general screening of pytochemicals present in

C. nutans leaves by HPLC in this study revealed not only phenolics and flavonoids, but also a wide range of other compounds, which may have overlapping absorption bands, thus, contributing to the sum of the peak areas.

The results from the colorimetric methods are consistent with previous reports on other species, including white tea (C. sinensis) (Venditti et al., 2010) and Salvia officinalis

(Lamiaceae) (Dent et al., 2013), showing that lower extraction temperatures resulted in more polyphenols compared to the higher temperatures. This may be due to the decreasing polarity of water with increasing temperature, which then reaches values comparable to those of aqueous organic solvent mixtures (Vergara-Salinas et al., 2012), or due to possible degradation of compounds, caused by hydrolysis, internal redox and polymerization processes (Alonso-Salces et al., 2001).

4.5 Conclusion

In conclusion, temperature can greatly influence the levels of phytochemical compounds in C. nutans leaves during drying. Likewise, the type of extraction solvent can also significantly affect the amounts of phytochemicals extracted from the leaves. In this study, an increase in both the TPC and the TFC was observed with increasing drying temperatures, with leaves dried at 100 and 80°C having the highest content of total phenolics and flavonoids, respectively. This suggests that low temperature drying may not be a suitable postharvest

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CHAPTER 4 method for drying the leaves of C. nutans. In general, more polar solvents resulted in more total phenolics and flavonoids being exracted from the leaves. The greatest quantities of both the TPC and the TFC were observed in the crude cH2O extract, indicating that cold water is the best solvent for the extraction of phenolics and flavonoids in C. nutans. The results obtained can contribute to gaining further knowledge on how the quality and the potential health benefits of C. nutans may be maximised in terms of the preparation variation and different postharvest processes.

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In vitro cytotoxic effect of different crude leaf extracts of Clinacanthus nutans (Burm. f.)

Lindau (Acanthaceae) on selected human cancer cell lines and anticancer activity of the crude cold aqueous leaf extract against D24 melanoma cells

5.1 Introduction

Cancer is considered as a major cause of mortality worldwide. The International Agency for

Research on Cancer (IARC) estimates that approximately 14 million new cancer cases are diagnosed each year worldwide and this number is predicted to increase by about 70% over the next two decades (Forman and Ferlay, 2014). Melanoma and breast cancers are among the most common cancers diagnosed globally, where 230,000 and 1.7 million incidences were estimated in 2012, respectively (Ferlay et al., 2013).

Breast carcinoma is the most common cancer in women (Hussein and Komarova, 2011), where invasive ductal and lobular carcinoma are the most common types (Carton et al., 2007).

Although screening programs and the introduction of adjuvant therapies have successfully reduced death related to breast cancer by about 24% in several developed countries, metastatic cases are often fatal (Berry et al., 2005). Breast carcinoma is a hormone-driven disease strongly related to lifetime oestrogen exposure (Carton et al., 2007). Virtually, all breast carcinomas are adenocarcinomas derived from the glandular epithelial cells lining the terminal duct lobular unit, which spread locally within the breast, via lymphatics to axillary nodes, and haematogenously to distant sites such as the liver, lungs and brain. The disease can be removed by surgical excision of the lesion but when metastasis is involved, it is usually incurable (Carton et al., 2007).

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Skin cancers can arise from any of the many cell types that are present within the skin, and thus, they are very diverse. Melanoma has the highest mortality rate, but is less common than the non-melanoma skin cancers (Cifola et al., 2013). Cancer of the skin is characterised by an imbalance towards insufficient apoptosis or over cell proliferation and survival of the epidermis and in the case of melanoma, the melanocytes (Lippens et al., 2011). Besides UV radiation, other causative agents such as viruses, mutagens in food and chemicals as well as genetic susceptibility also can lead to skin cancer (Freedman and Nierodzik, 2007, Marks and

Hanson, 2010). Nearly 50% of melanomas harbour BRAF mutations, while the other half bear different types (Johnson and Sosman, 2014). Among the BRAF mutations, over 90% are at codon 600, and among these, more than 90% are a single nucleotide mutation resulting in substitution of glutamic acid for valine (BRAF V600E) (Ascierto et al., 2012). BRAF V600E has been implicated in different mechanisms of melanomagenesis, and predominantly the deregulated activation of the downstream MEK/ERK effectors (Maurer et al., 2011).

Melanoma can be surgically removed or treated by anti-angiogenic therapy, which curbs tumour growth at early stages but once it developed into metastatic cancer, it is basically incurable (Batus et al., 2013, Chinembiri et al., 2014).

Cytotoxic drugs are commonly used in chemotherapy, by killing cells that are rapidly growing and dividing. Unfortunately, most of these drugs can also affect normal dividing cells, resulting in unwanted side effects and complications, such as immunosuppression, nervous system breakdown and to a greater extent, organ damage (Frank, 2009). Hence, anticancer agents, which selectively target tumour cells, without affecting normal cells are ideal drugs to be used in cancer therapies, with the aim of maximising the benefits, while minimising the side effects (Chua, 2007).

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An abundance of natural sources with medicinal values exist worldwide, of which many have yet to be explored for possible therapeutic applications. More than 50% of commercially available drugs worldwide, of which over 70% of anticancer agents, derived from natural sources, including animals, plants, microbes and marine (Chinembiri et al., 2014). Plants are one of the most utilised and still the leading natural source for novel cancer drugs (Newman and Cragg, 2012). The use of plants as herbal medicine is the foundation of primary health care delivery for many people (estimated 80% of the world’s population), especially those living in under developed, rural and remote areas, because of their availability, affordability and culture (Ekor, 2013). Otherwise, in more developed countries, these herbal medicines can complement modern pharmaceutical medicines. Herbal medicines have a long history of use in health maintenance and in disease prevention and treatment, particularly for chronic diseases, including cancer (WHO, 2013) because they generally have lesser side effects than synthetic drugs (Fennell et al., 2004).

Generally, cell death could be classified into two major forms: apoptosis, which is programmed cell death, and necrosis, defined as non-programmed or uncontrolled cell death

(Messner et al., 2012). In the last decade, programmed cell death has expanded to include autophagy and necroptosis (programmed necrosis) (Su et al., 2015). The former is a catabolic process, where double-membraned vesicle is formed, encapsulating cytoplasm and organelles, which then fuses with lysosomes to degrade the contents in the vesicle (Hippert et al., 2006).

The latter, a newly discovered mechanism, is a form of regulated necrosis, which is activated by ligands of death receptors through the kinase activity of RIP1 (Zhou and Yuan, 2014). In vitro studies on effects of anticancer agents on tumour cells have allowed researchers to identify the different major modes of cell death (Boccarelli et al., 2011). Characterising the mode of cancer cell death induced by anticancer agents is important, not only to provide a

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CHAPTER 5 basis for understanding the underlying mechanisms involved, but also to help evaluate their impact on tumour development and treatment response in vivo.

Cancer cells are known to have defective apoptosis regulation, which leads to uncontrollable cell proliferation (Erb et al., 2005). Therefore, apoptosis is a major target of anticancer chemotherapeutics (Chinembiri et al., 2014). Besides, the induction of apoptosis has been described as a standard and more efficient strategy in anticancer therapy (Kelly and Strasser,

2011, Strasser et al., 2011) because it allows the removal of cancer cells without causing damage to the cellular environment e.g. inflammation (Russo et al., 2006). Cell apoptosis is an autonomous cell death process, which can be induced by a variety of drugs and physical as well as chemical factors (Liu et al., 2013). It has been suggested that a cell undergoing apoptosis shows multitude of characteristic morphological and biochemical features, which differ depending on the inducer, cell type and the specific time at which the process of programmed cell death is observed (Wlodkowic et al., 2011). The morphological changes in apoptotic cells are accompanied by several biochemical modifications, including the expression of cell surface markers such as phosphatidylserine externalisation, which translocates to the outer side of the plasma membrane during early apoptosis and can be detected by Annexin V staining (Arur et al., 2003), while the loss of all surface enzyme occurs afterwards (Piva et al., 2012). The biochemical aspects of different cell death modes can be detected by several methods, such as cytofluorimetry, immunofluorescence microscopy, immunoblotting and colourimetry assays (Piva et al., 2000, Boccarelli et al.,

2011, Piva et al., 2012), while typical morphological changes of cells as shown in Table 5.1 can be identified by microscopy techniques, including optical, scanning and transmission electron (Elmore, 2007).

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Table 5.1. Comparison of morphological features of apoptosis and necrosis. Apoptosis Necrosis Single cells or small clusters of cells Often contiguous cells Cell shrinkage and convolution Cell swelling Chromatin condensation peripherally at Chromatin condensation not common the nuclear membrane Intact plasma membrane Loss of plasma membrane integrity Cytoplasm retained in apoptotic bodies Cytoplasm leakage through permeable cell from blebbing and budding membrane Cytoplasmic vacuoles not common Cytoplasmic vacuoles are common

There has been an increased interest in Clinacanthus nutans (Burm. f.) Lindau (Acanthaceae), not only to treat the various ailments and illnesses (Section 1.3.7 and 1.3.8), but also various cancers and several studies have reported its anticancer activity against different human cell lines, including K-562, HeLa and Raji (Yong et al., 2013, Arullappan et al., 2014). However, there is little known on the cytotoxic effect of different crude extracts of C. nutans leaves on human melanoma and breast carcinoma cell lines as well as the possible mode of cell death elicited by such extracts. Therefore, the aims of this study were to i) examine the cytotoxic effect of the different crude C. nutans leaf extracts (DCM, EtOH, MeOH, cH2O and hH2O) on melanoma (D24 and MM418C1) and breast carcinoma (MCF7 and BT474) cell lines, ii) evaluate the cytotoxicity and selectivity of the most active extract (cH2O) against the most sensitive cell line (D24) in a dose- and time-dependent manner and iii) investigate the possible death mode of D24 cells induced by the cH2O extract using biochemical and microscopy techniques.

5.2 Materials and methods

5.2.1 Plant materials

The collection of C. nutans leaves from TKC Herbal Nursery, Seremban, Negeri Sembilan,

Malaysia was previously described in Section 4.2.1.

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5.2.2 Preparation of crude extracts

The preparation of different crude extracts of C. nutans leaves is described in Section 4.2.3.

The crude DCM and MeOH extracts were dissolved in DMSO (Sigma, USA) at a final concentration of < 0.1% (v/v), while the crude EtOH extract was re-dissolved in ethanol

(Sigma) at a final concentration of < 1% (v/v). Both the crude cH2O and hH2O extracts were re-dissolved in sterilised distilled water. Each extract from one biological replicate was used in three independent experiments (analytical replication).

