Isolation and Characterization of Plant Based Pesticides

Saira Khan

CIIT/FA11-R62-001/ATD

PhD Thesis

Biotechnology

COMSATS University Islamabad Abbottabad Campus - Pakistan

Fall, 2017 COMSATS University Islamabad

Isolation and Characterization of Plant Based Pesticides

A Thesis Presented to

COMSATS University Islamabad, Abbottabad Campus

In partial fulfillment

of the requirements for the degree of

PhD

(Biotechnology)

Saira Khan

CIIT/FA11-R62-001/ATD

Fall, 2017

Isolation and Characterization of Plant Based Pesticides

A Post Graduate Thesis submitted to the Department of Environmental Sciences as partial fulfillment of the requirements for the award of Degree of Ph.D in Biotechnology.

Name Registration Number Saira Khan CIIT/FA11-R62-001/ATD

Supervisor

Dr. Mohammad Maroof Shah Professor, Biotechnology Program Department of Environmental Sciences COMSATS University Islamabad, Abbottabad Campus

Co-Supervisors 1. Dr. Raza Ahmad Associate Professor, Biotechnology Program Department of Environmental Sciences COMSATS University Islamabad, Abbottabad Campus

2. Dr. Guy Smagghe Professor, Department of Crop Protection Faculty of Bioscience Engineering, Ghent University, 9000 Ghent, Belgium

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DEDICATION To

My Parents, Husband and Kids

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ACKNOWLEDGEMENTS

I am first of all thankful to Almighty Allah (SWT) who gave me all the strength, courage, determination, and true guidance in conducting this study and completing my overall academic endeavors despite many hurdles and difficulties. I am grateful to my supervisor Prof. Dr. Mohammad Maroof Shah, for his excellent supervision, continuous guidance, relentless efforts and support throughout my research and academic career. He always pushed me forward and encouraged to have a belief in myself. I extend my gratitude to my co-supervisors, Dr. Raza Ahmad at CIIT, and Prof. Dr. Guy Smagghe, Department of Crop Protection, Ghent University, Belgium for their guidance and assistance in completing major part of my research. Thanks are extended to Prof. Dr. Sven Mangelinckx, Department of Sustainable Organic Chemistry and Technology, Faculty of Bioscience Engineering, Ghent University, Belgium for his guidance and support. I am thankful to the Higher Education Commission of Pakistan for providing me an opportunity to visit the Ghent University under International Research Support Initiative Program. I express deep gratitude to members of my supervisory committee including Dr. Amjad Hassan, Dr. Jamshaid Hussain and Dr. Sarfraz Shafiq for their great help and support in research work and completion of thesis at CIIT Abbottabad. I am very thankful to Dr. Ihsan ul Haq, Insect Pest Management Program, National Agriculture Research Center (NARC), Islamabad, for his guidance and support to complete a part of my research work. Thanks are also extended to Prof. Dr. Arshid Pervez, and to all faculty members of the Biotechnology program and Environmental Sciences Department for long-lasting interest and unbendable support throughout the course of my studies and research work. I am grateful to my friends and colleagues for their great and helpful support during research work especially Dr. Nighat Fatima, Dr. Irum Shehzadi, Miss Sadia Bibi, Miss Asma Khan, and Mr. Clauvis Nji Tizi Taning and Mr. Elias Bonneure. I am highly obliged to my parents and father in law, sisters and brothers who supported me throughout my life and academic career. My deepest gratitude goes to my parents who taught me the importance of having dreams, setting of goals, achieving the targets and the art of hard work. Profound, sincere and cordial thanks to my husband, Mr. Mohammad Asif Khan for his care, cooperation, encouragement and moral support during research and thesis work which enabled me to proceed and complete this task. At the end, I would like to thank my kids, Zaina Asif and Muhatasim Asif, who suffered a lot during my whole research period. .

Saira Khan CIIT/FA11-R62-001/ATD

ABSTRACT

Isolation and Characterization of Plant Based Pesticides With the pace of constant population growth, the demand for sufficient and safer food is continuously increasing around the globe. On the other hand, global loss to crops due to pests, diseases and weeds is significantly high, warranting excessive use of pesticides, threatning environmnet and food safety. The most frequently used pesticides are synthetic posing several associated pre and post application problems such as residual toxicity that results in compromising the safety of food and causing insect resistance. An alternative approach may be to utilize plant’s secondary metabolites that plants actually synthesize in their defense against pests and pathogens. The major aim of current research study was, therefore, to identify, isolate, and characterize at biochemical and molecular level the potent insecticidal compounds from plant sources. To achieve this aim, seven plants namely Cinnamomum camphora, Eucalyptus sideroxylon, Isodon rugosus, Boenninghausenia albiflora, Calotropis procera, Daphne mucronata, and Tagetes minuta were selected. The crude and purified extracts of each of these plants were used to screen for their toxic effects against six economically important agricultural pests, each representing a separate insect order; Acyrthosiphon pisum (Hemiptera), Drosophila melanogaster (Diptera), Tribolium castaneum (Coleoptera), Spodoptera exigua (Lepidoptera), Schizaphis graminum (Hemiptera) and Bactrocera zonata (Diptera). Aphids were the most susceptible insects with 100% mortality observed after 24 h for all the plant extracts tested. Further bioassays with lower concentrations of the plant extracts against aphids revealed that the extracts from Isodon rugosus (Lamiaceae) (LC50 36.2 ppm and LC90 102.1 ppm) and Daphne mucronata (Thymelaeaceae) (LC50 126.2 ppm and LC90 197.5 ppm) found out to be the most toxic to aphids, A. pisum. These most toxic and active plant extracts were further fractionated in different solvent fractions on polarity basis and their insecticidal activity was further evaluated. While all fractions showed considerable mortality in aphids, the most active was the butanol fraction from Isodon rugosus with an LC50 of 18 ppm and LC90 of 48.2 ppm. Further bioactivity guided fractionation of the butanol fraction results in isolation of bioactive principle compound that was identified through various spectroscopic techniques as rosmarinic acid with LC50 0.2 ppm and LC90 5.4 ppm. There was no significant difference between LCs of purified rosmarinic acid and of commercial rosmarinic acid. Further, two key genes, hydroxyphenylpyruvate reductase and rosmarinic acid synthase, known to involve in biosynthesis of rosmarinic acid were targeted to clone from Isodon rugosus. Only one of these genes, hydroxyphenylpyruvate reductase was successfully cloned in Isodon rugosus which consequently will open the way to explore all other genes responsible for biosynthesis of rosmarinic acid. The molecular knowledge regarding biosynthetic pathway will help in biotechnological production of rosmarinic acid and to produce aphid resistant plants through genetic engineering approaches. Considering the high mortality rate in aphids to a significantly low concentration of the rosmarinic acid from Isodon rugosus, could be exploited and further developed as a potential eco-friendly plant-based insecticide against sucking insect pests.

TABLE OF CONTENTS

1 Introduction…………………………………………………………... 1 1.1 Synthetic pesticides……………………………………………… 2 1.1.1. Organochlorine………………………………………….. 2 1.1.2 Organophosphate………………………………………... 3 1.1.3 Carbamate………………………………………………. 4 1.1.4 Pyrethroid………………………………………………. 4 1.1.5 Neonicotinoids…………………………………………. 5 1.1.6 Disadvantages of synthetic pesticides………………….. 6 1.2 Alternative approaches to overcome pest’s losses……………… 7 1.2.1 Microbial pesticides…………………………………….. 7 1.2.1.1 Entomopathogenic fungi……………………… 7 1.2.1.2 Viral biopesticides…………………………….. 8 1.2.1.3 Bacterial biopesticides………………………… 8 1.2.2 Plant incorporated protectants (PIPs)…………………… 9 1.2.3 Biochemical pesticides (Botanical pesticides)………….. 9 1.3 Botanical pesticides ……………………………………………... 10 1.3.1 History of botanical pesticides………………………….. 10 1.3.2 Types of botanical pesticides…………………………… 11 1.3.2.1 Pyrethrins (Pyrethrum/Pyrenone)…………….. 12 1.3.2.1.1 Mode of action…………………… 13 1.3.2.2 Rotenone……………………………………… 13 1.3.2.2.1 Mode of action…………………… 13 1.3.2.3 Nicotine………………………………………. 14 1.3.2.3.1 Mode of action…………………… 14 1.3.2.4 Sabadilla……………………………………… 14 1.3.2.4.1 Mode of action…………………… 15 1.3.2.5 Ryania………………………………………… 15

1.3.2.5.1 Mode of action…………………… 16 1.3.2.6 Limonene……………………………………… 16 1.3.2.6.1 Mode of action…………………… 16 1.3.2.7 Azadirachtin…………………………………… 16 1.3.2.7.1 Mode of action…………………… 17 1.3.3 Modes of application of botanical pesticides…………… 17 1.3.3.1 Fumigation toxicity…………………………… 17 1.3.3.2 Contact toxicity………………………………. 18 1.3.3.3 Repellent activity…………………………….. 18 1.3.3.4 Oviposition deterrent and adult emergence inhibition behavior…………………………… 18 1.3.3.5 Feeding deterrents……………………………. 19 1.3.4 Overview on plant products and extracts used as botanical pesticides…………………………………….. 19 1.4 Overview on selected plants…………………………………… 21 1.4.1 Boenninghausenia albiflora (Hook.) Rchb. ex Meisn. 21 (Rutaceae)……………………………………………… 1.4.2 Calotropis procera Aiton (Dryand). (Apocynaceae)…… 22 1.4.3 Tagetes minuta L. (Asteraceae)………………………… 23 1.4.4 Cinnamomum camphora (L.) J. Presl (Lauraceae)…….. 24 1.4.5 Eucalyptus sideroxylon A. Cunn. ex Woolls (Myrtaceae) 25 1.4.6 Daphne mucronata Royle (Thymelaeaceae)…………… 26 1.4.7 Isodon rugosus Wall. ex Benth (Labiatae)……………... 27 1.5 Overview on targeted insects…………………………………… 29 1.5.1 Bactrocera zonata Saunders (Diptera)…………………. 29 1.5.2 Drosophila melanogaster Meigen (Diptera)…………… 30 1.5.3 Tribolium castaneum Herbst (Coleoptera)……………… 31 1.5.4 Spodoptera exigua (Hubner) (Lepidoptera)…………….. 32

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1.5.5 Schizaphis graminum Rondani (Hemiptera)……………. 33 1.5.6 Acyrthosiphon pisum Harris (Hemiptera)………………. 34 1.6 Theoretical underpinning……………………………………….. 35 1.7 Objectives………………………………………………………. 38 2 Materials and Methods……………………………………………… 39 2.1 Pesticidal potential of selected plants against target pests……… 40 2.1.1 Plant material…………………………………………… 40 2.1.2 Preparation of plant extracts……………………………. 40 2.1.3 Target insects…………………………………………… 41 2.2 Bioassays mediated pesticidal potential of selected plants……... 42 2.2.1 Bactrocera zonata Saunders, 1842…………………….. 42 2.2.1.1 Rearing of Bactrocera zonata………………….. 42 2.2.1.2 Adult male fruit fly toxicity bioassay………… 43 2.2.1.3 Adul female fruitfly toxicity bioassay……….. 44 2.2.1.4 Repellence and oviposition deterrence effect bioassay……………………………………… 46 2.2.1.5 Data analysis………………………………… 47 2.2.2 Schizaphis graminum Rondani, 1852………………….. 48 2.2.2.1 Rearing of Schizaphis graminum……………. 48 2.2.2.2 Toxicity bioassay…………………………….. 48 2.2.2.3 Repellence bioassay………………………….. 49 2.2.2.4 Data analysis…………………………………. 49 2.2.3 Tribollium castaneum Herbst, 1797……………………. 50 2.2.3.1 Rearing of Tribolium castaneum…………….. 50 2.2.3.2 Impregnation bioassay……………………….. 50 2.2.3.3 Flour discs bioassay………………………….. 52 2.2.4 Drosophila melanogaster Meigen, 1830………………. 52 2.2.4.1 Rearing of Drosophila melanogaster………… 52

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2.2.4.2 Diet preparation………………………………. 52 2.2.4.3 Contact toxicity………………………………. 52 2.2.4.4 Data analysis…………………………………. 53 2.2.5 Spodoptera exigua Hubner, 1808……………………… 53 2.2.5.1 Rearing of Spodoptera exigua………………. 53 2.2.5.2 Diet preparation……………………………… 53 2.2.5.3 Contact toxicity……………………………… 54 2.2.5.4 Data analysis………………………………… 55 2.2.6 Acyrthosiphon pisum Harris, 1776…………………….. 55 2.2.6.1 Rearing of Acyrthosiphon pisum…………….. 55 2.2.6.2 Diet preparation……………………………… 55 2.2.6.3 Feeding toxicity bioassay……………………. 56 2.2.6.4 Data analysis…………………………………. 58 2.3 Isolation of active compounds from the selected plants responsible for pesticidal activity……………………………… 58 2.3.1 Daphne mucronata fractionation……………………… 58 2.3.2 Isodon rogusus fractionation………………………….. 59 2.3.3 Bioactivity of fractions of Dhaphne mucronata and Isodon rugosus………………………………………… 60 2.3.4 Sub-fractionation of 500 mg butanol fraction of I. rugous through first reverse phase automatic flash chromatography……………………………………….. 60 2.3.4.1 Bioactivity of sub fractions of butanol fraction from first reverse phase automatic flash chromatography……………………………… 62 2.3.5 Sub-fractionation of most active fraction 3A from first reverse phase automatic flash chromatography through preparative high performance liquid chromatography

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(prep-LC)……………………………………………… 62 2.3.5.1 Bioactivity of sub-fractions collected from prep-LC……………………………………… 62 2.3.5.2 Spectroscopic analysis of prep-LC most bioactive fraction, 3A-3……………………… 63 2.3.6 Sub-fractionation of 5 g butanol fraction of I. rugous through second reverse phase automatic flash chromatography……………………………………….. 63 2.3.6.1 Bioactivity of sub fractions of butanol fraction from second reverse phase automatic flash chromatography……………………………… 64 2.3.7 Acidic extraction of most bioactive fraction 1B from second reverse phase automatic flash chromatography… 65 2.3.7.1 Bioactivity of ethyl acetate and water phase obtained through acidic extraction………….. 65 2.3.8 Identification of most bioactive compound, rosmarinic acid……………………………………………………. 65 2.3.8.1 HPLC-MS of isolated bioactive compound, rosmarinic acid……………………………… 66 2.3.8.2 Specific optical rotation of isolated bioactive compound, rosmarinic acid…………………. 66 2.3.8.3 1H and 13C NMR spectra of isolated bioactive compound, rosmarinic acid…………………. 66 2.4 Characterization of the isolated compounds for their pesticidal activity using different fractions……………………………….. 66 2.4.1 Bioactivity of isolated rosmarinic acid and commercial rosmarinic acid………………………………………… 66 2.4.2 Comparison between growth of live aphids exposed to

plant extract treated and untreated diet after 24 h of bioassay 67 2.5 Identification of genes involved in the synthesis of selected active compound, rosmarinic acid…………………………….. 67 2.5.1 Plant material…………………………………………. 67 2.5.2 RNA extraction……………………………………….. 67 2.5.2.1 Gel electrophoresis and quantification of RNA 68 2.5.3 cDNA synthesis………………………………………. 68 2.5.4 Target genes…………………………………………… 68 2.5.4.1 PCR amplification of rosmarinic acid synthase gene (RAS)………………………………….. 69 2.5.4.2 PCR amplification of hydrooxyphenylpyruvate reductase (HPPR)……………………………. 70 2.5.4.3 Gel electrophoresis………………………….. 71 2.5.5 3′ and 5′ RACE PCR for sHPPR (HPPR from I. rugosus) 71 2.5.5.1 3′ and 5′ RACE cDNA synthesis……………. 71 2.5.5.2 3′ and 5′ RACE PCR…………………………. 72 2.5.5.3 Gel electrophoresis…………………………... 73 2.5.6 Expression of HPPR in leaves, roots and stems of I. rugosus………………………………………………… 73 2.5.6.1 RNA extraction……………………………… 73 2.5.6.2 cDNA synthesis……………………………… 73 2.5.6.3 RNA and cDNA quantification……………… 74 2.5.6.4 PCR amplification…………………………… 74 3 Results……………………………………………………………….. 75 3.1 Pesticidal potential of selected plants against target pests…….. 76 3.1.1 Bactrocera zonata Saunders, 1842……………………. 76 3.1.1.1 Adult mortality……………………………… 76

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3.1.1.2 Effect of treatments on settlement and repellence behavior of females……………… 77 3.1.1.3 Oviposition deterrence………………………. 78 3.1.1.3.1 Effect of plant extracts on recovery of pupae…………………………. 78 3.1.1.3.2 Effect of plant extracts on adult emergence………………………. 78 3.1.2 Schizaphis graminum Rondani, 1852………………….. 79 3.1.2.1 Toxicity bioassay……………………………. 79 3.1.2.2 Repellence bioassay…………………………. 81 3.1.3 Tribollium castaneum Herbst, 1797…………………… 83 3.1.3.1 Impregnation bioassay………………………. 83 3.1.3.2 Flour discs bioassay…………………………. 83 3.1.4 Drosophila melanogaster Meigen, 1830……………… 83 3.1.4.1 Contact toxicity……………………………… 83 3.1.5 Spodoptera exigua Hubner, 1808……………………... 84 3.1.5.1 Contact toxicity……………………………… 84 3.1.6 Acyrthosiphon pisum Harris, 1776…………………….. 85 3.1.6.1 Feeding toxicity bioassay……………………. 85 3.2 Isolation of active compounds from the selected plants responsible for pesticidal activity………………………………. 88 3.2.1 Daphne mucronata fractionation………………………. 88 3.2.1.1 Bioactivity of fractions………………………. 88 3.2.2 Isodon rogusus fractionation…………………………… 90 3.2.2.1 Bioactivity of fractions……………………….. 90 3.2.3 First reverse phase automatic flash chromatography of 500 mg butanol fraction of I. rugosus………………….. 92 3.2.3.1 Bioactivity of sub-fractions of butanol fraction

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of I. rugosus………………………………….. 94 3.2.4 Sub-fractionation of 3A fraction through prep-LC 96 3.2.4.1 Bioactivity of sub-fractions of fraction 3A collected through prep-LC…………………… 96 3.2.4.2 Spectroscopic analysis of fraction 3A-3 98 3.2.5 Sub-fractionation of 5 g butanol fraction of I. rugosus through second reverse phase automatic flash chromatography……………………………………….. 100 3.2.5.1 Bioactivity of sub fractions of butanol fraction from second reverse phase automatic flash chromatography…………………………….. 101 3.2.6 Acidic extraction of most bioactive fraction 1B from second reverse phase automatic flash chromatography… 103 3.2.6.1 Bioactivity of ethyl acetate and water phase of acidic extraction……………………………… 103 3.2.7 Identification of most active bioactive compound……... 105 3.2.7.1 HPLC-MS…………………………………….. 105 3.2.7.2 1H and 13C NMR……………………………… 106 3.3 Characterization of the I. rugosus rosmarinic acid for their pesticidal activity using different fractions……………………... 107 3.3.1 Bioactivity of I. rugosus rosmarinic acid and commercial rosmarinic acid…………………………………………. 107 3.3.2 Comparison between growth of live aphids exposed to plant extract treated and untreated diet after 24 h of bioassay………………………………………………… 109 3.4 Identification of genes involved in the synthesis of rosmarinic acid in Isodon rugosus……………………………………………….. 110 3.4.1 PCR amplification of rosmarinic acid synthase gene 111

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(RAS)…………………………………………………… 3.4.1.1 Amplification with first set of primers……….. 111 3.4.1.2 Amplification with second set of primers……. 111 3.4.1.3 Amplification with third set of primers………. 112 3.4.2 PCR amplification of hydrooxyphenylpyruvate reductase (HPPR)………………………………………………... 113 3.4.2.1 sHPPR sequence (HPPR from I. rugosus)…… 113 3.4.2.2 sHPPR phylogenetic tree…………………….. 114 3.4.2.3 Alignment of amplified sHPPR with the conserved region amplified from other reported plant species………………………………….. 115 3.4.3 3′ and 5′ RACE PCR for sHPPR………………………… 116 3.4.3.1 3′ RACE PCR product sequence for sHPPR…. 116 3.4.3.2 5′ RACE PCR product sequence for sHPPR…. 117 3.4.4 Expression of HPPR in leaves, roots and stems of I. rugosus…………………………………………………. 117 4 Discussion……………………………………………………………. 118 4.1 Pesticidal potential of selected plants against target pest……… 119 4.1.1 Bactrocera zonata Saunders, 1842……………………. 119 4.1.2 Schizaphis graminum Rondani, 1852…………………. 122 4.1.3 Tribollium castaneum Herbst, 1797…………………… 124 4.1.4 Drosophila melanogaster Meigen, 1830………………. 125 4.1.5 Spodoptera exigua Hubner, 1808……………………… 126 4.1.6 Acyrthosiphon pisum Harris, 1776…………………….. 127 4.2 Isolation of active compound (rosmarinic acid) from butanol fraction of Isodon rugosus……………………………………… 129 4.3 Active principle, rosmarinic acid………………………………. 130 4.4 Identification of genes involved in the synthesis of rosmarinic

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acid in Isodon rugosus………………………………………… 132 4.5 Conclusion…………………………………………………….. 136 4.6 Recommendations…………………………………………….. 137 5 References………………………………………………………….. 138

LIST OF FIGURES

Fig. 1.1 Pyrethrin I, R = CH3, Pyrethrin II, R = CO2CH3……………………….. 12 Fig. 1.2 Rotenone………………………………………………………………... 13 Fig. 1.3 Nicotine…………………………………………………………………. 14 Fig. 1.4 Sabadilla alkaloids (a) Cevadine (b) Veratridine………………………. 15 Fig. 1.5 Ryanodine………………………………………………………………. 15 Fig. 1.6 Linalool…………………………………………………………………. 16 Fig. 1.7 Azadirachtin A………………………………………………………….. 17 Fig. 1.8 Boenninghausenia albiflora………………………………………………….. 21 Fig. 1.9 Calotropis procera…………………………………………………….. 22 Fig. 1.10 Tagetes minuta…………………………………………………………. 23 Fig. 1.11 Cinnamomum camphora……………………………………………….. 24 Fig. 1.12 Eucalyptus sideroxylon………………………………………………… 25 Fig. 1.13 Daphne mucronata…………………………………………………….. 26 Fig. 1.14 Isodon rugosus…………………………………………………………. 27 Fig. 1.15 Bactrocera zonata……………………………………………………… 30 Fig. 1.16 Drosophila melanogaster……………………………………………… 31 Fig. 1.17 Tribolium castaneum…………………………………………………... 32 Fig. 1.18 Spodoptera exigua……………………………………………………... 33 Fig. 1.19 Schizaphis graminum………………………………………………….. 34 Fig. 1.20 Acyrthosiphon pisum…………………………………………………… 35 Fig. 2.1 Adult male fruit fly toxicity bioassay………………………………….. 44 Fig. 2.2 Adult female fruit fly toxicity bioassay………………………………... 45 Fig. 2.3 Repellence and oviposition deterrence effect bioassay………………… 47 Fig. 2.4 Shizaphis graminum repellence behavior………………………………. 49 Fig. 2.5 Impregnation bioassay against Tribolium castaneum………………….. 51 Fig. 2.6 Contact toxicity against Drosophila melanogaster…………………….. 53 Fig. 2.7 Contact toxicity against Spodoptera exigua……………………………. 55 Fig. 2.8 Feeding bioassay against A. pisum……………………………………… 57 Fig. 2.9 Daphne mucronata fractionation……………………………………..... 59 Fig. 2.10 Four different solvent fractions of Isodon rugosus…………………….. 60 Fig. 2.11 Biosynthetic pathway of isolated pure compound, rosmarinic acid……. 69 Fig.3.1 Mean percent repellence (%) against fruit flies, Bactrocera zonata…… 78 Fig.3.2 UV chromatogram of reverse phase automatic flash chromatography… 94 Fig. 3.3 Prep-LC chromatograms of three sub-fractions of fraction 3A………… 99 Fig. 3.4 UV chromatogram of second reverse phase automatic flash…………… 100 Fig. 3.5 Mass spectra of isolated rosmarinic acid and commercial rosmarinicacid with a pseudo molecular ion at m/z value of 106 359…………………..