5.2.3 Cell line and culture conditions

The C. nutans extracts were tested on four human cancer cell lines: D24 (wild type BRAF) and MM418C1 (BRAF V600E) melanoma cells; and MCF7 (wild type p53) and BT474

(temperature sentitive-p53 E285K) breast carcinoma cells; as well as human dermal fibroblasts (NHDF). The D24 and MM418C1 cells were cultured in RPMI 1640 medium containing L-glutamine (Gibco, Life Technologies, USA), whereas the MCF7, BT474 and

NHDF cells were cultured in high-glucose DMEM medium with pyruvate and L-glutamine

(Gibco). All culture media were supplemented with 10% foetal bovine serum (FBS) (Serana,

Melbourne, Australia) and 1% (v/v) penicillin/streptomycin (Gibco). For all the experiments, the cells were incubated for the indicated time under the indicated treatment at 37°C in a humidified atmosphere of 5% CO2.

5.2.4 Cytotoxicity assay

The cytotoxicity activity of all extracts was measured using the Cell Counting Kit-8 (CCK-8)

(Sigma), which is based on the conversion of a water-soluble tetrazolium salt, 2-(2-methoxy-

4-nitrophenyl)-3-(4-nitrophenyl)-5-(2,4-disulfophenyl)-2H-tetrazolium, monosodium salt

(WST-8), to a water-soluble formazan dye upon reduction by dehydrogenases in the presence

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CHAPTER 5 of an electron carrier. The cytotoxicity assay was performed according to the manufacturer’s protocol. Briefly, the cells were seeded in 96-well, flat bottomed plates (Greiner, Austria)

(100 µL/well) at a density of 5 × 103 cells/well for the D24, MM418C1 and NHDF cells, and

3 × 103 cells/well for the MCF7 and BT474 cells and incubated for 24 h. Subsequently, the cells were treated with the different extracts at 2 mg/mL and incubated for either 24 or 72 h.

Two types of control were used in this study i.e. untreated and vehicle. Untreated control cells were grown in the same supplemented medium lacking the C. nutans extracts, while vehicle control cells were treated with < 0.1% DMSO, < 1% ethanol or sterilised distilled water, but without C. nutans extracts. After 22.5 and 70.5 h, 10 µL of CCK-8 solution was added to each well and further incubated for 1.5 h at 37°C to reach a total of 24 and 72 h, respectively. Absorbance at 450 nm was measured using a microplate reader (CLARIOstar,

BMG Labtech, Germany) and data was obtained from the MARS data analysis software

V3.00 R2 (BMG Labtech). The percentage of viable cells was determined relative to the vehicle control cells using Equation 5.1. The vehicle controls were expressed as 100%.

Viable cell (%) = (absorbance of sample / absorbance of control) × 100 (5.1)

Only the two most active crude C. nutans extracts (cH2O and hH2O) against the most sensitive cell line (D24) as determined in this assay were used for EC50 determination

(Section 5.2.5).

5.2.5 Determination of half maximal effective concentration (EC50)

The D24 and NHDF cells were seeded in 96-well plates as described in Section 5.2.4. The cells were treated with different concentrations (0-2 mg/mL) of either the cH2O or hH2O extract and incubated at two time points, 24 and 72 h. At the end of each period, the

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CHAPTER 5 percentage of viable cells was determined using the CCK-8 method (Section 5.2.4). The viability of the untreated and vehicle (sterilised distilled water) control cells were not significantly different. Therefore, the percentage of viable cells was determined relative to the untreated control cells (100%). The EC50 values were determined from a non-linear regression model (curvefit) based on the sigmoidal dose-response curve (variable) and computed using GraphPad Prism version 6.05 (GraphPad Software, Inc., San Diego, USA)

(Lead et al., 2014). The extract with the lowest EC50 i.e. cH2O, was selected for subsequent experiments, including apoptosis evaluation by cytofluorimetry and microscopy analyses.

5.2.6 Observation of D24 cell morphology by phase contrast microscopy

Morphology of the untreated D24 cells and cells treated with 1 or 2 mg/mL of the cH2O extract for 24 and 72 h, in 96-well plates (Greiner), were observed using a Nikon Eclipse

TS100 (Japan) inverted microscope under 20× objective. Images were captured with the DS-

Fi 1 camera and DS-L2 control unit.

5.2.7 Evaluation of apoptosis by Muse cytofluorimetric analysis

Double staining with Annexin-7 and 7-AAD was performed using the Muse Annexin V/Dead

Cell Assay Kit (Merck Millipore, Germany). The untreated (control) and treated cells with 1 or 2 mg/mL of the cH2O extract for 24 and 72 h were washed twice with 1X PBS, dissociated with TrypLE reagent (Gibco), collected (both floating and adherent) in 1 mL microcentrifuge tubes, centrifuged at 800 × g for 5 min and resuspended in 100 µL of tissue culture medium.

Then, 100 µL of the fluorescent reagent was added to the cell suspension and incubated for

20 min at 22°C in the dark before being analysed for the detection of early and late apoptotic cells using a Muse Cell Analyzer (Merck Millipore). Based on the positivity of Annexin V, corresponding to phosphatidylserine externalisation in apoptotic cells and simultaneous

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CHAPTER 5 detection of dead cells, positive for the nuclear dye 7-AAD, the assay allows the differentiation of four populations in each sample by cytofluorimetric separation on a Muse

Cell Analyzer (Merck Millipore): i) viable (lower left quadrant: Annexin V−/7-AAD−), ii) early apoptotic (lower right quadrant: Annexin V+/7-AAD−), iii) late apoptotic/necrotic

(upper right quadrant: Annexin V+/7-AAD+) and iv) cell debris (upper left quadrant: Annexin

V−/7-AAD+) cells.

5.2.8 Detection of apoptosis and necrosis by Annexin V/PI double staining and confocal microscopy

The early and late apoptotic cells were detected using the Annexin V-FITC kit (Beckman

Coulter, USA), according to the manufacturer’s instructions with slight modifications. After treating the D24 cells with 1 or 2 mg/mL of the cH2O extract for 72 h, the cultures were stained with 1 µL of Annexin V-FITC (0.25 µg/mL) for 15 min, followed by 0.5 µL of PI

(0.125 µg/mL) for 5 min in the dark. The treated and untreated (control) cells were then observed using an inverted confocal microscope (Nikon Eclipse Ti-E A1, Japan) under 40× objective. The excitation wavelengths for Annexin V-FITC and PI used were 488 and 536 nm, respectively, while the emission wavelengths were 525 and 617 nm, respectively. Images were captured with a CCD camera and examined using the NIS-Elements Advanced

Research software (version 4.13). Based on the principles of this technique, the normal cells would not be stained by the two dyes (Annexin V-FITC−/PI−); the early apoptotic cells would only be dyed by Annexin V-FITC (Annexin V-FITC+/PI−); the late apoptotic cells would be positive in both Annexin V-FITC and PI staining (Annexin V-FITC+/PI+).

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5.2.9 Ultra structural analysis by TEM

The D24 melanoma cells were treated with the cH2O extract at 2 mg/mL for 72 h.

Subsequently, the treated and untreated (control) cells were washed twice with 1X PBS, dissociated with TrypLE reagent (Gibco), collected in 1 mL microcentrifuge tubes, centrifuged at 800 × g for 5 min and resuspended in 100 µL of 2.5% (v/v) glutaraldehyde with 2% (w/v) paraformaldehyde in 0.1 M sodium cacodylate buffer, pH 7.3 at 22°C for 30 min. Cells were then pelleted at 800 × g for 5 min and rinsed with 100 µL of 0.1 M sodium cacodylate buffer for 5 min and repeated three times. The cells were fixed in 100 µL of 1%

(w/v) osmium tetroxide with 1.5% (w/v) potassium ferrocyanide at 22°C for 1 h on an orbital shaker. After the period, the cells were centrifuged at 800 × g for 5 min and the supernatant was discarded. Then, the cells were washed by resuspending them in 100 µL of distilled water and were left on the orbital shaker for 10 min before centrifugation at 800 × g for 5 min.

This step was repeated twice. Dehydration was performed as follows: the cells were first resuspended with 100 µL of 50% (v/v) ethanol and left on an orbital shaker for 15 min before centrifugation at 800 × g for 5 min. The supernatant was discarded and followed by resuspension in 100 µL of 70% (v/v) ethanol for 15 min, 100 µL of 90% (v/v) ethanol for 15 min, 100 µL of 95% (v/v) ethanol for 15 min, 100 µL of 100% (v/v) ethanol for 30 min twice and finally 100 µL of 100% (v/v) acetone for 30 min twice. Infiltration was carried out twice with the acetone:Spurr’s resin (1:1) mixture on a rotator, first overnight and then for 2 h at

22°C. This followed by vacuum infiltration with fresh 100% Spurr’s resin for 2 h, repeated twice. Finally, the cells were polymerised at 70°C for 24 h. The cells were sectioned to a thickness of 1 µm using a UCT ultramicrotome (Leica Ultracut, Germany). The sections were observed at 80 kV with a JEOL JEM 1010 (Japan) transmission electron microscope and images were examined using the Gatan Microscopy Suite software version 2.3 (Gatan Inc.,

Pleasanton, USA).

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5.2.10 Statistical analysis

All cell experiments were performed in triplicate and the results presented as mean ± SD.

Data was analysed using statistical software Minitab 17 (Minitab, 2013). Significant difference was determined using ANOVA Fisher’s test at p ≤ 0.05 significance level.

Correlations were calculated according to Pearson’s correlation coefficient (r).

5.3 Results

5.3.1 Cytotoxic effect of the different crude C. nutans leaf extracts on the melanoma and breast carcinoma cells

The five different crude C. nutans leaf extracts were screened for their cytotoxic effect on the two melanoma (D24 and MM418C1) and two breast carcinoma (MCF7 and BT474) cell lines at the highest test concentration, which was 2 mg/mL for 24 and 72 h. The results obtained from WST-8 assay were shown in Figure 5.1. Among the four cancer cell lines, D24 was the most sensitive to these extracts. All the crude extracts exhibited cytotoxicity against the D24 cells, where the DCM, EtOH, cH2O and hH2O extracts showed significant (p ≤ 0.05) cell killing at 72 h (Figure 5.1A). The cH2O extract had the highest cytotoxicity against the D24 cells at 72 h, which caused 42.9% cell death, twice that of the MeOH extract with only 20.6% cell death. In the MM418C1 cells, both the cH2O and hH2O extracts showed cytotoxicity, particularly the hH2O extract (p ≤ 0.05) at 72 h, but not the DCM, EtOH and MeOH extracts

(Figure 5.1B). In fact, these crude extracts, especially the DCM and MeOH increased (p ≤

0.05) the percentage of viable cells at 72 h, about 1.5 times more compared to the control.

Although the cH2O and hH2O extracts showed cytotoxicity against the MCF7 cells at 24 h, where the hH2O extract caused significant (p ≤ 0.05) cell death, these extracts did not have any cytotoxic effect at 72 h (Figure 5.1C). On the contrary, the DCM extract displayed slight cytotoxicity against the MCF7 at longer treatment time. No cytotoxic effect on the MCF7

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CHAPTER 5 cells was observed for the MeOH and EtOH extracts. In the BT474 cells, all the crude extracts did not show any cytotoxicity, even though a slight cytotoxic effect was seen with the hH2O extract at 24 h (Figure 5.1D).

Figure 5.1. Percentage of viable D24 (A), MM418C1 (B), MCF7 (C) and BT474 (D) cells treated with the different crude C. nutans leaf extracts at 2 mg/mL for 24 and 72 h. Cell viability was determined by WST-8 assay. Data represents the mean ± SD from three independent experiments (n = 3). Samples that do not share a letter are significantly different at p ≤ 0.05 level according to the Fisher LSD method. * p ≤ 0.05, significantly different from vehicle controls (100%).

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5.3.2 Cytotoxicity of the crude cold and hot aqueous C. nutans leaf extracts against the

D24 melanoma and NHDF cells

The two most active crude C. nutans leaf extracts, i.e. cH2O and hH2O, along with the cell line with the highest sensitivity to treatments, i.e. D24 cells (Section 5.3.1) were used in further experiments, including the determination of the EC50. Following 24 h post-treatment with the extracts at concentrations ranging from 0.25 to 2 mg/mL, only 1 and 2 mg/mL showed significant (p ≤ 0.05) decrease in the percentage of viable cells compared to the untreated control (Figure 5.2A). The highest test concentration i.e. 2 mg/mL of the cH2O extract caused 45.7% cell death, relative to control after 24 h treatment, whereas the lowest test concentration (0.25 mg/mL) showed slight cytotoxicity with only 1.6% cell death.

Meanwhile, 2 and 0.25 mg/mL of the hH2O extract also showed cytotoxicity with 39.3 and

6.5% cell death, respectively.

The percentage of viable cells after 72 h treatment with both the extracts was significantly (p

≤ 0.05) reduced for all concentrations compared to the untreated control. Treatment with 2 mg/mL of the cH2O and hH2O extracts for 72 h demonstrated cytotoxicity with 57.6 and 48.6% cell death, respectively. However, the cells treated with the lowest test concentration (0.25 mg/mL) of the cH2O and hH2O extracts only showed 15.6 and 12.7% cell death, respectively.

Statistical analysis revealed no significant difference between treatment times but there was a significant (p ≤ 0.05) difference in cytotoxicity between the two aqueous extracts at 2 mg/mL for 72 h. All the extracts had EC50 values > 2 mg/mL, except the cH2O extract, which had a

72 h EC50 value of 1.63 mg/mL.

In addition, low cytotoxicity of both the cH2O and hH2O extracts against the normal cell line,

NHDF was observed. Treatment of the NHDF cells with 2 mg/mL of the cH2O and hH2O

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Figure 5.2. Cytotoxic effect of the crude cH2O and hH2O extracts on the D24 melanoma (A) and the NHDF cells (B). Data represents mean ± SD from three independent experiments (n = 3). * p ≤ 0.05, significantly different from untreated control. # p ≤ 0.05, significantly different between treatment times for the same extract concentration. † p ≤ 0.05, significantly different between extracts of the same time-point.

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5.3.3 Morphological changes in the D24 melanoma cells treated with the crude old aqueous C. nutans leaf extract

The untreated (control) D24 melanoma cells were adherent and exhibited a smooth surface and elongated structure. These cells also showed a finely granulated cytoplasm (Figure 5.3A

& D). In contrast, the cells treated with the cH2O extract displayed cell shrinkage with irregular and rough form and the cell number were lower than that of controls (Figure 5.3B, C,

E & F). More noticeable morphological changes were observed when the cells were exposed for longer treatment times and at higher concentrations, which corresponded to the results of the cytotoxicity assay.

Figure 5.3. Morphological changes (arrows) in the D24 melanoma cells treated with the crude cH2O extract. Phase contrast images (20×) of cells after 24 h exposure to no treatment (A), extract at 1 mg/mL (B) and 2 mg/mL (C); 72 h exposure to no treatment (D), extract at 1 mg/mL (E) and 2 mg/mL (F).

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5.3.4 Induction of mode of cell death in the treated D24 melanoma cells by the crude cold aqueous C. nutans leaf extract

As seen earlier (Section 5.3.2), the cH2O extract had a lower 72 h EC50 value, indicating higher cytotoxicity against the D24 cells than the hH2O extract. As a result, the cH2O extract was chosen for cytofluorimetric analysis. Treatment of the D24 cells with the extract at 1 mg/mL for either 24 or 72 h did not induce high affinity for both Annexin V and 7-AAD, indicating that most cells (> 50%) were viable (bottom left quadrant) (Figure 5.4A & B1). On the other hand, treatment with 2 mg/mL of the extract for 24 or 72 h increased the number of cells that were positive to both Annexin V and 7-AAD (upper right quadrant). This was also shown in Figure 5.4B2. When the cells were treated with 2 mg/mL of the extract for 24 or 72 h, there was a significant (p ≤ 0.05) increase in the percentage of cells that were undergoing or had undergone apoptosis (more on the latter) compared to the controls. This indicated that a high concentration of the cH2O extract was more likely to induce apoptosis in the D24 cells.

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Figure 5.4. Effect of the crude cH2O extract on the D24 melanoma cells. Representative apoptosis profile plots of untreated and treated cells after 24 or 72 h exposure to 1 or 2 mg/mL of the extract (A). The percentage of viable (B1), early apoptotic and late apoptotic/necrotic (B2) cell populations of untreated and treated groups. Data represents mean ± SD from three independent experiments (n = 3). * p ≤ 0.05, significantly different from control.

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Confocal microscopy was used to provide a qualitative identification of both apoptotic and necrotic deaths of the D24 cells treated for 72 h with 1 and 2 mg/mL of the cH2O extract

(Figure 5.5). Thr control (untreated) cells (Figure 5.5A-D) did not stain following the addition of both Annexin V and PI, which showed that they were viable. When the cells were treated with 1 or 2 mg/mL of the extract, they were shown to be positive to both Annexin V and PI. The cells treated with 2 mg/mL of the extract had a higher level of these fluorescent stains than compared to those treated with 1 mg/mL, which suggests that the modes of D24 cell death were dose-dependent. At lower test concentration (1 mg/mL), most of the cells were viable and hence, they were negative to both Annexin V and PI stains (Figure 5.5E-H).

However, several cells were positive to both stains, which appeared green and red, signifying the occurrence of late apoptosis/necrocis (Figure 5.5H). On the contrary, most of the cells treated with the higher test concentration (2 mg/mL) were positive to Annexin V and negative to PI, which appeared green (Figure 5.5J & L), indicating the presence of early apoptotic cells, and there were also a few that were negative to Annexin V and positive to PI, which appeared red (Figure 5.5K). The results obtained from confocal microscopy were similar to that obtained from the cytofluorimetric analysis.

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Figure 5.5. Confocal images (40 ×) of the untreated cells with no uptake of any stains (A-D) and the induction of early and late apoptosis/necrosis in the D24 cells after treatment with 1 mg/mL (E-H) and 2 mg/mL (I-L) of the crude cH2O extract for 72 h. Continuous arrows indicate early apoptotic cells and dotted arrows indicate late apoptotic/necrotic cells.

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5.3.5 Ultrastructural changes in the D24 melanoma cells treated with the crude cold aqueous C. nutans leaf extract

As the cH2O extract has shown to induce cell death in the treated D24 cells (Section 5.3.4), structural changes in the cell were further analysed electron microscopically to detect more details. In the D24 cells, clear morphological changes were observed between the untreated control and cells treated with the extract (2 mg/mL) for 72 h (Figure 5.6). The untreated cells displayed normal cell characteristics, such as microvilli, intact plasma membrane and nucleus with evenly distributed chromatin (Figure 5.6A). In contrast, in the treated cells, there were cell shrinkage, loss of microvilli, marked peripheral chromatin condensation at the nuclear membrane (Figure 5.6B & C), segmented/lobulated nucleus and irregular plasma membrane with extensive blebbing (Figure 5.6C & D). Furthermore, vacuoles were also observed in these cells (Figure 5.6C).

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MV Cr

Nu NM

PM NM

PM PM

NM * Nu NM Cr * Nu *

Figure 5.6. Transmission electron micrographs of the control (untreated) D24 cell (A) and the cells treated with the crude cH2O extract (2 mg/mL) for 72 h (B-D). Distinct morphological changes, including plasma membrane alteration (white arrow), chromatin condensation (white arrowheads), blebbing (black arrow), segmented/lobulated nucleus (black arrowheads) and formation of vacuoles (asterisks) were observed in the treated cells. Cr: chromatin, MV: microvilli, NM: nuclear membrane, Nu: nucleus, PM: plasma membrane, Va: vacuole.

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5.4 Discussion

5.4.1 The crude C. nutans leaf extracts exhibit different effects on the viability of melanoma and breast carcinoma cells

This study is the first to evaluate the effect of the different crude leaf extracts of C. nutans extracted with solvents (DCM, MeOH, EtOH, cH2O and hH2O) of varying polarity on the melanoma: D24 (wild type BRAF) and MM418C1 (BRAF V600E); and breast carcinoma:

MCF7 (wild type TP53) and BT474 (ts-TP53 E285K) cell lines. The melanoma D24 cells were the most sensitive to the cH2O and hH2O extracts. The extracts were shown to be cytotoxic to all the cell lines, with an initial loss of cell viability at 24 h in the MCF7 and

BT474 cells after which, they recovered. This suggests that stimulation of some proliferative effects in surviving cells occurred, or the cells eventually adapted to the treatment. Although most tumour cells initially respond to treatments, after an unpredictable period, they developed resistance to treatments through several mechanisms, including alteration of drug metabolism, derangement of intracellular signalling pathways, crosstalk between different membrane receptors, modification of apoptotic signalling and interference with cell replication (Fodale et al., 2011). Additionally, the higher responsiveness of the wild type

BRAF melanoma cell line (D24) to treatments compared to that of BRAF V600E-mutant

(MM418C1) suggests that the extracts affect the MAPK/ERK signalling pathway differently

(Ascierto et al., 2012). The DCM, EtOH and MeOH extracts of C. nutans leaves in general, showed little or no cytotoxicity against the tested cancer cell lines. In fact, these extracts enhanced the proliferation of the MM418C1, MCF7 and BT474 cell cultures at 72 h. It is known that medicinal plants are beneficial to human health, either as alternative treatments or preventive measures for a number of diseases, such as cancers (Sofowora et al., 2013).

However, it has been reported that some plant extracts have harmful effects on certain cancer cell lines by stimulating cell proliferation (Hasan and Al Sorkhy, 2014). For example,

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CHAPTER 5 ecdysterone, saffcomin A, psoralen and isopsoralen, isolated from Chinese herbs, were found to significantly increase proliferation of MCF7 cells, by enhancing the transition from G1 to

S phase (Niu et al., 2007). Danggui Buxue Tang (DBT), a Chinese herbal decoction containing Radix Astragali and Radix Angelicae sinensis (Apiaceae), was shown to stimulate the proliferation and differentiation of MG-63 human osteosarcoma cells (Choi et al., 2011).