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Fig. 3.6 Structure of rosmarinic acid isolated from I. rugosus...... 107 Fig. 3.7 Comparison between growth of live aphids exposed to plant extracttreated and untreated diet after 24 h of 110 bioassay……………………… Fig. 3.8 Multiple bands appearence for RAS…………………………………... 111 Fig. 3.9 Purified band for RAS at 786 bp………………………………………. 112 Fig. 3.10 PCR products (multiple bands) for RAS in gradient PCR using 10annealing temperatures between 50◦C to 112 60◦C………………………… Fig. 3.11 Purified PCR product for sHPPR at 438 bp from I. rugosus………….. 113 Fig. 3.12 Phylogenetic tree for sHPPR (HPPR from I. rugosus) with HPPR in other plant species; Perilla frutescens, Solenostemon scutellarioides, 114 Salvia officinalis and Salvia miltiorrhiza……………………………… Fig. 3.13 Purified 5′ and 3′ RACE PCR products for sHPPR…………………… 116 Fig. 3.14 PCR products of HPPR and actin from I. rugosus leaves, roots and stems……………………………………………………………………. 117

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LIST OF TABLES

Table 2.1 Details of plant species used to evaluate pesticidal potential………… 40 Table 2.2 Yields of plant’s crude methanolic extracts………………………….. 41 Table 2.3 Details of target insects used for study……………………………….. 42 Table 2.4 First reverse phase automatic flash chromatography conditions of butanol fraction of Isodon rugosus……………………………….... 61 Table 2.5 Second reverse phase automatic flash chromatography conditions of butanol fraction of Isodon rugosus………………………………...... 64 Table 2.6 Primers used in amplification of rosmarinic acid synthase gene…… 70 Table 2.7 Primers used in amplification of hydrooxyphenylpyruvate reductase.. 71 Table 2.8 sHPPR gene specific primers for 3’ and 5’ RACE PCR amplification 72 Table 2.9 Actin and HPPR primers used in PCR amplification of HPPR and actin gene from roots, leaves and stem of Isodon rugosus…………… 74 Table 3.1 Mean percentage mortality of males and females of Bactrocera zonata exposed to methanolic extracts of different plants in female protein baits and male lures under laboratory conditions…………….. 77 Table 3.2 Mean number of Bactrocera zonata pupae recovered and adults emerging for guava fruit treated with various plant extracts and exposed for oviposition for 48 h……………………………………… 79 Table 3.3 Time course mean percentage mortality of Schizaphis graminum exposed to methanolic extracts of different plants for 12, 24. 36 and 48 hunder laboratory conditions in toxicity bioassay……………...... 81 Table 3.4 Mean percentage repellence of Schizaphis graminum exposed to methanolic extracts of different plants for 12, 24, 36 and 48 h under laboratory conditions in repellence bioassay…………………………. 83 Table 3.5 Time course mean percentage mortality of Drosophila melanogaster exposed to methanolic extracts of different plants at 2% for 24, 48 and 72 h under laboratory conditions………………………………… 84 Table 3.6 Time course mean percentage mortality of Spodoptera exigua exposed to methanolic extracts of different plants for 24. 48 and 72 h under laboratory conditions...... 85 Table 3.7 Toxicity of crude methanolic extracts against newborn (<24 h old) nymphs of pea aphids (Acyrthosiphon pisum) after exposure for 24 h on artificial diet supplemented with different concentrations of crude extracts………………………………………………………………… 87 Table 3.8 Toxicity of Daphne mucronata solvent fractions against newborn (<24 h old) nymphs of the pea aphid (Acyrthosiphon pisum) after exposure for 24 h on artificial diet supplemented with different concentrations of solvent fractions...... 89

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Table 3.9 Toxicity of Isodon rugosus solvent fractions against newborn (<24 h old) nymphs of pea aphids (Acyrthosiphon pisum) after exposure for 24 h on artificial diet supplemented with different concentrations of solvent fractions………………………………………………………. 91 Table 3.10 Sub-fractions collected from first reverse phase automatic flash chromatography……………………………………………………….. 93 Table 3.11 Toxicity of sub fractions of butanol fraction from first reverse phase automatic flash chromatography against Newborn (< 24 h old) nymphs of pea aphids (Acyrthosiphon pisum) after exposure for 24 h on artificial diet supplemented with different concentrations of solvent fractions...... 95 Table 3.12 Toxicity of sub-fraction 3A against Newborn (<24 h old) nymphs of pea aphids (Acyrthosiphon pisum) after exposure for 24 h on artificial diet supplemented with different concentrations of solvent fractions.. 97 Table 3.13 Fractions from second reverse phase automatic flash chromatography 100 Table 3.14 Toxicity of sub fractions of butanol fraction from second reverse phasenautomatic flash chromatography against newborn (<24 h old) nymphs of pea aphids (Acyrthosiphon pisum) after exposure for 24 h on artificial diet supplemented with different concentrations of solvent fractions………………………………………………………. 102 Table 3.15 Toxicity of ethyl acetate and water phase of acidic extraction againstnewborn (<24 h old) nymphs of pea aphids (Acyrthosiphon pisum) after exposure for 24 h on artificial diet supplemented with different concentrations of solvent 104 fractions………………………………. Table 3.16 Toxicity of isolated rosmarinic acid and commercial rosmarinic acid against newborn (<24 h old) nymphs of pea aphids (Acyrthosiphon pisum) after exposure for 24 h on artificial diet supplemented with different concentrations of isolated rosmarinic acid and commercial rosmarinic acid………………………………………………………... 108

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LIST OF ABBREVIATIONS

µg Microgram µg/mL Micro gram per mililiter µL Microliter µM Micromole Ach Acetylcholine ANOVA Analysis of variance AW-IPM Area-wide basis-Integrated Pest Managment BAT Protein Bat Annihilation Technique bp Base pair Bt Baccilus thuringiensis ChE Cholinesterase CI Confidence interval cm Centimeter d Days DDT Dichlorodiphenyltrichloroethane DDVP 2, 2-dichlorovinyl dimethyl phosphate EPA Environmental Protection Agency EtOAc Ethyl acetate EU European Union FAO Food and Agriculture Organization FDI Feeding deterrent index g Gram g/mol Gram per mole GF 120 Commercial protein bait containing Spinosad h Hour HF Heterogeneity factor HPLC-MS Preparative High performance liquid chromatography-mass spectrometry HPPR Hydroxyphenylpyruvate reductase IPM Integrated Pest Managment IR-RA Isodon rugosus rosmarinic acid KT50 Lowest knockdown effect at 50% LC Lethal Dose LC50 Lethal concentration causes 50% mortality LC90 Lethal concentration causes 90% mortality LD50 Lethal dose causes 50% mortality MAT Male Annihilation Technique ME methyl eugenol mg Miligram mg/L Miligram per liter mg/mL Miligram per milliliter MHz Mega hertz min Minutes mL Mililiter

xxv mM Milimole NMR Nuclear Magnetic Resonance PCR Polymerase chain reaction PIPs Plant incorporated protectants Ppm Parts per million Prep-LC Preparative high performance liquid chromatography RA Rosmarinic acid RACE Rapid amplification of cDNA ends RAS Rosmarinic acid synthase RH Relative humidity RNA Ribonucleic acid RNAi RNA interference TE Triss EDTA TEPP Tetra ethyl pyrophosphate UPM universal primer mix US United States WHO World Health Organization μg/gm Microgram per gram

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Chapter 1

Introduction

The current world population is about 7.3 billion that is projected to be approximatly 9.1 billions in 2050. For the estimated increase of additional 2 billion people, more serious efforts will need to be realised and warrant increasing the food production sources. Apart from this, 795 million people in the world were facing the problem of severe under nourishment in 2014-2016 (Rosen et al., 2016). To overcome these problems, about 30% more food is required at the global level. Furthermore, in the last decades besides the demand for more food production there was also demand for sustainable agricultural production including environmental friendly, socially fair and economically favorable (United Nations, 2009). Out of 36.5% total average loss of production worldwide, losses caused by insects, diseases and weeds were 10.2%, 14% and 12.2% respectively (Dang et al., 2012). Including pests, diseases and weeds, around 67000 different crop’s pests were reported (Oerke et al., 1994). During previous decades maximum efforts were made to enhance the agricultural productivity by reducing the damage caused by insects, diseases and weeds. This resulted in increased utilization of pesticides. Due to the intensive use of synthetic pesticides since 1960’s, in Europe 70% and in USA 100% crop yield increased with improvement in plant varieties, crop nutrition and irrigation through the management of pests (Pretty, 2008). Pesticides played an important role in agricultural productivity but their excessive and steadily rising use cannot be neglected and consequently resulted in serious threats to the environment and consumers (Pavela, 2016).

1.1. Synthetic pesticides There are four major groups of synthetic pesticides that are important because of their broad spectrum effects, persistency and toxicity in the environment. These are organochlorines, organophosphates, carbamates and pyrethriods (Guleria and Tiku, 2009).

1.1.1. Organochlorines Organochlorines are divided into two main sub groups, dichloro diphenyl trichloro ethane (DDT) and chlorinated alicyclics with different mechanism of action. DDT acts on peripheral nervous system. After activation and depolarization of membrane, DDT stops the gate closure at the axon’s sodium channel (Coats, 1990).

Chlorinated cyclodienes consists of dieldrin, endrin, aldrin, heptachlor, endosulfan and chlordane. These pesticides inhibit chloride flow through the nerve by binding to GABA sites in the gamma aminobutyric acid chloride ionophore complex. These insecticides act by disrupting the nervous system of the insects, results in tremors formation malfunctioning and ultimately death (Coats, 1990).

These insecticides are insoluble, thus become persistent in the soil and ground water. Therefore they can bio-accumulate in the body tissues of vertebrates and invertebrates that can ultimately affect the top predators through entering into the food chain. Their low solubility and adsorption in lipids and particles result in years to decay (Brooks, 1974). Organochlorine pesticides have diverse half lives ranging from 1 year to 10 years (Wauchope et al., 1992; Mackay et al., 1997). Due to these reasons in the late 1990’s, the use of organochlorines was stopped in industrialized countries but still in use in developing countries (Guleria and Tiku, 2009).

1.1.2. Organophosphates During World War II, these insecticides were originated from the compounds like tetra ethyl pyrophosphate (TEPP) and parathion as nerve gases by the Germany. These nerve gases were used as weapons in the war. Parathion, methyl parathion, melathion, chlorpyrifos, dichlorvos, diazinon, phosmet, tetrachlorvinphos, azinphos-methyl, azamethiphos and fenitrothion included in this group are splendidly used among all the organophasphate pesticides (Maugh and Thomas, 2010). Among all of them melathion is mostly used in public areas, gardens, agriculture and for mosquito control (Bonner et al., 2007). These insecticides inhibit cholinesterase (ChE) that at the nerve synapse breaks down the acetylcholine (Ach). Inhibition of cholinesterase results in hyperactivity and insects paralysis that leads to the death (Guleria and Tiku, 2009).