The results of this study are in agreement with other research on different plant species, showing higher cytotoxicity effect of aqueous extracts than that of alcoholic extracts. For instance, studies by Madhulika and Ajitkumar (2010) and Moghe et al. (2011) on the whole plant of Cassia occidentalis (Fabaceae) and leaves of Bryonia laciniosa (Cucurbitaceae) respectively, reported that the cytotoxicity of aqueous extracts against several human cancer cell lines, including colon (HCT-15, SW-620, COLO 205), ovary (OVCAR-5), prostate (PC-

3), lung (HOP-62), MCF7 and cervix (SiHa), was higher than the corresponding alcoholic extracts. Uddin et al. (2011) also observed that aqueous leaf extracts from Bruguiera gymnorrhiza (Rhizophoraceae), Hibiscus tiliaceus (Malvaceae) and Limnophila indica

(Plantaginaceae) were more cytotoxic than methanol extracts when added to human gastric

(AGS), colon (HT-29) and breast (MDA-MB-435S) cancer cells. Similarly, aqueous Pouteria torta (Sapotaceae) leaf extracts were more effective when used at lower concentrations, with the most significant activity against human oral squamous carcinoma (OSCC-3) and MCF7 cells than treated with the corresponding hexane and ethanol extracts (Elias et al., 2013).

Saetung et al. (2005) found no cytotoxic activity in the aqueous extracts of Bridelia ovata

(Euphorbiaceae) leaves on lung adenocarcinoma (CORL-23) and PC3 cells. In addition,

Yong et al. (2013) showed that the chloroform extracts of C. nutans leaves were more cytotoxic against HepG2, NCI-23, gastric (SNU-1), colon adenocarcinoma (LS-174T), HeLa,

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K-562, and Raji cell lines than were the corresponding methanol and aqueous extracts. The extraction procedure (e.g. extraction time), treatment and handling of the crude extracts as well as the sampling location, growth/environmental conditions and harvest time of C. nutans leaves may affect the phytochemical components. Therefore, this may contribute to the observed differences in cytotoxic activity between this and the previous study.

Furthermore, the cytotoxicity of the cH2O and hH2O extracts, which were the most active against all cancer cell lines, may relate to the phytochemical composition of these extracts.

The phenolic, flavonoid and phytochemical (in general) levels were higher in the aqueous extracts, although the bioactive compounds that may be present are yet to be identified.

However, although the MeOH extract of the leaves had high concentrations of polyphenols, it was not very cytotoxic against the cell lines tested. Interestingly, the EtOH extract, which had the lowest concentrations of phytochemicals, showed moderate activity, for instance against the D24 cells. The extraction of bioactive phytochemical compounds from plant materials is significantly influenced by the solubility of each specific structure in the selected solvent

(Naczk and Shahidi, 2006). This suggests that each extract may consist of a unique set of compounds. Moreover, crude extracts are composed of several substances, which have not been purified and fully characterised. Thus, there is a possibility of molecules with opposing or synergistic effects found in a particular extract and not the others, which may contribute to the discrepancies in activity (Elias et al., 2013).

5.4.2 The crude cold and hot aqueous C. nutans leaf extracts exhibit selective cytotoxicity against the D24 melanoma cells

The cH2O and hH2O extracts of C. nutans leaves were shown to be cytotoxic to the D24 melanoma cells, but not to the NHDF cells. This suggests that the extracts contain

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CHAPTER 5 phytochemicals, which are selective against tumour cells but not normal cells. Results of this study are in accordance with earlier investigations of C. nutans (Yong et al., 2013,

Arullappan et al., 2014), which were shown to have cytotoxic effect against tumour cells. The cytotoxicity of the extracts might be due to the bioactive polyphenols present in the leaves, which at the moment, are still unknown and yet to be determined.

Many bioactive polyphenols are found in the leaves and leaf-derived products are important dietary sources of such compounds (Lavola et al., 2012). Epidemiological studies and associated meta-analyses strongly suggest that long term consumption of diets rich in plant polyphenols offer protection against the development of cancers, cardiovascular diseases, diabetes, osteoporosis and neurodegenerative diseases (Pandey and Rizvi, 2009). Effects of polyphenols on human cancer cell lines, including those from the mouth, stomach, duodenum, colon, liver, lung, breast and skin, are usually protective and they reduce the number of tumours or their growth through different mechanisms of action (Johnson et al., 1994, Yang et al., 2001). Reported polyphenols, which showed anticancer activity, were mostly flavonoids, such as catechin, apigenin, luteolin, kaempherol, myricetin, quercetin, rutin, orientin, vitexin, isoflavones and cyanidin as well as other phenolic compounds, including ellagic acid, resveratrol and curcumin (Pandey and Rizvi, 2009, Zhou et al., 2009, Lee et al.,

2012, Batra and Sharma, 2013, Guo et al., 2014).

5.4.3 The crude cold aqueous C. nutans leaf extract induces apoptosis in the treated D24 melanoma cells

This study is the first to examine the mode of D24 melanoma cell death that was induced by the cH2O extract in vitro. Cytofluorimetric analysis using Annexin V and 7-AAD staining showed that there was a significant increase in early and late apoptosis/necrosis as well as a

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CHAPTER 5 significant decrease in the number of viable cells when the cells were treated with 2 mg/mL extract when compared to the untreated controls following treatment. Visual assessment of the Annexin V/PI double-stained D24 melanoma cells by confocal microscopy showed early apoptosis in the cells 72 h post-treatment with 2 mg/mL of the extract. Late apoptotic/necrotic cells were also observed in these treated cultures. The results suggest that the cH2O extract induces apoptotic cell death in the D24 cells in dose- and time-dependent manner.

In order to further confirm the occurrence of various apoptotic and necrotic features of the

D24 cells induced by the cH2O extract, these cells were observed under TEM. Electron micrographs of the treated D24 cells revealed typical morphological features of apoptosis, such as marked peripheral chromatin condensation at the nuclear membrane, segmented/lobulated nucleus and irregular plasma membrane with extensive blebbing, These suggest that the extract induced apoptotic cell death. Kerr et al. (1972) explained that with cell shrinkage, the earliest recognised morphological changes during apoptosis are condensation, segregation and margination of chromatin at the nuclear membrane, which is the main characteristic feature of the process. Progression of condensation is accompanied by convolution of the nuclear and cell outlines followed by breaking up of the nucleus into discrete fragments and by budding of the cell as a whole to produce membrane-bound apoptotic bodies, consisting of cytoplasm with tightly packed organelles with or without nuclear fragments (Kerr et al., 1972, Elmore, 2007). The bioactive compound(s) present in this extract still remain to be identified. However, the presence of vacuoles in the treated cells suggests that some cells may have died by necrosis (Tan et al., 2005, Elmore, 2007). The mechanism for the apoptotic and necrotic cell deaths observed in the treated D24 cells still remain to be elucidated. A number of studies on other plant species have also reported

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CHAPTER 5 combination of cell death modes induced by crude leaf extracts, such as that shown with methanol extract of Pereskia bleo (Cactaceae) on breast carcinoma (T47-D) cells (Tan et al.,

2005) and hot aqueous extract of Cratoxylum formosum (Hypericaceae) on HepG2 cells

(Issara-Amphorn and T-Thienprasert, 2014).

5.5 Conclusion

A number of notable conclusions can be drawn from this study on the cytotoxic effects of the different crude extracts of C. nutans leaves against the melanoma (D24 and MM418C1) and breast carcinoma (MCF7 and BT474) cell lines as well as the anticancer activity of the crude cH2O extract against the D24 cells. First, results revealed variation in the effect of the different extracts at 2 mg/mL on the cell viability of the four cell lines. This indicates that the solubility of bioactive compounds in the extracts is significantly influenced by the extraction method. The D24 cell line was the most sensitive to treatment and the cH2O and hH2O extracts were the most active against all cancer cell lines, although they showed cytotoxicity against both the breast carcinoma cell lines only at 24 h after which, the cells recovered.

Second, the aqueous C. nutans leaf extracts, particularly the cH2O extract, were more cytotoxic against the cancer cells, i.e. D24 melanoma, than towards the non-cancerous NHDF cells. This suggests that these extracts may contain anticancer compounds. Third, based on the biochemical and morphological changes detected by cytofluorimetry and microscopy approaches, respectively, the cH2O extract predominantly induced the D24 cells to undergo apoptosis in a dose- and time-dependent manner. Moreover, necrosis was also detected in the treated cells, probably due to the combination of different bioactive compounds present in the crude cH2O extract. These observations suggest that the crude cH2O extract could be used as an alternate adjunctive regimen for cancer prevention or treatment.

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However, further research work is needed to isolate and identify the structure(s) of the active compound(s) or component(s) in the crude cH2O extract of C. nutans, as well as to determine their effectiveness as anticancer agents. In addition, further studies to understand the underlying mechanism in cell death, especially the cellular signalling pathways involved and in vivo testing of the observed anticancer activity are essential to explore the full potential of

C. nutans for therapeutic opportunities.

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Comparative cytotoxicity of Clinacanthus nutans (Burm. f.) Lindau (Acanthaceae) leaves obtained from different locations and anticancer activity of the crude methanol leaf extract against D24 human melanoma cells

6.1 Introduction

Cancer remains the leading cause of morbidity and mortality worldwide, with an estimated 14 million new cases and 8.2 million cancer related deaths in 2012. The number of new cases is predicted to rise by ~ 70% over the next 20 years (Forman and Ferlay, 2014). Melanoma is a type of skin cancer, characterised as a neoplastic disorder of the epidermal pigment- producing cells known as melanocytes (Uong and Zon, 2010). Melanoma is the least common but most fatal form of skin cancer and it has increased two-fold in the past two decades (Cifola et al., 2013). It is one of the most common cancers diagnosed in men and women worldwide, where 232,130 incidences were estimated in 2012 (Ferlay et al., 2013) and is the fourth most commonly diagnosed cancer in Australia, with more than 11,000 cases reported in 2010 (Australian Institute of Health and Welfare, 2014). About 50% of melanomas bear mutations in the BRAF gene, mainly at codon 600, resulting in constitutive activation of the MAPK signalling pathway, which plays a key role in the regulation of gene expression, cellular growth and survival (Jang and Atkins, 2013). Other melanomas harbour mutations in different genes, such as CDKN2A, PTEN and NRAS (Stark and Hayward, 2007).

Although melanoma can be surgically treated at early stages, it is an aggressive tumour with advanced disease defined by widespread metastatic lesions and the tumour has been reported to be resistant to most forms of cancer treatment (Batus et al., 2013). Therefore, it has become an increasingly critical public health issue, not just for Australia, but also globally and novel treatment options have become an important requirement.

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Besides surgery, radiation and systemic therapies have a role in the treatment of metastatic melanoma, cytotoxic chemotherapeutic agents with satisfactory antitumour efficacy, which include alkylating agents, platinum complexes and microtubular toxins, have also been used either alone or in combination for the treatment of the disease for more than three decades

(Bhatia et al., 2009). According to the National Cancer Institute, the currently approved melanoma drugs, which act on different cellular pathways include dabrafenib, dacarbazine, pembrolizumab and ipilimumab (NCI, 2015). Nonetheless, the main problems with chemotherapeutic agents are severe adverse effects and multi-drug resistance through drug efflux systems, amplifications of drug targets or changes in drug kinetics (Iyer et al., 2013,

Kunjachan et al., 2013, Markman et al., 2013). Although various strategies, such as the use of nanoparticles, liposomes and micellar as drug delivery vehicles, have been reported to have successfully overcame drug resistance (Markman et al., 2013), side effects caused by the high toxicity of chemotherapy drugs still remain a major dilemma that is unacceptable to many cancer patients (Chinembiri et al., 2014). Therefore, there is a need for the breakthrough of more effective and less toxic therapeutic and preventive strategies.