Some of these are not persistent, thus not bio-accumulates in the body of organisms as well as in the environment (Guleria and Tiku, 2009). Organophosphate degrades rapidly than organochlorides, but they can cause acute toxicity to people who are at the risk of their excessive exposure. Their degradation varies with the temperature, pH, sunlight and microbial composition. As these pesticides irreversibly inhibit acetylcholine esterase activity, they can criticaly affect nerve function both in insects and

3 humans. They are fatal to fetus and young children even if in very low doses, by effecting brain development. They can enter into the body via food or by in contact to the skin or due to inhalation of mist. In 2001, EPA banned their use in residential areas, but in agriculture they are still in use on many crops (Goodman and Brenda, 2011).

1.1.3. Carbamates Carbamates are derived from carbamic acid, their use as insecticide began in 1950’s. Insecticides included in this group are aldicarb, carbaryl, carbofuran, fenobucarb, methomyl and oxamyl. These insecticides act by reversibly inactivating the actylcholinestaerse enzyme (Metcalf, 2005). Carbamyl was the first carbamate insecticide introduced in 1956, blocks acetyl choline receptors through its effects on cholinesterase that ultimately effect nerve transmission in insects. They are broad spectrum, have medium lethality and persistence. They have less ability of bio accumulation; therefore do not cause major damage to the environment (Kuhr and Dorough, 1976; Guleria and Tiku, 2009).

Mostly carbamates are highly lethal to Hymenoptera, to avoid exposure to parasitic wasps and foraging bees, precautionary measures must be taken. Main route of carbamate entry is either by ingestion or by inhalation or through dermal route. Ingestion and inhalation routs cause more toxicity than dermal rout. Mostly reported symptoms are muscles weakness, sweating, minor body discomfort and dizziness. At higher doses of exposure, headache, nausea, abdominal pain, salivation and diarrhea are prominent symptoms (Agarwal and Sharma, 2010).

1.1.4. Pyrethroids Pyrethroids are synthetic insecticide whose structure are based on the natural insecticide i.e. pyrethrum. They were introduced in 1960’s consisting of extensively used agriculture compounds i.e. tetramethrin, resmethrin, fenvalerate, permethrin and delta methrin (Guleria and Tiku, 2009). Pyrethroids consists of almost one thousands insecticides. Although they are analogs of natural pyrethrins but their synthesis involves high chemical modification that make them more lethal and less degradable. They are mainly classified into two groups namely Type I and Type II on the basis of their physical properties and toxic effects. Type I pyrethroides derived from pyrethrin that do

4 not contain cyano group, elicit tumors. Type II pyrethroids contain cyano group and exhibit salivation and chloreoathetosis. Other compounds like piperonyl sulfoxide, piperonyl butoxide and sesame are used in the formulation of pyrethroids to increase the efficiency that acts as synergists. Formulated pyrethroids are highly toxic due to presence of other inert substances (Agency for Toxic Substances and Disease Registry, 2003). Pyrethroid presence was justified in rain water as these insecticides come along the water (Beenakumari, 2007). Due to these effects precautions must be taken during their use. They are broad spectrum insecticides used against insects belonging to orders like Diptera, Coleoptera, Hemiptera, Lepidoptera, Orthoptera, Hymenoptera and Thysanoptera, but lethal to some natural enemies of the pest as well. They have low persistence and mammalian toxicity but have very toxic effects on fish and aquatic invertebrates (Elliot et al., 1978; Guleria and Tiku, 2009). In soil they are degraded aerobically in 3 to 96 days and anaerobically in 5 to 430 days (Laskowski, 2002).

1.1.5. Neonicotinoids

Neonicotinoids are derived from nicotine insecticides. Two scientists, Shell and Bayer in 1980s and 1990s respectively, worked on development of these pesticides (Kollmeyer et al., 1999). These pesticides include clothianidin, nitenpyram, acetamiprid, imidacloprid, nithiazine, thiamethoxam and thiacloprid. Among all of these widely used insecticide is imidacloprid (Yamamoto et al., 1999). These insecticides and their byproducts results in less toxicity to mammals as compared to carbamates and organophosphates (Tomizawa and Casida, 2005). Imidacloprid is efficient in case of sucking insects, soil insects, fleas and few chewing insects (Gervais et al., 2010). It is used to irrigate plants by dissolving it in the water (Adak et al., 2012).

Like nicotine these insecticides have the ability to bind cell at nicotinic acetylcholine receptors and generate a reaction by that cell. In case of mammals these receptors are found in the cells of both peripheral nervous system and central nervous systems, but in insects these receptors are found only in cells of central nervous system. Abnormality in activation of these receptors results in their blockage due to high levels of overstimulation and nervous stimulation, which ultimately cause paralysis and finally death (Yamamoto et al., 1999; Gervais et al., 2010).

1.1.6. Disadvantages of synthetic pesticides

The use of many of the above-mentioned synthetic pesticides has been banned in many countries due to their toxicity and persistence in the environment (Smith and Gangolli, 2002; Walker and Lynch, 2007) but still many developing countries are using these banned toxic pesticides (Stoytcheva, 2011). Each year almost 2.5 million tons of pesticides are applied on crops and because of their excessive use worldwide loss reaches to $100 billion annually (Koul et al., 2008). As estimated by World health organization (WHO), due to mishandling of pesticides, annually 3 million pesticide poisoning cases take place that results in more than 250000 deaths (Stoytcheva, 2011). Due to residual effect of pesticides on animal and human health, there are many epidemiological studies indicating the link between toxicology of pesticides and cancer development (Settimi et al., 1990; Wolff et al., 1993; Dewailly et al., 1994). Pesticides may results in cancer either by non genotoxic manner through process such as promotion, peroxisome abundance and hormonal imbalance or affect the genome and increase the proliferation of neoplastic cells (Hodgson and Levi, 1996; Stoytcheva, 2011).

Many of the insecticides have negative effects on reproductive, respiratory, renal and nervous systems of both men and women (Stoytcheva, 2011). Due to some similarities between nervous system of insects and mammals, many insecticides that designed to target nervous system of insects, have their lethal effects on nervous system of many mammals (Tanner and Longston, 1990).

Synthetic pesticides to control pests all around the world have negative impact on human and animal health. It destroy the environment, develop pesticide-resistant organisms, gives secondary pest revival, are toxic to non-target organisms and lead to soil, water and air contamination, destroy aquatic and bird life, and, in worst case, have a direct lethal effects to users of the pesticides (Stoytcheva, 2011; Naqqash et al., 2016). For instance more than 500 arthropod pest species are reported to have developed resistance to one or more pesticides (Hajek, 2004).

1.2. Alternative approaches to overcome pest’s losses Due to above mentioned problems associated with the application of synthetic pesticides, EU commission, 2009 made legislative changes that aimed to decrease the utilization of synthetic pesticides or to search for new alternatives of these risky insecticides. To overcome this problem currently many alternatives exists to control insects. Best potential alternative to synthetic chemical pesticides are biopesticides. Biopesticides could eliminate pest population and they are living organisms, their products including both plants and microbial products or their byproducts. For safer production of food and sustainable agriculture, there is more demand and acceptability for biopesticides. Biopesticides are significantly considered as safer alternative, eco friendly, biodegradable and target specific (Kamble et al., 2016).

There are three main groups of biopesticides:

a) Microbial pesticides b) Plant incorporated protectants (PIPs) c) Biochemical pesticides (Botanical pesticides)

1.2.1. Microbial pesticides Microbial pesticides include microorganisms i.e. bacteria, virus and fungi, that can control many kinds of pests. Each pesticide have specific active ingredient against specific target pest. Some of pesticides are active by out-competing the pests. Microbial pesticides have to be regularly monitored to make sure that they should not become harmful to humans and other non target organisms (Mazid et al., 2011).

1.2.1.1. Entomopathogenic fungi Entomopathogenic fungi act as mycoinsecticides against many different insect pests and have vital role in insect population regulation. These mycoinsecticides act by penetrating through the cuticle of host, reach hemolymph, where they produce toxins and utilize nutrients in the haemocoel in order to grow and to avoid insect immune responses (Hajeck and Leger, 1994). Entomopathogenic fungi have been applied in the form of mycelium or conidia that will be sporulates following application (Mazid et al., 2011).

1.2.1.2. Viral biopesticides There are about 1600 different kinds of viruses that infect 1100 species of different insect pests. Baculovirus that infects more than 100 different insects belongs to special class of viruses and constitute about 10% of all the virus pathogens. Baculovirus have rod shaped particles that have DNA. Many viruses have protein coat to make inclusion body. Protein coat can get dissolve in the midgut of insects due to alkaline nature and thus virus released from the inclusion bodies. These viruses combine with the epithelial cells of midgut and kill the host by multiplying there (Usta, 2013). The registered baculovirus pesticides increases steadily, there are more than fifty formulations till now. Some of them have the same formulation, used with different names in different countries. GVs, NPVs are commonly used pesticides but the pesticides originates from nucleopolyhedrosis virus are considered the major group of pesticides (Mazid et al., 2011). Viral pesticides are more expensive than chemical pesticides and infect only specific pests that can serve as their hosts. That is why they cannot be used to control many different pests. Furthermore their effect on insects is very slow and their formulations degrade rapidly in ultraviolet light under the sun (Usta, 2013).

1.2.1.3. Bacterial biopesticides Bacterial pesticides are commonly used and cheaper among microbial pesticides. It can infect specific individual species of the flies, butterflies, moths, mosquitoes and beetles. In order to be effective, bacterial pesticides must be in contact with the target organism and must be ingested by the pest. Mostly bacterial pathogens that infect insects belong to bacillus that are spore forming rod shaped bacteria live in the soil (Usta, 2013). Most commonly used bacterial pesticide is Bacillus thuringiensis. This bacterium has the mechanism to produce pesticidal toxins. Mostly these toxins are coded by cry genes (Schnepf et al., 1998). After its discovery in 1901, this bacterial pesticide is widely used to control many different pests of medicine, forestry and agriculture. Cry proteins that are produced during sporulation have pesticidal property. Until now more than one hundred pesticides based on B. thuringiensis have been produced that have control on dipteran, lepidopteran and coleopteran larvae. The genes code for cry proteins have been successfully transferred to other crops. The toxicity of cry proteins is induced due to the

8 osmotic cell lysis because of the development of ion channels or transmembrane pores (Roh et al., 2007).

1.2.2. Plant incorporated protectants (PIPs) To eliminate or reduce the destruction of crops by pest many plants have been genetically modified with genes encoding insecticidal toxins to kill the pests. The acceptance of genetically modified crops has increased in last eleven years. The genetically modified plants with Bacillus thuringiensis that produces δ-endotoxins were commercialized in 1996 in the US. These genetically modified crops that express insecticidal toxins confers protection against number of insects (Shelton et al., 2000). The toxicity of this toxin solely depends on the alkaline nature of the midgut of the environment; due to this property they cannot affect humans. This toxin target pests of various crops including cotton, potato, maize, tomato, rice and tobacco. This provides more protection by their presence and their effect in those parts of the plant where foliar spray cannot reach (Shelton et al., 2000). There are many strains of Baccilus thuringiensis that can produce different types of cry proteins; about 60 different cry proteins identification has been done. For example, Bt maize hybrids expressing Cry1Ab protein, some expressing Cry1Ac or Cry9C protein can provide protection against a serious pest of maize in Europe and North America that is European corn borer. Many other maize hybrids expressing Cry3Bb1 provides protection against corn rootworm complex. Cotton express Cry1Ac protein provides protection against cotton bollworm (Usta, 2013; Gómez et al., 2002).

1.2.3. Biochemical pesticides Biochemical pesticides include natural substances that can act as pesticides against number of insects e.g. plant extracts that can act as pesticide in non-toxic way. Biochemical pesticides also comprise of materials that have the ability to hinder the mating or growth, like plant growth regulators or materials that attract or repel the pests includes pheromones (Mazid et al., 2011).

Plant extracts or other plant materials as pesticides have some benefits over other biopesticides. Other biopesticides need special formulations to act as pesticides before used by farmers, whereas plant extracts can be used by local farmers without any special

9 formulation. Other biopesticides are specific against particular insects e.g. viral biopesticides can be used against only those insects that can serve as a host of that particular virus. Almost 16 insect species become resistant in laboratory analysis and in the field, Busseola fusca, Spodoptera frugiperda and Helicoverpa zea showed resistance to Bacillus thuringiensis (Tabashnik et al., 2009). Farmers need special knowledge to use these biopesticides except botanical pesticides. The best alternative approach is to utilize plant’s secondary metabolites that plants actually synthesized in their defense against pests and pathogens (Miresmailli and Isman, 2014).

1.3. Botanical pesticides Many plants have the property to evolve extensive range of physical and chemical defenses against insects by producing secondary metabolites e.g. alkaloids, terpenoids, polyphenols and phenols. By using various extraction methods e.g. simply macerate the plants in water, using different polarity organic solvents, different distillation procedures and supercritical fluid extraction, these metabolites can be isolated (Dubey, 2011). Active substances in some plants that are isolated using these methods become known as botanical pesticides (Regnault-Roger et al., 2012; Isman and Grieneisen, 2014; Pavela, 2015). As compared to synthetic pesticides, whose formulation is usually based on a single compound, crude plant extracts have more potential to deal with resistant population of insects because it is very difficult for the insects to develop resistance against complex mixture of compounds (crude extract). Another benefit of using crude plant extracts, farmers can easily use them. Especially in developing countries where farmers cannot afford synthetic pestides, they can use locally available plants against insects. By having the knowledge of bioactive compound in crude plant extract, formulations can be made in which bioactive compound typically range from 1 to 5% which is acceptable range for all chemical compounds with in crude extract mixture to apply against insects (Isman, 2008). The toxicity of a pesticide, however, will be determined by its chemical nature regardless of its source.

1.3.1. History of botanical pesticides The use of plants to protect against insects dates back to more than 3000 years in Europe, perhaps the history of using them is not known exactly. People initially used

10 various modified parts of many aromatic plants as insect repellents against many damaging insects including ectoparasites or anthelmintics (Isman, 2006; Pavela, 2016). Many people used them to gain protection of stored products against insects (Isman, 2006; Grzywacz et al., 2014). The best known example that people used historically is finally ground chrysanthemum that gave protection against louses and fleas at that times. Some studies report that in 400 B.C children were deloused by using a dried flowers powder of plant known as pyrethrum (Abd El Ghany, 2012). Aromatic plants e.g. juniper, rosemary, myrrh were used to get fumigation of granaries in Ancient Rome. Due to these results people came to know the repellent effect of aromatic plants (Dubey, 2011). Due to use of many plants at that time the people came to know the use of baits prepared from Helleborus niger roots against rodents. In Persia, many oils from plants were used to treat scabies caused by some mites i.e. Sarcoptes scabiei (El-Wakeil, 2013). Later on with the agriculture development some plants were used to gain protection against phytophagous pests. In 17th century first commercial botanical insecticide, nicotine that gave protection against plum beetle was isolated from tobacco leaves. Later new insecticide from plant timbo-Derris spp was introduced in around 1850 (Abd El Ghany, 2012). But after world war II further development in botanical pesticides stopped in Europe due to advent of cheap synthetic insectides (Ware and Whitacre, 2004) and on other continents, the use of botanical pesticide has been preserved to a limited level today (Pavela, 2016).

1.3.2. Types of botanical pesticides Botanical pesticides can be classified into two groups:

I. This group includes botanical pesticides that are not produced commercially but people made them on the basis of previously available traditional recipes. People use the knowledge about pesticidal potential of some plants from generation to generation (Grzywacz et al., 2014; Sola et al., 2014). It is therefore difficult to know exact amount of plant material that is used in the preparation of insecticides. However to get information regarding pesticidal potential of local flora some ethanobotanical studies have tried to trace back

the information about their use in protection against insects (Asase et al., 2005; Odalo et al., 2005). II. This group includes botanicals that are produce commercially by small companies of local value. Until today various products are produced based on active substances in plants.

1.3.2.1. Pyrethrins (Pyrethrum/Pyrenone) Pyrethrum extracted as dried powder from Chrysanthemum cinerariaefolium (Asteraceae) (Fig. 1.1). Pyrethrum is the crude flower extract whereas pyrethrins are six insecticidal compounds isolated from this crude flower extract. Pyrethrins act by rapid knockdown in flying insects and convulsions and hypersensitivity in insects. In pure form pyrethrins are moderately toxic to mammals with oral LD50 of 350 to 500 mg/kg (Casida and Quistad, 1995). Pyrethrum as predominant botanical contributes 80% of the total botanicals in the market (Isman, 2005, 2006).

As they are less toxic to mammals and rapidly photo degraded, they are favorable to use in case of vegetables and dairies that are near to harvest, where less mammalian toxicity and no residual activity is in demand (El-Wakeil, 2013). It is toxic to bees and fish (Guleria and Tiku, 2009). Pyrethrum has been tested and found effective against caterpillars, aphids, leafhoppers, spider mites, bugs, cabbage worms and beetles (Casida, 1973; Glynne Jones, 2001).

Figure 1.1: Pyrethrin I, R = CH3, Pyrethrin II, R = CO2CH3 (Pavela, 2016)

1.3.2.1.1. Mode of action Pyrethrins affect central nervous system by destroying the potassium and sodium ion exchange in insects, and disturbing the normal transmission of nerve impulses. They cause paralysisrapidly in insects by acting after immediate application. However insects have enzymes in their guts that can metabolize pyrethrin and insects recover paralysis and not die after some time. Due to this reason pyrethrin are mixed with piperonyl butoxide or PBO that protect pyrethrin from metabolizing enzymes (Guleria and Tiku, 2009; El-Wakeil, 2013).

1.3.2.2. Rotenone Rotenone is the most lethal among all the botanical pesticides, isolated from the roots of Derris and Lonchocarpus (Fig. 1.2). It is very poisonous to aquatic life and fish, hence used for fish poisoning in water management programs. It has LD50 of almost 350 mg/kg. It is degraded rapidly in sunlight and air. Rotenone has more toxicity towards mammals through inhalation rather than by ingestion, skin contact results in irritation and inflammation of mucous membranes (Guleria and Tiku, 2009). Rotenone was reported to be active against bugs, aphids, potato beetles, spider mites and carpenter ants (Cabras et al., 2002; Cabizza et al., 2004).

Figure 1.2: Rotenone (El-Wakeil, 2013)

1.3.2.2.1. Mode of action Rotenone inhibits cellular respiration. It primarily acts on nerve and muscle cells and stop insects from feeding. It blocks the biochemical process at cellular level to stop

13 getting oxygen by insects required for body functions and therefore results in transmission of nerve impulses (Hollingworth et al., 1994; El-Wakeil, 2013).

1.3.2.3. Nicotine Nicotine is highly toxic to mammals, extracted from stems and leaves of tobacco plant and found commonly as nicotine sulfate (Fig. 1.3.). It usually enters through skin contact, eyes and mucous membrane. It is extremely lethal to warm blooded animals as well as for insects. It has oral LD50 of around 50 mg/kg (Isman, 2006). It is reported to be effective against aphids, thrips and caterpillars (Casanova et al., 2002).

Figure 1.3: Nicotine (Pavela, 2016)

1.3.2.3.1. Mode of action It acts by lowering the heart beat at elevated dose level but increasing the heart beat at small doses through interacting with nervous system. Death in insects occurred due to respiratory failure through improper functioning of chest muscles (El-Wakeil, 2013).

1.3.2.4. Sabadilla Sabadilla extracted from ripe seeds of Schoenocaulon officinale. In air and sunlight it breaks down rapidly with no residual effect (Fig. 1.4). It is broad spectrum insecticide acts mainly through contact poisoning and has some activity by acting as stomach poison. Sabadilla oral LD50 is around 5000 mg/kg. To human sabadilla cause irritation to skin, respiratory track and can be absorbed through skin contact. It has mainly cevadine and veratridine alkaloids that have effect as nerve poison (Guleria and Tiku, 2009). Sabadilla was analyzed against grasshoppers, codling moths, armyworms, aphids, cabbage loopers and squash bugs (Bloomquist, 1996, 2003).

Figure 1.4: Sabadilla alkaloids (a) Cevadine (b) Veratridine (El-Wakeil, 2013)

1.3.2.4.1. Mode of action Alkaloids in sabadilla acts on cell membrane, resulting in the loss of nerve function, ultimetly causing paralysis and finally death. Sabdilla kills insects immediately, if survive they will be in the state of paralysis for many days and then die (El-Wakeil, 2013).

1.3.2.5. Ryania Ryania has been extracted from roots of Ryania speciosa. Ryania shows low level of toxic effects to mammals and breaks down gradually (Fig. 1.5). The LD50 for ryania is around 750 mg/kg (Guleria and Tiku, 2009). Ryania was reported to be effective against codling moths, potato aphids, onion thrips, corn earworms and silkworms (Copping and Menn, 2000; Isman, 2006).

Figure 1.5: Ryanodine (El-Wakeil, 2013)

1.3.2.5.1. Mode of action Ryania acts as stomach poison but not produce rapid effect. It can slowly paralyze the insects and insect does not stop feeding immediately after ingesting it (El- Wakeil, 2013).

1.3.2.6. Limonene

Limonene has been extracted from citrus oils (Fig. 1.6). It has LD50 of more than 5000 mg/kg. Closely related compound linalool also isolated from orange and other citrus fruits peel. These citrus oils combined with insecticidal soaps to use against aphids and mites as contact poison. As they evaporate quickly from the surface, they have no residual effects (Guleria and Tiku, 2009).

Figure 1.6: Linalool (Pavela, 2016)

1.3.2.6.1. Mode of action Both limonene and linalool act as contact poison (Guleria and Tiku, 2009). It is thought that limonene cause sudden increase in sensory nerves activity. Central nervous system has been affected that result in motor nerves stimulation, ultimatly cause rapid knockdown paralysis (Leslie, 1994).

1.3.2.7. Azadirachtin Neem has important insecticidal compound, azadirachtin (Fig. 1.7). It is liminoid or tretranor triterpenoid compound (Isman, 2005). Due to its complex chemical structure, it cannot be synthesized. Bioactivity of neem products was reported against armyworms, cutworms, stemborers, bollworms, leaf miners, caterpillars, aphids, whiteflies, leafhoppers, psyllids, scales, mites and thrips (Dimetry et al., 1993, 2010).

Figure 1.7: Azadirachtin A (Pavela, 2016)

1.3.2.7.1. Mode of action Neem powerfully acts as feeding and oviposition deterrent and insect growth regulator. It is structurally like insect hormone, ecdysone. During insect growth and molting, azadirachtin interfere with production and reception of this hormone. It inhibit the molting cycle of insects, results in their death (Mordue and Blackwell, 1993; Guleria and Tiku, 2009).

1.3.3. Modes of application of botanical pesticides The modes of application of botanical pesticides can be described in terms of fumigation toxicity, contact toxicity, repellent activity, oviposition and adult emergence inhibition behavior and feeding deterrents.