Over the years, there has been an increasing interest in the areas of natural products for novel and bioactive molecules for cancer drug discovery due to their general availability, safety and low toxicity, which may cause lesser side effects (Pratheeshkumar et al., 2012). Pytochemical compounds from roots, bulbs, barks, stems, flowers and leaves have been shown to have cancer protective effects against carcinogens (Pratheeshkumar et al., 2012) through the detoxification and enhanced excretion of carcinogens (Johnson, 2007). Besides, these compounds can be potential sources of anticancer agents or lead compounds for new drug synthesis (Chinembiri et al., 2014). Several compounds from various natural sources

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(summarised in Table 6.1) have been found to possess anticancer activity, specifically on

melanoma (Chinembiri et al., 2014).

Table 6.1. Melanoma therapeutic compounds obtained from natural sources (dietary components, phytochemicals and crude extracts). Apoptosis promoter Antiproliferative Antimetastatic Quercetin Kaempferol Amentoflavone Kaempferol Epigallocatechin gallate Hinokiflavone Epigallocatechin gallate Apigenin β-carotene Apigenin Vitamin A, C, D and E Fucoxanthin β-carotene Emodin from Aloe Vitamin A and C Fucoxanthin Eupatilin from Artemisia Resveratrol Vitamin C Extracts: Sulforaphane Resveratrol Ganoderma lucidum Carnosol and ursolic acid from Curcumin Coriolus versicolor Rosmarinus officinalis Sulforaphane Hypericum perforatum Extracts: Withaferin A from Withania Melaleuca alternifolia Withania somnifera somnifera Calendula officinalis Viscum album Eupatilin from Artemisia spp. Alpinia oxyphylla Calendula officinalis Galangin from Alpinia officinarum Alpinia galangal Extracts: Ganoderma lucidum Coriolus versicolor Melaleuca alternifolia Zingiber officinale

As most cancer cells have defective apoptosis mechanisms, which result in uncontrollable

cell proliferation, this pathway is a key target of chemotherapeutics (Chinembiri et al., 2014).

Apoptosis (programmed) is another main form cell death, besides necrosis (non-

programmed), where it is involved in tissue homeostasis and tumorigensis (Kerr et al., 1972).

The process is generally characterised morphologically by chromatin condensation and

formation of apoptotic bodies, which can be identified using microscopy techniques, and

biochemically by DNA fragmentation into oligonucleosome-sized fragments and

phosphatidylserine externalisation (Wyllie, 1997, Arur et al., 2003, Elmore, 2007), which can

be detected using cytofluorimetry, immunofluorescence microscopy, immunoblotting and

separation techniques (Boccarelli et al., 2011).

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Melanoma cells, like most of the cancer cells, can undergo apoptosis, where the molecular components of this cell death consist of positive (apoptotic) and negative (anti-apoptotic) regulators (Elmore, 2007). The former include molecules such as p53, Bid, Noxa, PUMA,

Bax, TNF, TRAIL, Fas/FasL, PITSLRE, interferons and c-KIT/SCF, while the latter include

Bcl-2, Bcl-XL, Mcl-1, NF-KB, survivin, livin and ML-IAP (Hussein et al., 2003). The key molecular components and mechanisms of apoptosis in melanoma have been discussed in previous reviews (Hussein et al., 2003, Soengas and Lowe, 2003, Erb et al., 2008).

As seen earlier (Chapter 5), postharvest method i.e. extraction solvent was found to influence the cytotoxic effect of C. nutans leaves on the human melanoma (D24 and MM418C1) and breast carcinoma (MCF7 and BT474) cell lines. The D24 melanoma cell line was the most sensitive to treatment, while the crude cH2O extract was the most active extract, which predominantly induced apoptotic cell death in the treated D24 cells. However, the effect of collection sites with different environmental conditions on the cytotoxicity of C. nutans leaf extract against cancer cells is still unknown. Therefore, the aims of this study were to i) compare and investigate the cytotoxicity of the crude MeOH extracts of C. nutans leaves collected from 11 different locations with varying environmental characteristics against the

D24 melanoma cells, ii) evaluate the cytotoxic effect and selectivity of the extract against the

D24 cells in a dose- and time-dependent manner and iii) examine the possible death mode of the D24 cells induced by the extract using biochemical and microscopy techniques.

6.2 Materials and methods

6.2.1 Plant materials

The collection of C. nutans leaves from 11 different locations in Malaysia, Thailand and

Vietnam was previously described in Section 3.2.1.

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6.2.2 Preparation of crude extracts

The preparation of the crude MeOH extract of C. nutans leaves from each location was previously described in Section 3.2.2. The extract was dissolved in DMSO (Sigma, USA) at a final concentration of < 0.1% (v/v). Each extract from one biological replicate was used in three independent experiments (analytical replication), unless otherwise stated.

6.2.3 Cell line and culture conditions

The extracts were tested on the D24 melanoma cells as they were shown to be the most sensitive towards treatment from previous study (Section 5.3.1) as well as the NHDF cells.

D24 and NHDF cells were cultured in RPMI 1640 medium containing L-glutamine (Gibco,

Life Technologies) and high-glucose DMEM medium with pyruvate and L-glutamine

(Gibco), respectively, supplemented with 10% FBS (Serana) and 1% (v/v) penicillin/streptomycin (Gibco). For all the experiments, the cells were incubated for the indicated time under the indicated treatment at 37°C in a humidified atmosphere of 5% CO2.

6.2.4 Cytotoxicity assay

The cytotoxicity assay was performed following the procedure as described in Section 5.2.4.

The percentage of viable cells was determined relative to the vehicle control (< 0.1% DMSO) using Equation 5.1. The vehicle control was expressed as 100%. Only the most active C. nutans sample (CT5) against the D24 cells as determined in this assay were used for further experiments.

6.2.5 Determination of EC50

The determination of EC50 was performed following the procedure as described in Section

5.2.5. The percentage of viable cells was determined relative to the vehicle control (100%).

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6.2.6 Observation of D24 cell morphology by phase contrast microscopy

As per Section 5.2.6.

6.2.7 Evaluation of apoptosis by Muse cytofluorimetric analysis

As per Section 5.2.7.

6.2.8 Detection of apoptosis and necrosis by Annexin V/PI double staining and confocal microscopy

As per Section 5.2.8.

6.2.9 Ultra structural analysis by TEM

As per Section 5.2.9.

6.2.10 Statistical analysis

All cell experiments were performed in triplicate, unless otherwise stated, and the results presented as mean ± SD. Data was analysed using statistical software Minitab 17 (Minitab,

2013). Significant difference was determined using ANOVA Fisher’s test at p ≤ 0.05 significance level. Correlations were calculated according to Pearson’s correlation coefficient

(r).

6.3 Results

6.3.1 Comparison of cytotoxicity among the crude methanol extracts of C. nutans leaves obtained from different locations against the D24 melanoma cells

There were differences in levels of cytotoxicity of the MeOH extracts (2 mg/mL) of C. nutans leaves from 11 different locations against the D24 cells after 72 h exposure (Figure

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6.1). All the extracts except for CT1 and CT2, were shown to be cytotoxic (p ≤ 0.05).

Cytotoxicity of all extracts varied from 19.8 to 91.0%, with a comparative mean effect of

52.5 ± 25.2%. Higher variation was observed between the Thai samples compared to the

Malaysian samples, where the latter possessed middle-range cytotoxicity levels. For the

Malaysian samples, CE4 from Kota Kinabalu had the highest activity, which was about 2.5- fold greater than that of the sample with the lowest activity (CE2 from Sandakan). In addition, even samples collected from the same region i.e. CE1 and CE2, showed significant (p ≤ 0.05) difference in their cytotoxic effects. For the Thai samples, CT5 from Chiang Dao was 5-fold more cytotoxic than CT2 from Map Khae, which had the least cytotoxicity. Samples CT1 and

CT2, although collected from the same region, had no significant difference in activity. Of the 11 C. nutans samples studied, only CE1, CE4, CT4, CT5 and CV1 at 2 mg/mL caused >

50% cytotoxicity when compared to the vehicle control (< 0.1% DMSO).

Figure 6.1. Effect of the crude MeOH leaf extract of C. nutans samples from 11 different locations at 2 mg/mL on the D24 cells after 72 h exposure. Cell viability was determined by WST-8. Data represents mean ± SD from three independent experiments (n = 3). * p ≤ 0.05, significantly different from vehicle control (< 0.1% DMSO).

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6.3.2 Cytotoxicity of the crude methanol C. nutans leaf extract against the D24 melanoma and NHDF cells

In Section 6.3.1, it was observed that the MeOH extract of CT5 was the most cytotoxic.

Hence, this sample was selected for further experiments, including determination of the EC50 for the leaf extract. The cells were treated with the extract (0-2 mg/mL) for 24 h and viability was measured by WST-8 assay. The MeOH extract at 1 and 2 mg/mL showed significant (p ≤

0.05) cytotoxicity, which caused 66.5 and 48.0% cell death, respectively, when compared to the vehicle control (< 0.1% DMSO) (Figure 6.2A). Viability of the D24 cells fell sharply after exposure for 24 h to lower concentrations (0.25 and 0.5 mg/mL) of the extract, even lower than those exposed for 72 h. However, increasing the extract concentration above 1 mg/mL showed a more gradual reduction of viable cells.

Meanwhile, all test concentrations of the MeOH extract significantly (p ≤ 0.05) reduced the percentage of viable D24 cells after 72 h exposure, when compared to the vehicle control (<

0.1% DMSO) (Figure 6.2A). Treatment with 2 mg/mL of the extract caused 88.0% cell death, almost 6-fold more cytotoxic than that with 0.25 mg/mL (15.3% cell death). Furthermore, there was a significant (p ≤ 0.05) difference in cytotoxic effect between 24 and 72 h at 2 mg/mL, suggesting that the observed effect was time dependent. The EC50 values for 24 and

72 h were 0.95 and 0.77 mg/mL, respectively.

As shown in Figure 6.2B, the MeOH extract showed low cytotoxicity against the normal

NHDF cells, where 24 and 72 h of exposure to 2 mg/mL of the extract caused 12.0 and 22.2% cell death, respectively. The EC50 values for the extract at 24 and 72 h were > 2 mg/mL, suggesting that the extract was more selective for the D24 melanoma cells than the normal cells.

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Figure 6.2. Cytotoxic effect of the crude MeOH C. nutans leaf extracts on the D24 melanoma (A) and the NHDF cells (B). Data represents mean ± SD from two independent experiments (n = 2). * p ≤ 0.05, significantly different from vehicle control (< 0.1% DMSO). † p ≤ 0.05, significantly different between treatment times.