1.3.3.1. Fumigation toxicity Botanicals applied as vapor or gaseous phase on target insects called fumigants (Rajendran and Sriranjini, 2008). Fumigation has been applied mainly to protect stored food against insect’s attack. Both invitro and invivo fumigation techniques has been applied by different methods by different workers. Most commonly used methods were impregnated paper assay in which treated filter papers were placed inside the containers (Abd ElSalam, 2010a, b; Aboua et al., 2010). During fumigation most of the botanicals acts by disturbing the function of neuromodulator octapamine and results in stopping the nervous system to work properly (Kostyukovsky et al., 2002). Some studies report the

17 blocking of nerve impulse transmission through inhibiting the acetylcholinesterase enzyme activity (Houghton et al., 2006), results in paralysis and death. For fumigation purpose mostly, many plant’s essential oils were used due to their volatile nature and they enter quickly through the respiratory system of insects (Regnault-Roger and Hamraoui, 1995).

1.3.3.2. Contact toxicity In contact toxicity botanicals exert their effect on contact with insects. To analyze contact toxicity different types of plant products e.g. powders, essential oils, extracts and seed oils were used by different workers against different insects. Methods used for contact toxicity includes residual film assay, impregnated paper assay, direct topical application and dipping method. Botanicals on contact may blocks spiracles and insects die due to asphyxiation (Denloye, 2010) or may penetrate through respiratory system (Ofuya and Dawodu, 2002). Botanicals if apply in fine particles on insects, it increases contact due to even distribution on the body of insects and insects may also die due to dehydration (Kedia et al., 2015).

1.3.3.3. Repellent activity Botanicals acts as repellent through deterring insects from landing over or flying to food materials, either by acting locally or from a distance (Nerio et al., 2010). Hundreds of the plants have been analyzed as repellents in many studies in last 50 years (Sukumar et al., 1991). Repellency has been analyzed either through filter paper method (Talukder and Howse, 1994) or through olfactometer assay (Shukla et al., 2011).

1.3.3.4. Oviposition deterrent and adult emergence inhibition behavior Botanicals act as oviposition deterrent by inhibiting females to lay eggs on the food materials. They are reported to effects adversely ovariole of female insects (Dodia et al., 2008). Oviposition deterrence may occur due to dying of females or due to failure of females to lay eggs on contact with botanical products (Shukla et al., 2011). Egg lying capacity of insects may deter due to changes in their physiology and behavior. These plants chemicals are known as semiochemicals and have important role in integrated pest management instead of using toxic chemicals (Kumar et al., 2009). Plant products may penetrate through chorion into the eggs and interfere with the physiological and

18 biochemical processes, results in suppression of embryonic development (Raja et al., 2001). Adult number may also reduce due to low hatchability of eggs, and failure of hatching may result due to mortality of eggs or may be due to changes in surface tension and oxygen tension within the eggs (Abdullahi et al., 2011). Oviposition deterrence stops the spreading of insect’s population at the initial stages of their life cycle.

1.3.3.5. Feeding deterrents Botanical products act as feeding deterrent by stopping the insects from feeding and they die due to starvation. Some plant products effect peristaltic movements and insects die due to starvation. Neem products due to presence of azadirachtin cause antiperistaltic effect in the alimentary canal of many insects (Immaraju, 1998). A number of studies reported the deterrent effect of plant powders, essential oils and extracts as feeding deterrent index (FDI) (Jayakumar, 2010). Plant products that have the potential to act as feeding deterrent can cause high adult mortality, oviposition deterrence effect, high mortality of eggs and low adult emergence (Kumar et al., 2008; Lale and Mustapha 2000; Raja et al., 2001).

1.3.4. Overview on plant products and extracts used as botanical pesticides Over the years, from 235 plant families, more than 6000 plant species have been screened for their pesticidal potential against many pests (Copping and Menn, 2000; Walia and Koul, 2008). Plants were applied as crude extracts, as powders and as aqueous or organic solvent extracts against insects (George et al., 2008). For primary health care and pest control, 80% of the population belongs to developing countries rely on traditional control measures as estimated by WHO survey. For the development of potential products, botanicals are needed to be explored and screened (Pandey and Dushyant, 2011). Botanicals have low or no residual property and thus readily break down in the soil, thus not bio accumulate in animal and plant tissues. The parts of plants used for the preparation of insecticides include roots, leaves, tubers, stems, seeds, flowers, pods, sap and wood. Mostly leaves (62 species) were utilized followed by roots (16 species) and tubers (12 species) in bioassays against insects. Different studies were reported that the plants use for insecticidal activity belong to families i.e. Asteraceae, Annonaceae, Asclepiadaceae, Fabaceae and Euphorbiacea (Boulogne and Petit, 2012).

Several plants reported to have insecticidal activity, inlcuding neem, pongamia, adathoda, chrysanthemum, turmeric, Indian privet, onion, garlic, deodarcedar, tobacco, ocimum, custard apple, ginger, citrus fruits and many other plants (Rahuman et al., 2009; Osipitan and Oseyemi, 2012). Spinosad produced due to fermentation of fungus and azadirachtin isolated from neem tree, have insecticidal potential against fall webworm Hyphantria cunea (Brudea et al., 2012). Garlic has been grown as border inter crop to protect the main crop from pest infestation due to its repellent activity against insects. Garlic and onion bulbs are used as extract or as powdered form against insects in the fields and granaries. Plants including Vitex negundo, Pongamia glabra, Adathoda vasica and Acorus calamus were reported to protect stored food against pests (Sadek, 2003). Similarly, the effective control against early third instar larvae of Anopheles stephensi, malaria mosquito has been provided by using extracts of Pomoea cornea fistulosa, Calotropis gigantea and Datura strumarium that contained active principles (Arivoli et al., 2012). Flower extracts of Michelia champaca showed potential control against mosquito larvae (Boulogne et al., 2012). Similarly, Strychnos nuxvomica leaf extract had the larvicidal potency to control Culex quinquefascaitus (Arivoli and Samuel, 2012). Leaf extracts of many plants including Lantana camara, Ocimum basilicum, O. sanctum and Vetivera zizanoides were reported to be useful in providing protection against leaf miners in potato, brinjal, beans, chillies and tomato. Tagetes erecta crushed leaves protect against root-knot nematodes after applying to soil in mulberry gardens (Chitwood, 2002). According to some reports, Annona squamosa and Citrus paradise seed extracts have been useful to control diamond moth and Colorado potato beetle. Similarly, Spodoptera litura and Heliothis armigera were effectively controlled through the use of bark extract of Meliaazedarach that was acted as antifeedant (Wheeler and Isman, 2001; Nathan, 2006). Extracts from the leaves of Argemone Mexicana, Cymbopagon citrates, Artemesia absinthium, Cassia occidentalis and Sieges beckiia orientalis were reported to be strong antifeedants against Crocidolomia binotalis (Abdelgaleil et al., 2008). Growth of bacteria was inhibited by root extract of Moringa oleifera (Fahey, 2005). Extracts from the plants i.e. Garcinia kola, Zingiber officinale, Azadirachta indica and Allium sativum used to control Xanthomonas campestris, responsible for the bacterial leaf spot to protect two varieties of Solanum i.e. S. gilo and S. torvum (Opara and Obani, 2010).

1.4. Overview on selected plants

1.4.1. Boenninghausenia albiflora (Hook.) Rchb. ex Meisn. (Rutaceae) Boenninghausenia albiflora is known by common name as White Himalayan Rue and locally (Abbottabad region) as Pissu Mar Booti (Fig. 1.8). This plant is a perennial herb, woody at the base, commonly found in north of Pakistan and grow in shady forests.

Figure 1.8: Boenninghausenia albiflora (Pissu Mar Booti)

B. albifora is traditionally used to kill flea. This plant is reported to posses various insecticidal activities. An insecticidal coumarin, murraxocin (7-methoxy-8-[1′- ethoxy-2′-hydroxy-3′-methyl-but-3′enyl]-coumarin), isolated from the leaves of B. albifora and was analyzed and active against forest pests, Plecoptera reflexa, Clostera cupreata and Crypsiptya coclesalis (Sharma et al., 2006). Sharma et al.(2007) evaluated methanol and hexane extract against Spodoptera litura and have isolated compounds, 2- Octadecanone, palmitic acid, p-methoxy-methyl cinnamate and 3-(3í,4ídihydroxyphenyl) propyl hexadecanoate and two known coumarins, murralongin and albiflorin-3 as an antifeedent through bioactivity guided isolation procedure. Negi et al. (2012) evaluated plants, Boenninghausenia albiflora, Skimmia anquetillia, Glycosmis arborea, Vitex negundo, Premna barbata, Callicarpa arborea and Clerodendron indicum against Plecoptera reflexa, Clostera cupreta, Crypsiptya coclesalis and Spodoptera litura. They found that B. albifora and S. anquetillia showed potent insecticidal activities.

1.4.2. Calotropis procera Aiton (Dryand) (Apocynaceae) Calotropis procera is evergreen perennial shrub. It is commonly known as Sodom Apple and locally as Ak (Fig. 1.9). It belongs to flowering plants and is native to Tropical Africa, South Asia, North Africa, Western Asia and Indochina.

Figure 1.9: Calotropis procera (AK)

Begum et al. (2011) reported the insecticidal activity of ethanolic extracts of leaves from C. procera against Musca domestica in different concentrations and found that the C. procera showed LC50 value of 282.5 mg/L. Phytochemical analysis showed that alkaloids were found in the maximum amount. Chloroform, hexane, ethanol, methanol and ethyl acetate extracts from the leaves of C. procera were evaluated for their antifeedent activity against Spodotera litura and found that the choloroform extract had more antifeedent effect as compared to all other analyzed extracts (Bakavathiappan et al., 2012). Calotropis procera also showed pronounced insecticidal activity against Tribolium castaneum (Khan et al., 2012). Barati et al. (2013) evaluated the insecticidal activity of C. procera against Bemisia tabaci grown on tomato plants under controlled conditions in glass house. From leaves and latex of C. procera, important pharmaceutical compounds have been isolated like calotropon, uscharine, calotropin, calctin, calotropagenin and uscharidin (Chopra et al., 1956; Pathak et al., 2014). Pathak et al.

(2014) reported that C. procera has repellent effect against mite Eriophyes cheriani massee.

1.4.3. Tagetes minuta L. (Asteraceae) Tagetes minuta is an annually grown plant. It is commonly known as Mexican Marigold and locally as Gul-e-Sad Barg (Fig. 1.10). It is grown widely in tropical and temperate regions of the world.

Figure 1.10: Tagetes minuta (Gul-e-Sad Barg)

The aerial parts of T. minuta were analyzed for the insecticidal activity against sand fly, Phlebotomus duboscgi and significant mortalities were observed (Njeru, 2011). The leaf powder of Tagetes minuta was evaluated against stored grain pest, Sitophilus zeamais and was assesses after 14, 28, 42, 56 and 70 d (Muzemu et al., 2013). The aqueous and methanolic extracts of T. minuta were evaluated for their toxic and repellent effect against Tribolium castaneum. It was found that T. minuta showed strong repellent effect against T. castaneum (Padin et al., 2013). Phoofolo et al. (2013) reported the aphicidal effect of T. minuta crude extract from water, acetone, methanol and mixture of water, acetone and methanol against Brevicoryne brassicae and found that mixture extract showed more toxic effect than all the extracts evaluated. Ground leaf powder of T. minuta evaluated against Sitophilus zeamais to determine the effect of powder on percent weight loss, insect infestation, percent germination, grain odor and grain color up

23 to 192 days. They found that plant powder prevent only grain damage and insect infestation (Musundire et al., 2015). In one study methanolic extracts of T. minuta synergized with Sesamum indicum and one without synergizing, evaluated against Periplaneta americana and found that with contact toxicity and fumigation synergized extract had more effect than non synergized extract (Otengo et al., 2015). Motazedian et al. (2014) reported significant fumigant activity of essential oil from T. minuta against cabbage aphid, Brevicoryne brassicae and found limonene (13%) as a most abundant compound. Tagetes minuta essential oil depicted insecticidal activity against diamondback moth, Plutell xylostella and abundant compound observed was limonene (59.5%) (Reddy et al., 2015)

1.4.4. Cinnamomum camphora (L.) J. Presl (Lauraceae) Cinnamomum camphora is evergreen tall tree, native to China, Japan, Taiwan, Korea, Veitnam and many other countries (Fig.1.11). It is commonly known as Camphor tree and locally as Kafoor.