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6.3.3 Morphological changes in the D24 melanoma cells treated with the crude methanol

C. nutans leaf extract

The untreated (control) D24 melanoma cells were adherent and exhibited a smooth surface and elongated structure. These cells also showed a finely granulated cytoplasm (Figure 6.3A

& D). In contrast, the cells treated with the extract displayed notable cell shrinkage with irregular and rough form and the cell number were lower than that of controls (Figure 6.3B, C,

Figure 6.3. Morphological changes (arrows) in the D24 melanoma cells treated with crude methanol C. nutans leaf extract. Phase contrast images (20×) of cells after 24 h exposure to no treatment (A), extract at 1 mg/mL (B) and 2 mg/mL (C); 72 h exposure to no treatment (D), extract at 1 mg/mL (E) and 2 mg/mL (F).

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6.3.4 Induction of mode of cell death in the treated D24 melanoma cells by the crude methanol C. nutans leaf extract

Treatment of the D24 cells with the extract at 1 mg/mL for either 24 or 72 h did not induce high affinity for both Annexin V and 7-AAD, indicating that most cells (> 50%) were viable

(bottom left quadrant) (Figure 6.4A & B1). Similarly, 24 h exposure to the extract at 2 mg/mL also showed that most of the D24 cells were viable. However, treatment with 2 mg/mL of the extract for 72 h remarkably increased the number of cells that were positive to both Annexin V and 7-AAD (upper right quadrant). This was also shown in Figure 6.4B2.

When the cells were treated with 2 mg/mL of the extract for 72 h, there was a significant (p ≤

0.05) increase in the percentage of late apoptotic/necrotic cells compared to the control.

Besides, when the cells were exposed to 2 mg/mL of the leaf extract for 72 h, the number of cells that had undergone apoptosis increased by almost 4-fold than those exposed for 24 h.

This indicated that a long exposure to a high concentration of the MeOH extract was more likely to induce late apoptosis/necrosis rather than early apoptosis in the D24 cells.

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Figure 6.4. Effect of the crude MeOH C. nutans leaf extract on the D24 melanoma cells. Representative apoptosis profile plots of the untreated and treated cells after 24 or 72 h exposure to 1 or 2 mg/mL of the extract (A). The percentage of viable (B1), early apoptotic and late apoptotic/necrotic (B2) cell populations of untreated and treated groups. Data represents mean ± SD from three independent experiments (n = 3). * p ≤ 0.05, significantly different from control. † p ≤ 0.05, significantly different between treatment times.

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Confocal microscopy was used to provide a qualitative identification of both apoptotic and necrotic deaths of the D24 cells treated for 72 h with 1 and 2 mg/mL of the extract (Figure

6.5). The control (untreated) cells (Figure 6.5A-D) did not stain following the addition of both Annexin V and PI, which showed that they were viable. Most of the D24 cells treated with 1 mg/mL of the extract were also negative to both fluorescent stains (Figure 6.5E-H).

However, early apoptosis (Annexin V +ve; PI -ve) (Figure 6.5F) as well as late apoptosis/necrosis (Annexin V +ve; PI +ve) (Figure 6.5H) were seen in several of the D24 cells treated with 1 mg/mL of the extract. On the other hand, most of the cells treated with the higher test concentration (2 mg/mL) were positive to both Annexin V and PI, which appeared red (Figure 6.5L), indicating the presence of late apoptotic/necrotic cells. The cells treated with 2 mg/mL of the extract had a higher level of these fluorescent stains than compared to those treated with 1 mg/mL, which suggests that the modes of D24 cell death were dose- dependent. The results obtained from confocal microscopy were similar to that obtained from the cytofluorimetric analysis.

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Figure 6.5. Confocal images (40 ×) of the untreated cells with no uptake of any stains (A-D) and the induction of early and late apoptosis/necrosis in the D24 cells after treatment with 1 mg/mL (E-H) and 2 mg/mL (I-L) of crude MeOH C. nutans leaf extract for 72 h. Continuous arrow indicate early apoptotic cells and dotted arrows indicate late apoptotic/necrotic cells.

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6.3.5 Ultrastructural changes in D24 melanoma cells treated with the crude methanol C. nutans leaf extract

As the crude MeOH C. nutans leaf extract was shown to induce cell death in the D24 cells

(Section 6.3.4), structural changes in the cells were further analysed electron microscopically to detect more details. In the D24 cells, clear morphological changes were observed between the untreated control and cells treated with the extract (2 mg/mL) for 72 h (Figure 6.6). The untreated cells displayed of normal cell characteristics, such as microvilli, intact plasma membrane and nucleus with evenly distributed chromatin (Figure 6.6A). In contrast, in the treated cells, there was cell shrinkage, loss of microvilli, marked peripheral chromatin condensation at the nuclear membrane (Figure 6.6B), segmented/lobulated nucleus and irregular plasma membrane with extensive blebbing (Figure 6.6C).

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PM NM

PM PM NM

Cr Nu Nu

Figure 6.6. Transmission electron micrographs of the control (untreated) D24 cell (A) and cells treated with the crude MeOH extract (2 mg/mL) for 72 h (B & C). Distinct morphological changes, including plasma membrane alteration (white arrow), chromatin condensation (white arrowhead), blebbing (black arrow) and segmented/lobulated nucleus (black arrowhead) were observed in the treated cells. Cr: chromatin, MV: microvilli, NM: nuclear membrane, Nu: nucleus, PM: plasma membrane.

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6.4 Discussion

6.4.1 Cytotoxicity against the D24 melanoma cells varies among the C. nutans leaves obtained from different locations and it correlates with the environment and concentration of phytochemical content

This study is the first to show variation in the cytotoxicity among the crude MeOH extracts of

C. nutans leaves collected from 11 different locations in Malaysia, Thailand and Vietnam, with diverse geographical and climatic conditions, against the D24 melanoma cells. Besides, this study is also the first to reveal significant correlations between cytotoxicity and several parameters, including elevation (Figure A3A, Appendix 5) and annual mean temperature

(Figure A3B) of sampling sites as well as the concentration of total phenolic (Figure A3D), flavonoid (Figure A3E) and polyphenol (in general) (Figure A3F) contents in the leaf extracts.

Current findings suggest that geographical origin of C. nutans may not be a major determinant for its cytotoxicity, but differences in elevation and climatic conditions within a geographical area may contribute to their variation. As reported (Section 3.3.1) and discussed

(3.4.1 and 3.4.2) previously, elevation and annual mean temperature significantly affected the total polyphenol contents in C. nutans leaves, thus its bioactivity, suggesting that samples from higher elevations and cooler climates are likely to have a higher cytotoxic effect against the D24 cells than from samples collected from lower elevations and warmer air temperature.

It has been suggested that environmental factors that influence growing conditions are important because they can interfere with the metabolic pathways in plants and therefore on the final concentration of bioactive compounds, which in turn affects the levels of their bioactivities (Akula and Ravishankar, 2011, Radušienė et al., 2012). The results of this study are in agreement with previous research, reporting variations in the levels of cytotoxicity of samples of different geographical origins with varying environmental conditions. Ayob et al.

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(2014) showed that crude methanol extracts of Justicia gendarussa (Acanthaceae) leaves from five different locations in Malaysia had different cytotoxic effects on MDA-MB-231 and -468 breast cancer cell lines. In another study, Basar et al. (2015) observed considerable variation in the cytotoxicity of methanol root extracts of Glycyrrhiza glabra L. (Fabaceae) samples from nine different countries, which were tested against immortal human keratinocyte (HaCaT), lung adenocarcinoma (A549) and HepG2 cell lines.

6.4.2 The crude methanol C. nutans leaf extract exhibits selective cytotoxicity against the D24 melanoma cells

The crude MeOH extract of C. nutans showed significant cytotoxicity against the D24 melanoma, but not the NHDF cells, which suggests that the extract is selective against cancer cells but not normal cells. Results of this study are in agreement with previous findings, including the results from Chapter 5 (crude cH2O extract), reporting that C. nutans leaves do have anticancer properties, although different extract and cancer cell line were used in the current study. Yong et al. (2013) tested three crude leaf extracts (chloroform, methanol and aqueous) on different human cancer cell lines i.e. HepG2, neuroblastoma (IMR-32), NCI-23,

SNU-1, LS-174T, HeLa, K-562, and Raji, and found that chloroform extract had the highest antiproliferative effect against the latter two cell lines. Another study by Arullappan et al.

(2014) also tested three crude leaf extracts of C. nutans (petroleum ether, ethyl acetate and methanol) on HeLa and K-562 cells, and found that petroleum ether extract had the strongest cytotoxic activity against both cell lines. The cytotoxic activity of C. nutans leaves may be due to the presence of flavonoids, such as C-glycosyl flavones, which have been shown in

Mimosa pudica (Fabaceae) (Li et al., 2011b) and Isodon lophanthoides var. gerardianus

(Lamiaceae) (Zhang et al., 2015) to have inhibitory effect on the proliferation of cancer cells.

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6.4.3 The crude methanol C. nutans leaf extract induces apoptosis in the D24 melanoma cells

This study is the first to examine the cell death modes of D24 melanoma cells treated with the crude MeOH extract in vitro. Muse cytofluorimetric analysis using the characterisation of biochemical features (Annexin V to detect apoptotic cells that have expressed phosphatidylserine externalisation on the cell surface and 7-AAD to distinguish dead cells) revealed a significant increase in the percentage of late apoptotic/necrotic and a significant decrease in viable cell populations, particularly at the highest test concentration (2 mg/mL) and longest treatment time (72 h), compared to the untreated controls. Besides, visual assessment of the confocal images of the Annexin V/PI double-stained D24 cells treated with the leaf extract confirmed the presence of late apoptotic/necrotic cells, although a few early apoptotic cells were observed at the lowest test concentration. Therefore, results of this study suggest that the extract induces late apoptotic/necrotic cell death in the D24 cells in a dose- and time-dependent manner.

Although utilisation of Annexin V, 7-AAD and PI is a standard procedure to monitor the progression of apoptosis, this method is not able to distinguish between late apoptotic and primary necrotic cells, since both groups are positive to Annexin V and 7-AAD/PI (Brauchle et al., 2014), as shown from the results of Muse cytofluorimetric and confocal analyses.

Therefore, to further determine the mode of cell death of D24 induced by the crude MeOH extract of C. nutans, detection of cell morphology changes using other types of microscopy techniques, including phase-contrast and transmission electron were used.

Optical and electron microscopy have been used to identify the various morphological changes that occur during apoptosis, although the latter gives better definition of subcellular

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CHAPTER 6 changes (Elmore, 2007). Phase contrast images and electron micrographs of the treated D24 cells revealed typical morphological features of apoptosis, including cell shrinkage as a result of condensation of organelles and the density of cytoplasm; chromatin condensation peripherally at the nuclear membrane; fragmented and/or lobulated nucleus; and extensive blebbing of plasma membrane, budding into apoptotic bodies consisting of cytoplasm with tightly packed organelles with or without nuclear fragments (Chen et al., 2001, Ziegler and

Groscurth, 2004, Elmore, 2007, Svejda et al., 2010). It is not always easy to distinguish apoptosis from necrosis, especially using conventional histology, and both events can happen simultaneously depending on factors like the concentration and exposure time of stimulus, the degree of ATP depletion and the availability of caspases (Zeiss, 2003). Regardless, some of the classical features of necrosis include cell swelling, highly vacuolated cytoplasm and disrupted cell membrane that becomes permeable, resulting in the release of cellular contents

(Ziegler and Groscurth, 2004, Elmore, 2007). However, the treated D24 cells did not show these ultrastructural features when observed under a transmission electron microscope and hence, results of this study may suggest that apoptotic cell death as the most likely mechanism involved, although the compound or component of the crude MeOH leaf extract of C. nutans that triggered apoptosis is unknown. Further studies and time points need to be investigated to confirm that the crude MeOH extract induces the apoptotic pathway in these cells.

Several medicinal plant species of the family Acanthaceae have been reported to induce apoptotic cell death in different cancer cell lines. For example, Justicia spicigera

(Acanthaceae) leaves have shown to induce apoptosis in mouse fibroblasts (3T3), human cervical carcinoma (CALO and INBL) (Cáceres-Cortés et al., 2001) and HeLa (Alonso-

Castro et al., 2012) cells, while crude leaf extracts of Ruellia tuberosa (Acanthaceae) and

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Andrographis paniculata (Acanthaceae) exhibited potent apoptogenic activity on HePG2

(Dey et al., 2013) and KB (Saengkhae and Meewassanasuk, 2009) cells, respectively. It has been described that a synergistic activity of IFN-γ and tumour necrosis factor (TNF)-α strongly induces apoptosis in HaCaT keratinocyte cells (Konur et al., 2005). However,

Thongrakard and Tencomnao (2010) showed that the crude ethanol C. nutans leaf extract significantly inhibited the induction of apoptosis by IFN-γ/TNF-α in HaCaT cells.

Nevertheless, it should be noted that the stereotype outcome either as apoptosis or necrosis cannot always be expected. This is because induction or inhibition of cell death modes depends on a number of factors, such as the plant species, preparation methods (crude extracts, fractions and isolated compounds), concentrations and exposure durations of stimuli, cell types and the nature of the cell death signal (Hirsch et al., 1997, Zeiss, 2003, Ahmad et al., 2010, Cock, 2011).

6.5 Conclusion

Several noteworthy conclusions can be drawn from this study on the comparative cytotoxicity of C. nutans leaves from various locations and the anticancer activity of the crude MeOH leaf extract against the D24 melanoma cells. First, the crude MeOH extracts of leaves from 11 different locations in Malaysia, Thailand and Vietnam, with diverse geographical and climatic conditions showed variation in cytotoxicity against the D24 melanoma cells as assessed by WST-8 assay. The leaves of plants grown at higher elevations and lower air temperatures exhibited higher levels of cytotoxicity than those grown at lower elevations and higher air temperatures. This suggests that the anticancer potential of C. nutans leaves can be considerably variable depending on the collection sources. The results could assist in the selection of the most favourable environmental conditions to produce C. nutans with the highest medicinal value. Second, the crude MeOH leaf extract of the most active sample

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(CT5 from Chiang Dao, Chiang Mai, Thailand) showed significant cytotoxicity against the cancer cells (D24 melanoma), but considerably less towards the non-cancerous NHDF cells, suggesting selective killing of cancer cells by the extract. Third, the extract induced cell death in the D24 cells, most likely by apoptosis in a dose- and time-dependent manner, as shown by the biochemical and morphological changes detected by cytofluorimetry and microscopy approaches, respectively. These observations suggest that the crude MeOH C. nutans leaf extract could be used to supplement current regimens used for cancer prevention or treatment.

However, further work is needed to elucidate the structure(s) of the active compound(s) or component(s) in the crude MeOH C. nutans leaf extract, as well as to determine their anticancer efficacies. In addition, further research to understand the underlying mechanism in cell death, especially the cellular signalling pathways involved in induced apoptosis and in vivo testing of the observed anticancer activity are essential to unveil the full potential use of

C. nutans in cancer therapy.

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Synopsis and future directions

7.1 Influence of collection sites with different environmental characteristics on the phytochemical constituents and anticancer activity of C. nutans leaves

The leaves of C. nutans grown at higher elevations with lower air temperatures (i.e. CT5 from Chiang Dao, Chiang Mai, Thailand) had higher TPC and TFC as determined using the colourimetric (Folin-Ciocalteu and AlCl3) (Yuann et al., 2012, Ghasemzadeh et al., 2014,

Tiew et al., 2014, Wong et al., 2014, Lusia Barek et al., 2015, Mustapa et al., 2015) and

HPLC (Chelyn et al., 2014, Ghasemzadeh et al., 2014, Mustapa et al., 2015) methods, compared to those grown at lower elevations with higher air temperatures. Results demonstrated that the level of phytochemical constituents in C. nutans leaves may be associated with growing conditions, where higher elevations and lower air temperatures are beneficial to the overall production of phytochemicals (Kishore et al., 2010, Dong et al., 2011,

Guo et al., 2011, Bernal et al., 2013). The cytotoxicity of the C. nutans leaf extracts against

D24 melanoma cells was measured using the WST-8 assay. The cytotoxicity was found to be significantly (p ≤ 0.05) correlated to the total phytochemical content in the leaves. Therefore, the crude MeOH extracts of C. nutans leaves from higher elevations and lower air temperatures exhibited higher cytotoxicity against the cancer cells. The leaf extract of C. nutans grown in Chiang Dao, Chiang Mai, Thailand, showed the highest activity (24 h EC50:

0.95 mg/mL and 72 h EC50: 0.77 mg/mL). The leaf extract from Chiang Dao was more cytotoxic against the cancer cells but not dermal fibroblasts. Moreover, the extract induced apoptotic cell death in the D24 melanoma cells in a dose- and time-dependent manner as determined by cytofluorimetry (Engel et al., 2014, Jiang et al., 2014, Nair et al., 2014) and confocal microscopy (Ismail et al., 2013, Issara-Amphorn and T-Thienprasert, 2014) using

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Annexin V, 7-AAD and PI fluorescent stains. In addition, ultrastructural changes showing typical characteristics of apoptosis were observed in the D24 cells treated with 2 mg/mL of the extract using TEM (Tan et al., 2005, Ait-Mohamed et al., 2011, Echiburú-Chau et al.,

2014). The results of this study may suggest that apoptosis as the most likely mechanism involved, although the compound or component of the crude MeOH leaf extract of C. nutans that triggered the mode of cell death is unknown.

Genetic study of C. nutans using RAPD and microsatellite markers revealed high genetic similarity, with an average of 77.0% and 91.6%, respectively, among the vegetatively propagated C. nutans samples from Malaysia, Thailand and Vietnam, with identical genetic profiles even though they were geographically distant. However, PCR-RFLP of the rbcL region could not discrimate the C. nutans samples, suggesting that this technique is not suitable to analyse the genetic variation in C. nutans. The rbcL region is known to be highly conserved with low gene substitution rates in perennial angiosperm taxa (Bousquet et al.,

1992, Frascaria et al., 1993), which makes it more suited for genetic studies of higher taxonomic levels (intergeneric to subclass) but not lower levels (inter- to intraspecies) (Arca et al., 2012). The low genetic diversity in C. nutans may be due to a combination of evolutionary history or the long-term artificial vegetative propagation by breeders. PCR-

RFLP, RAPD and microsatelite marker techniques were able to differentiate A. paniculata from C. nutans at the intergeneric level (Lakshmi et al., 2002). However, only RAPDs and microsatellites were able to distinguish C. siamensis from C. nutans, suggesting that these molecular markers are more suitable for identifying the three plant spcies, which may be useful for authentication and quality assessment (Li et al., 2007, Khan et al., 2011, Ding et al.,

2012, Cichorz et al., 2014).

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These results suggest that the cytotoxic activity of C. nutans leaf extract against the D24 melanoma cells may be determined by its phytochemical constituents, which can be influenced by environmental variables, particularly elevation and air temperature, rather than genetic factors. This relationship has also been observed by others (Grass et al., 2006, Šamec et al., 2015), describing that variation in phenolics is mainly caused by environmental factors than genetic factors. This study provides a better understanding on the effect of collection sites with different environmental characteristics on the phytochemical constituents and hence, the bioactivity of C. nutans. Results will provide useful information for cultivars on the optimum growing conditions for C. nutans in order to obtain the highest medicinal value and the maximum therapeutic benefits from the plant. Besides, observations from the cytotoxicity study suggest that the crude MeOH C. nutans leaf extract could be used to supplement current regimens used for cancer prevention or treatment.

7.2 Influence of postharvest processes on the phytochemical constituents and anticancer activity of C. nutans leaves

The effect of postharvest treatments, including drying temperature and extraction solvent, on the phytochemical constituents of C. nutans leaves was also successfully investigated using colourimetric and HPLC methods. However, only the effect of extraction solvent on the viability of four human cancer cell lines: D24 (wild type BRAF) and MM418C1 (BRAF

V600E) melanoma cells; and MCF7 (wild type p53) and BT474 (ts-p53 E285K) breast carcinoma cells, was examined in the current study. Temperature was found to significantly influence the level of phytochemicals in the C. nutans leaves during drying (Hansen et al.,

1993, Periche et al., 2015). An increase in the amount of phytochemical constituents was observed with increasing drying temperatures, where leaves dried at 100°C and 80°C had the highest TPC and TFC, respectively. HPLC analysis also showed the greatest amount of

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CHAPTER 7 phytochemicals in the C. nutans leaves dried at 80°C. This suggests that the use of low temperatures may not be a suitable postharvest method for drying C. nutans leaves (Chen et al., 2011, Shotorbani et al., 2013, Lopez et al., 2013, Periche et al., 2015). Likewise, the type of extraction solvent also affected the total phytochemcial content extracted from the leaves reviews (Tanko et al., 2005, Naczk and Shahidi, 2006, Stalikas, 2007, Rocha et al., 2011).

The most polar solvent (H2O) resulted in higher total phytochemical content being extracted from the leaves compared to MeOH, DCM and EtOH (Nyirenda et al., 2012). Moreoever, the temperature of extraction solvent also significantly affected the total phenolic and flavonoid content being extracted from these leaves (Naczk and Shahidi, 2006, Stalikas, 2007). The crude cH2O C. nutans leaf extract had the greatest quantities of both phenolics and flavonoids, suggesting that cold extraction (22°C) with water is the best method to prepare the crude extract of these leaves with maximum phenolics and flavonoids (Venditti et al., 2010, Dent et al., 2013). However, HPLC analysis revealed greater amount of phytochemicals in the crude hH2O extract than that of the cH2O extract. This may be due to the presence of other compounds, besides phenolics and flavonoids, which may have overlapping absorption bands, thus, contributing to the sum of the peak areas.