Figure 1.11: Cinnamomum camphora (Camphor or Kafoor)

Cinnamomin is ribosome inactivating protein isolated from seeds of C. camphora, evaluated against bollworm larvae through feeding bioassay, exhibiting LC50 of 1839 ppm (Zhou et al., 2000). Its toxicity was also observed against domestic silkworm through feeding bioassay. (Wei et al, 2004). Powders from the aerial parts of the C.

24 camphora were analyzed against Tribolium castaneum and Trogoderma granarium. Trogoderma granarium was more susceptible to C. camphora plant powders than T. castaneum (Nenaah and Ibrahim, 2011). Essential oil from C. camphora showed strong fumigant and contact toxicity against Tribolium castaneum and Lasioderma serricorne. GC-MS analysis revealed the presence of, 1,8-cineole (4.3%), α-terpineol (3.8%), D- camphor (28.1%) and linalool (22.9%) (Guo et al., 2016). Jiang et al. (2016) investigated the chemical composition, contact and repellent activities of essential oil from leaves, seeds and twigs of C. camphora against cotton aphid. The major components present in the leaves and twigs were eucalyptol, camphor, 3,7-dimethyl-1,3,7-octatriene and linalool and in seeds were methyleugenol (20%), eucalyptol (20.9%), camphor (5.5%) and linalool (14.7%). In case of contact toxicity, LC50 values were 275, 245.79 and 146.78 mg/L exhibited by twings, leaves and seeds, respectively. Seed essential oil gave highest repellent effect of 89.9% after 24 h of treatment.

1.4.5. Eucalyptus sideroxylon A. Cunn.ex Woolls (Myrtaceae) Eucalyptus sideroxylon is a medium to tall tree. It is commonly known as Red Ironbark and locally as Gond or Safeda (Fig. 1.12).

Figure 1.12: Eucalyptus sideroxylon (Gond or Safeda)

Fumigant and repellent activities of essential oil from E. sideroxylon were evaluated against first instar of Blattella germanica. The lowest knockdown effect at

50% KT50 for E. sideroxylon was 62 minutes but exhibited lower repellent effect as compared to positive control used in the experiment (Alzogaray et al., 2011). Tolozoa et al. (2010) evaluated the fumigant activity of E. sideroxylon essential oil against head louse, Pediculus Humanus capitis and observed that it has knockdown time, KT50 of 24.8 min.

1.4.6. Daphne mucronata Royle (Thymelaeaceae) Daphne mucronata is an evergreen shrub. It is commonly known as Kashmir Daphne and locally as Kutti lal (Fig. 1.13). It is native to Pakistan, specifically the northern areas of Pakistan.

Figure 1.13: Daphne mucronata (Kashmir Daphne; Kutti Lal) Insecticidal activities for this plant are not reported before in any previous study. It is well known medicinal plant and has been used for the treatment of many diseases. Daphne mucronata has been used to treat bone, skin diseases and allergies (Ali and Qaiser, 2009; Afzal et al., 2009; Hussain et al., 2012). It has been used to treat skeleto- muscular and rheumatism problems (Hamayun, 2007; Murad et al., 2011). It consists of important bioactive compounds including flavonoids (Ullubelen, 1986; Baba et al., 1986; Baba et al., 1987), coumarins (Baba et al., 1986), triterpenoids (Baba et al., 1986), cumarinolignans (Ullah et al., 1999; Rasool et al., 2009), lignin (Ullah et al., 1999), glucosides, aquillochin, daphecin, umbelliferone and daphnine (Rasool et al., 2010; Hussain et al., 2012). The ethanolic extracts from different parts of the plant depicted

26 antimicrobial activity against Pseudomonas aeruginosa, Escherichia coli, Bacillus subtilis and Staphylococcus aureus (Javidnia et al., 2003). Ethanolic extract exhibited promising breast anticancer activity (Hedayati et al., 2005) and long term used reported to completely eliminate the tumor (Hedayati et al., 2003). Daphne mucronata contains an important bioactive diterpene compound, gnidilatimonoein that was reported to be active against many cancer cell lines like HL-60, CCRF-CEM, K562, MOLT-4 leukemia cell lines, LNaP-FGC-10 prostate cancer cell lines and Wehi-164 a mouse BALB/C fibrosarcoma cell line (Yazdanparast and Sadeghi, 2004). Uysal et al. (2016) evaluated the antibacterial activity of essential oil from D. mucronata against Bacillus subtilis, Bacillus cereus, Staphylococcus epidermidis, Staphylococcus aureus, Escherichia coli, Streptococcus faecalis, Proteus mirabilis, Pseudomonas aeruginosa, Proteus vulgaris and Salmonella typhi. The main components in the oil were identified as nootkatin, nootkatone and daphnauranol C.

1.4.7. Isodon rugosus Wall.ex Benth (Labiatae) Isodon rugosus is a deciduous shrub. It is commonly known as Wrinkled Leaf Isodon and locally as Boi (Fig. 1.14). It is distributed in Afghanistan, Arabia, Pakistan, China and northern temperate regions of Himalaya.

Figure 1.14: Isodon rugosus (Boi)

Isodon rugosus has been known to treat many diseases but its insecticidal activities are not reported in any other previously reported study. Ethnobotanically, the

27 bark has been used to treat dysentery and cure generalized pain of the body (Shuaib and Khan, 2015). Dried leaves were used to treat toothache (Akhtar et al., 2013) and fresh leaves used to cure earache (Sabeen and Ahmad, 2009). Ethnomedically, it has been used to treat abdominal pain and gastric problems (Ahmad et al., 2014). It was reported to be used for different infections, pyrexia, blood pressure, rheumatism and microbial infections (Khan and Khatoon, 2007; Adnan et al., 2012; Shuaib et al., 2014). Isodon rugosus has been reported to be used pharmaceutically as bronchodilator, antidiaiarrheal and hyporglycaemic (Sheret al., 2011; Ajmal et al., 2012; Janbaz et al., 2014). Rauf et al. (2012b) reported the antibacterial activity and phytotoxic effect of crude methanolic and subsequent fractions of I. rugosus. As compared to antibacterial activity, I. rugosus showed significant phytotoxic effect. Phytotoxic and cytotoxic effect of crude methanolic extract and subsequent fractions were reported against seeds of radish and brine shrimp and found that crude extract was most phytotoxic while chloroform and saponin fractions gave more cytotoxic effect (Zeb et al., 2014a). Isodon rugosus crude extracts and its fractions were evaluated for their inhibitory activities of butyrylcholinesterase and acetylecholinestrase, oxidative stress and evaluated the total phenolic and flavonoid contents. Flavonoid and chloroform fractions gave the maximum inhibitory effect against butyrylcholinesterase and acetylecholinestrase. Fraction flavonoid, chloroform and ethylacetate gave highest antioxidant activity, while crude, ethylacetate and flavonoid showed highest amount of flavonoids and phenolic contents (Zeb et al., 2014b). Analgesic potential of crude and different fractions were evaluated and revealed that chlorofom fraction showed more analgesic effect. Most prominent components like sugiol, benzyl alcohol, myristic acid, phytol, sebacic acid, tocopherol, α- Amyrin, and stigmastero were isolated as analgesic compound through GC-MS analysis (Zeb et al., 2016).

1.5. Overview on targeted insects

1.5.1. Bactrocera zonata Saunders (Diptera) The peach fruit fly, Bactrocera zonata Saunders, 1842 (Diptera: Tephritidae), is one of the most economically important insect pests that causes economic loss by damaging fruit and by interfering international horticultural trade (Shehata et al., 2008)

(Fig. 1.15). It is native to Asia where it causes severe damage to over 50 species of fruit crop. Its most preferred host is guava, in which losses may reach 50%, if effective control measures are not adopted (Awad et al., 2014)

Like many other species in the genus Bactrocera, males of B. zonata show strong attraction to a phenylpropanoid compound, methyl eugenol (ME; 1, 2-dimethoxy-4-(2- propenyl) benzene), which occurs naturally in many plant species (Tan & Nishida, 2012). ME is used to monitor and suppress populations of fruit flies by a male lure and kill approach (male annihilation technique; MAT) (Steiner et al., 1970). The MAT relies on attracting males from the field population in devices containing ME and insecticides (Vargas et al., 2010). To suppress female populations in conjunction with MAT, sprays of protein baits containing insecticides (bait application technique; BAT) can also be used. As components of integrated pest management, both MAT and BAT are more efficient when used on an area-wide basis (AW-IMP). In addition to these control measures, farmers routinely apply synthetic insecticides. However, synthetic insecticide application is undesirable because of adverse effects to the environment, poisonous residues in fruit and issues for international trade (El-Aw et al., 2008).

Due to the limitations of each control strategy as a stand-alone technique, it is recommended that an IPM approach be adopted (Vargas et al., 2015). Although, MAT and BAT are components of IPM and these are environmentally benign techniques, overuse of baits with synthetic insecticides deposits a huge quantity of insecticides into the environment, so to protect the environment safer insecticides i.e. Spinosad are incorporated into baits. Therefore, exploring plant based insecticides may lead to the discovery of safer insecticides either for direct application or for incorporation into the baits. Plant extracts can potentially be eco-friendly alternatives to synthetic insecticides in IPM of fruit fly populations.

Figure 1.15: Bactrocera zonata

1.5.2. Drosophila melanogaster Meigen (Diptera) The common fruitfly, Drosophila melanogaster is a polyphagous pest, and in Pakistan it mostly damage pulpy fruits especially banana and guava (Fig. 1.16). D. melanogaster feed on pulp and thus make the fruit useless. Mostly in summer their population reached the maximum and thus cause more harm to fruits. With a short life cycle of 10 days from egg to adult and the possibility of many generations in one season, D. melanogaster, if not controlled, can rise to significantly high numbers with potential economic impact on fruit production and storage (Yasmin et al., 1995). It is also noxious pest in homes, fruit markets and restaurants. They feed on animal and human excrets, also on uncooked food and thus serve as serious disease carrier (Anjum et al., 2010). It is causative agent involved in spreading of pathogen, Staphylococcus aureus (Nedham et al., 2004). Survey done by FAO reported that pests and rodents collectively cause 10- 25% of world harvested food loss annually (Khan et al., 2015). In Pakistan, annual loss caused by fruit flies to growers reaches to about 7 million rupees. Therefore it is an important economic and health concern to control these flies.

For the control of these flies there are many reports in the literature (Khan et al., 1999). This pest faces a lot of insecticide application because of its cosmopolitan distribution, found in temperate and tropical regions and ability to occupy all type of environment (Fournier-Level et al., 2016). Due to lots of insecticide application this pest has developed resistance like towards DDT (Le Goff and Hilliou, 2016).

Figure 1.16: Drosophila melanogaster

1.5.3. Tribolium castaneum Herbst (Coleoptera) The red flour beetle, T. castaneum is worldwide pest that can cause damage to stored products including grains, chocolate, cereals, flour, pastas, beans and nuts in food warehouses, mills, urban homes and retail stores, and it can survived more than 3 years (Via, 1999; Weston and Rattlingourd, 2000; Rees, 2004) (Fig. 1.17). It can damage stored grains either through feeding or by reducing their quality severely through larval feces. At adult and larval stages it posed serious damage to flour and cereals, it also causes damage to seeds and make colour changes and bad odor to flour. Their long term presence on flour and cereal grains usually leads to mould development (Bennett, 2003; Baldwin and Fasulo, 2004).