The study on the effect of extraction solvent on the viability of the four tested cancer cell lines revealed variation in the effect of the different crude extracts at 2 mg/mL. This indicates that the solubility of bioactive compounds in the extracts is significantly influenced by the extraction method (Naczk and Shahidi, 2006). The D24 melanoma cell line was the most sensitive to treatment and both the crude cH2O and hH2O extracts were the most active against all cancer cell lines, whereas the crude MeOH, EtOH and DCM extracts showed low to no cytotoxicity against these cell lines (Madhulika and Ajitkumar, 2010, Moghe et al.,

2011, Uddin et al., 2011, Elias et al., 2013). Further evaluation on the cytotoxic effect of the

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cH2O and hH2O extracts on the D24 cells showed that the former (72 h EC50: 1.63 mg/mL) had greater selective effect than the latter (72 h EC50: > 2 mg/mL). Both extracts exhibited selective cytotoxicity towards the cancer cells but not the normal dermal fibroblasts. Results are in agreement with that of the colourimetric methods, suggesting that the extraction with water at 22°C is the best method to extract the bioactive compound(s) with anticancer property from C. nutans leaves, which remains to be isolated and identified. According to the biochemical and morphological changes detected by cytofluorimetry and microscopy approaches, respectively, the crude cH2O extract predominantly induced apoptotic cell death in the D24 cells in a dose- and time-dependent manner. Moreover, necrosis was also detected in the treated cells, possibly due to the combination of different bioactive compounds present in the crude cH2O extract. This study provides crucial information on the optimisation of postharvest methods to prepare crude extracts that preserve the bioactive phytochemicals from dry C. nutans leaves. This may increase the potential of C. nutans as a valuable medicinal product.

C. nutans holds much promise as a therapeutic agent against melanoma. However, its efficiency is currently not optimised due to limited information on preservation of bioactive compounds (e.g. phenolics and flavonoids). This study has dispelled current beliefs and contributed significantly to improve C. nutans products in future by advising primary producers and manufacturers on best practices.

7.3 Future directions

Since there is a lack of genetic diversity in C. nutans at the present time, genetics are not significant in the designing of conservation management plans for this plant. The most appropriate measure is to reduce continuous breeding of the plant by vegetative propagation

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CHAPTER 7 and continuing it through mature stage with the development of reproductive features, where possible. With reproductive features, sexual reproduction of C. nutans may be feasible, hence, increasing its genetic diversity, which in turn, may aid in long-term survival and evolution of the species. Future genetic studies would benefit from a larger sample size as well as the use of more primers to analyse diversity in order to draw more robust conclusions about the genetic variability of C. nutans.

Current findings suggest that the phytochemical constituents in C. nutans leaves, which determined their cytotoxic effects against cancer cells, are more likely to be affected by environmental factors, such as elevation and air temperature, instead of genetic factors.

Certainly, other environmental factors may also influence the levels of phytochemicals in C. nutans. The leaves of C. nutans used in this study were from plants grown in their natural environment or in the field. The interaction between the biosynthesis of secondary metabolites in C. nutans and the natural environment can be complicated by multiple effects, which these effects are difficult to separate from each other. In order to study specific environmental factors on the phytochemical constituents of C. nutans, it is recommended to grow the plant under a controlled environment (greenhouse system) to facilitate a better understanding of single factors. It is also critical to investigate the effect of the environment on individual phytochemicals to assist improved production of bioactive compounds.

However, structural elucidation and identification of the bioactive compounds are necessary before the determination of the relevant metabolic pathways that can be influenced by environmental changes. Besides, a larger sample size i.e. collection of more than one plant from a single location, as well as a selection of wider environmental range may be useful to produce a stronger conclusion on the effect of environmental factors on the levels of phytochemicals in C. nutans leaves.

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One of the major outcomes for this study is that the crude extracts (cH2O and MeOH) of C. nutans leaves exhibited cytotoxic activity against the D24 melanoma cells. Although results suggest that the bioactive compound that is responsible for the activity may be a phenolic or a flavonoid, the specific compound is still unknown. Therefore, further studies are needed to elucidate the structure of the bioactive compounds or components in the crude cold aqueous and methanol C. nutans leaf extracts and evaluate their efficacies in vitro and in vivo.

However, pure compounds isolated from plants may not have greater bioactivity and efficacy than crude extracts due to the synergistic effect of several components in the crude mixture, such as that observed with Aloe vera (Cock, 2011), but at present remains unknown for C. nutans. Therefore, there is a need to investigate this potential discrepancy between the crude extract and bioactive compound isolated from C. nutans.

The crude cH2O and MeOH C. nutans leaf extracts showed higher cytotoxic effect on D24 melanoma cells, which bear the wild type Braf gene than on MM418C1 cells, which harbour the mutated Braf gene at codon 600. This suggests that these crude leaf extracts may have bioactive compouds that affect the Raf/MEK/ERK cellular signalling pathway in different manners but this effect is unknown. Future studies are needed to investigate the responsible compounds and their mode of action on this pathway.

Although this study has shown the induction of apoptotic cell death in the treated D24 melanoma cells by the crude cH2O and MeOH C. nutans leaf extracts, the exact underlying mechanisms and the cellular signalling pathways (e.g. intrictic or extrinsic) involved remain to be identified. Besides, both apoptotic and necrotic cell deaths were induced in the treated

D24 cells by the crude cold aqueous leaf extract. This may be due to the combination of effects of the different components in the crude extract. However, this finding needs to be

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CHAPTER 7 clarified. Identifying the signalling pathways that initiate the cell death in the treated D24 cells is an important step in understanding not only melanoma, but cancer as a whole. This may lead to designing effective strategies for the treatment of cancer, resulting in the development of more effective anticancer drugs.

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Appendices

Appendix 1

Table A1. Similarity matrix of the twelve C. nutans, one C. siamensis and two A. paniculata samples and the outgroup based on the RAPD data calculated using Dice’s coefficient. The genetic similarity coefficient value between samples ranged from 0 – 1, where 0 indicated no similarity and 1 indicated that the two samples were genetically identical. CP1 CP2 CE1 CE2 CE3 CE4 CT1 CT2 CT3 CT4 CT5 CV1 CS AP1 AP2 MC CP1 1.0000 CP2 1.0000 1.0000 CE1 1.0000 1.0000 1.0000 CE2 1.0000 1.0000 1.0000 1.0000 CE3 1.0000 1.0000 1.0000 1.0000 1.0000 CE4 1.0000 1.0000 1.0000 1.0000 1.0000 1.0000 CT1 0.8000 0.8000 0.8000 0.8000 0.8000 0.8000 1.0000 CT2 0.8000 0.8000 0.8000 0.8000 0.8000 0.8000 1.0000 1.0000 CT3 0.8000 0.8000 0.8000 0.8000 0.8000 0.8000 1.0000 1.0000 1.0000 CT4 0.8000 0.8000 0.8000 0.8000 0.8000 0.8000 1.0000 1.0000 1.0000 1.0000 CT5 0.7273 0.7273 0.7273 0.7273 0.7273 0.7273 0.8889 0.8889 0.8889 0.8889 1.0000 CV1 0.7692 0.7692 0.7692 0.7692 0.7692 0.7692 0.7273 0.7273 0.7273 0.7273 0.6667 1.0000 CS 0.2222 0.2222 0.2222 0.2222 0.2222 0.2222 0.2857 0.2857 0.2857 0.2857 0.2500 0.4000 1.0000 AP1 0.0000 0.0000 0.0000 0.0000 0.0000 0.0000 0.0000 0.0000 0.0000 0.0000 0.0000 0.1538 0.0000 1.0000 AP2 0.2667 0.2667 0.2667 0.2667 0.2667 0.2667 0.3077 0.3077 0.3077 0.3077 0.2857 0.2500 0.0000 0.5333 1.0000 MC 0.0000 0.0000 0.0000 0.0000 0.0000 0.0000 0.0000 0.0000 0.0000 0.0000 0.1111 0.0000 0.0000 0.0000 0.0909 1.0000

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Appendix 2

Table A2. Similarity matrix of the twelve C. nutans, one C. siamensis and two A. paniculata samples and the outgroup based on the microsatellite data calculated using Dice’s coefficient. The genetic similarity coefficient value between samples ranged from 0 – 1, where 0 indicated no similarity and 1 indicated that the two samples were genetically identical. CP1 CP2 CE1 CE2 CE3 CE4 CT1 CT2 CT3 CT4 CT5 CV1 CS AP1 AP2 MC CP1 1.0000 CP2 1.0000 1.0000 CE1 1.0000 1.0000 1.0000 CE2 1.0000 1.0000 1.0000 1.0000 CE3 1.0000 1.0000 1.0000 1.0000 1.0000 CE4 1.0000 1.0000 1.0000 1.0000 1.0000 1.0000 CT1 0.8667 0.8667 0.8667 0.8667 0.8667 0.8667 1.0000 CT2 0.8667 0.8667 0.8667 0.8667 0.8667 0.8667 1.0000 1.0000 CT3 0.8667 0.8667 0.8667 0.8667 0.8667 0.8667 1.0000 1.0000 1.0000 CT4 0.8667 0.8667 0.8667 0.8667 0.8667 0.8667 1.0000 1.0000 1.0000 1.0000 CT5 0.8667 0.8667 0.8667 0.8667 0.8667 0.8667 0.9333 0.9333 0.9333 0.9333 1.0000 CV1 0.8966 0.8966 0.8966 0.8966 0.8966 0.8966 0.9655 0.9655 0.9655 0.9655 0.8966 1.0000 CS 0.5926 0.5926 0.5926 0.5926 0.5926 0.5926 0.6667 0.6667 0.6667 0.6667 0.6667 0.6923 1.0000 AP1 0.2963 0.2963 0.2963 0.2963 0.2963 0.2963 0.2963 0.2963 0.2963 0.2963 0.2963 0.3077 0.1667 1.0000 AP2 0.2308 0.2308 0.2308 0.2308 0.2308 0.2308 0.3077 0.3077 0.3077 0.3077 0.3077 0.2400 0.0870 0.8696 1.0000 MC 0.2000 0.2000 0.2000 0.2000 0.2000 0.2000 0.2667 0.2667 0.2667 0.2667 0.2667 0.2759 0.2222 0.2222 0.2308 1.0000

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Appendix 3

Figure A1. Standard curves of gallic acid (A) and quercetin (B).

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Appendix 4

Figure A2. Scatter plots showing correlations between environmental factors, including elevation (A), annual mean temperature (B) and rainfall (C) and the TPC and TFC of C. nutans as well as between the TPC and TFC (D). ** p ≤ 0.01 and NS not significant.

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Appendix 5

Figure A3. Scatter plots showing correlations between cytotoxicity of the crude MeOH leaf extracts of 11 C. nutans samples and different parameters, including elevation (A), annual mean temperature (B), rainfall (C), total contents of phenolic (D), flavonoid (E) and phytochemical level (in general) as determined by HPLC at 332 nm (F). * p ≤ 0.05.

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