Many chemical insecticides and fumigants has been used to control T. castaneum and other stored product pests, and control through chemicals need special care during the whole period of the storage and transportation (Story, 1984). Due to continuous use of chemical insecticides, T. castaneum become resistant worldwide (Dyte and Blackman, 1970; Champ and Dyte, 1976; Rossi et al., 2010; Singh and Prakash, 2013). The genus Tribolium become resistant to almost all chemical insecticides including lindane (Champ and Campbell-Brown, 1970), melathion (Champ and Campbell-Brown, 1970; Assie et al., 2007; Bughio and Wilkins, 2004), phosphine (Campbeel, 2010; Jagadeesan et al., 2012), cyclodiene (Andreev, 1999) and deltamethrin (Singh and Prakash, 2013).

Figure 1.17: Tribolium castaneum

1.5.4. Spodoptera exigua (Hubner) (Lepidoptera) The beet armyworm, Spodoptera exigua is a major and polyphagous pest, damaging major agriculture areas including field, flower and vegetable crops (Taylor et al., 2008) (Fig. 1.18). Due to its polyphagous ability, vast distribution and strong migratory properties (Stewart et al., 2002), it has the ability to attack vast areas of horticultural cultivations and field crops (Han et al., 2008). It has many host crops including sugar beet, tobacco, cotton, soy bean in Mediterranean and African countries (Mushtaq et al., 2008) and other hosts includes lettuce, cabbage, egg-plant, potato, pea, spinach, alfalfa and corn (Dingha et al., 2004; Rizwan- ul- Haq et al., 2009). Though it has irregular outbreaks, at late larval stages its population increases because of improper management. If not properly managed, its control becomes difficult because of its multivoltine behavior and short life span. Spodoptera exigua damages 60 species belonging to 31 families of weeds and farm plants (Talhouk, 2003).

Spodoptera exigua controlled strategy involves the use of chemical insecticides and their continuous use results in resistance development that made its control difficult (Van Laecke and Degheele, 1991). Development of resistance causes major problem for the management of S. exigua because the parts of the plant on which it usually attacks are repeatedly treated with chemical insecticides. The abundance of this insect is due to frequent use of insecticides and it is assumed to act as secondary or induced insect in many crops (Eveleens et al., 1973).

Figure 1.18: Spodoptera exigua

1.5.5. Schizaphis graminum Rondani (Hemiptera) Green bug aphid, Schizaphis graminum is dominant specie in Pakistan, having a wide host range of at least 60 plant species including wheat, sorghum, corn and barley (Bowling et al., 1998, Rustamani et al., 1999) (Fig. 1.19). Green bug aphid like other aphids consumes phloem sap as food by inserting its stylet into phloem sieve elements (Miles, 1999; Burd, 2002). It primarily develops red or necrotic spots on sorghum and wheat crops and eventually followed by general necrosis and plant death (Porter et al., 1997; Miles, 1999). Carbon assimilation rate, total chlorophyll and transpiration decreases significantly by the aphid that badly affects the nitrogen and protein contents, weight of 1000 grains, number of grains per year (Ciepiela, 1993) and reduces plant biomass ultimately (Holmes et al., 1991). By sucking sap aphids can cause 35-40% loss directly and by transmission of fungal and viral diseases cause 20-80% loss indirectly (Rossing et al., 1994).

To overcome Schizaphis graminum a number of methods have been investigated. To control these aphids, insecticides have been used on routine basis. Due to these control measures resistance to insecticides increases and many of them are toxic to non target insects (Van Emden and Harrington, 2007). These aphids have developed resistance to many of old insecticides including DDT, pyrethroids and parathion-methyl over the past 20-30 years (Gubran et al., 1992; Gao et al., 1992). This is most likely that these aphids can aquire resistance against modern pesticides.

Figure 1.19: Schizaphis graminum

1.5.6. Acyrthosiphon pisum Harris (Hemiptera) The pea aphid, Acyrthosiphon pisum (Hemiptera: Aphididae), is one of the most important groups of insect pests in the world (Fig. 1.20). About 4700 aphid species have been described up to date, out of which 250 species are serious pests of ornamental plants and various crops (Alford 2000; Blackman and Eastop 2007). The pea aphid causes direct damage by sucking the phloem of plants and indirect damage by transmitting plant pathogenic viruses and allowing the production of fungi on the excreted honey dew (Dixon 1998; Van Emden and Harrington 2007; The International Aphid Genomics Consortium 2010). A. pisum is responsible for crop damage worth hundreds of million dollars every year and is a vector of more than 30 viruses, including red clover vein mosaic virus, pea streak virus and bean yellow mosaic virus (Barnett and Diachun 1986; Jones and Proudlove 1991; Brault et al., 2010).

The use of synthetic broad spectrum insecticides to control A. pisum has led to the development of resistance, negative effects on non-target organisms, humans and the environment (Unal and Jepson 1991; Ware et al., 2003). The increased environmental awareness and the pollution potential and health hazards of many of the synthetic chemical pesticides, support the search for new natural insecticides to control pests in an environment friendly approach (Edwards et al., 2008).

Figure 1.20: Acyrthosiphon pisum

1.6. Theoretical underpinning Due to mounting hazards and cost of synthetic pesticides, natural products derived from plants provide the hope for sustainable pest management with reduced environmental and health hazards in the future. The potential plant insecticidal extracts when combined with other environmentally friendly approaches i.e. biological control, cultural practices and trap crops can provide effective control against many economically damaging insect pests to be applied in Integrated Pest Management Programs (IPM). As, plants give additive or synergistic effects of many strong and weak insecticidal activities, the purification and structure determination of the active principle is necessary to improve bioefficacy of active principles. From the knowledge of active principle in crude extracts, different formulations of crude extracts against insects can be made based on the quantitity and efficacy of active principle in it. If the structures of the active principles are known it will easier to set up a synthesis instead of isolating it from the plant. Since these natural pesticides usually are very active, they may be found only in minute quantities in the plant. This may be a limitation by its own since it will necessitate the need of huge crops of the plants to provide enough material for practical use. Moreover, isolating new pesticidal compounds from plants remains a difficult and time consuming task, because this process requires interdisciplinary efforts and skills to purify, characterize, structure elucidation, synthesize and screen the compounds for biological activity.

Along with chemistry, genetic knowledge is also necessary to aknowledge in which part of the plant the active principle is most abundant through evaluating the expression analysis in all parts of plant. If it is more expressed in roots then active principle can be produced through hairy root culture. If expression is more abundant in leaves, then cell suspension culture can be made to produce active principle on large scale and if the expression is more in stems, then multiple organogeneses can be used for its higher production. Genetic engineering approaches can be used to overexpress effective pesticidal compound in the same plant to maximize its production. Another approach is to transfer gene of pesticidal compound into the crops, so that those transgenic crops become resistant by themselves to the targeted insect pests.

Alternative approaches for sustainable and reproducible production should be adopted for plant species with specialized activity, other than traditional agriculture and collection of wild plants. In last few years there has been a great progress in development of cell cultivation methods to produce insecticidal compounds (George et al., 2000; Hitmi et al., 2000). In plant cell cultivation method plants can be maintained for indefinite period of time and can be safe from adverse environmental conditions. Recently used biotechnological approaches to increase the productivity of botanical pesticides in cell culture includes the use of elicitors and metabolic engineering methods (Prakash and Srivastava, 2011; Shilpa et al., 2010; Yendo et al., 2010; Pandey et al., 2012). To scale up production of botanical pesticides under controlled conditions in suspension cultures through bioreactors, there are many studies related to economical, technological and scale up aspects (Yesil-Celiktas et al., 2010). Due to problems like slow growth rate, low economic feasibility and low product yield in cell culture, their exploitation on commercial level has not been successful.

Instead of using cell culture, organ culture methods like fast growing hairy root cultures and adventitious root cultures are successful to sustainably produce specific plant metabolites under invitro conditions through transformation with Agrobacterium rhizogenes. These cultures can be maintained for lengthy duration and production is also high. Due to stable production of plant metabolites these techniques can be exploited industrially to upscale the cultures in bioreactors (Baque et al., 2012; Georgiev et al.,

2007; Guillon et al., 2006; Pistelli et al., 2010; Srivastava and Srivastava, 2007). Some commercial botanical insecticides have been produced through hairy root cultures e.g. azadirachtin (Azadirachta indica) (Allan et al., 2002; Srivastava and Srivastava, 2012), thiophenes (Tagetes patula) (Rajasekaran et al., 2004), gossypol (Gossypium hirsutum) (Verma et al., 2009), phytoecdysteroids (Ajuga reptans, Ajuga turkestanica) (Matsumoto and Tanaka, 1991; Chenget al., 2008) and nicotine (Nicotiana rustica) (Hamil et al., 1986; Zhao et al., 2013).

This biotechnological approach is also used to produce pesticidal compounds from endangered plant species e.g. the cultivation under optimized conditions and transformation through Agrobacterium rhizoghenes provides the biotechnology approach for the production of metabolites from Aremisia granatensis which is endangered plant specie. The phytochemical analysis of this plant revealed the presence of sesquiterpenic lactones, monoterpenes and poliacetilenic spiroacetals showing antifeedent effect against aphids (Barrero et al., 2013).

Another approach is aeroponically produced plants under controlled environmental conditions for the production of valuable metabolites from the aerial parts and roots (Martı´n-Laurent et al., 1999). At present, this system has been used to produce biomass to mainly get medicinal compounds on commercial scale (Kim et al., 2012; Hayden, 2006; Xu et al., 2009; Xu et al., 2011).

This study reported the isolation of insecticidal compounds from potential plants and biochemical and molecular characterization of isolated insecticidal compounds.

1.7. Objectives The objectives of this study were to:

• Evaluate pesticidal potential of selected plants against target pests. • Isolate active compounds from the selected plants responsible for pesticidal activity. • Characterize the isolated compounds for their pesticidal activity using different fractions. • Identify genes involved in the synthesis of selected active compounds.

Chapter 2

Materials and Methods

2.1. Pesticidal potential of selected plants against target pests

2.1.1. Plant material The leaves of Cinnamomum camphora (L.) J. Presl (Lauraceae), Eucalyptus sideroxylon A. Cunn. ex Woolls (Myrtaceae), and aerial parts of Isodon rugosus Wall. ex Benth (Labiatae), Boenninghausenia albiflora (Hook.) Rchb. ex Meisn. (Rutaceae), Calotropis procera Aiton (Dryand).(Apocynaceae), Daphne mucronata Royle (Thymelaeaceae), Tagetes minuta L.(Asteraceae) were collected from northern Pakistan (34.1558° N, 73.2194° E) (Table 2.1). All the selected plant species were identified by Dr. Zafar Jamal, Chairman Botany Department, Government College, Abbottabad, KPK, Pakistan.

Table 2.1: Details of plant species used to evaluate pesticidal potential

Scientific Name Common Name Family Part used Daphne mucronata Kashmir Daphne (Kutti Lal) Thymelaeaceae Aerial parts

Tagetes minuta Mexican Marigold (Gule Asteraceae Aerial parts Sad Barg) Calotropis procera Sodom Apple (Ak) Apocynaceae Aerial parts

Boenninghausenia White Himalayan Rue Rutaceae Aerial parts albiflora (Pissu Mar Booti) Eucalyptus sideroxylon Red Ironbark (Gond or Myrtaceae Leaves Safeda) Cinnamomum camphora Camphor Tree (Kafoor) Lauraceae Leaves Isodon rugosus Wrinkled Leaf Isodon (Boi) Lamiaceae Aerial parts

2.1.2. Preparation of plant extracts Plants were dried in the shade. Dried plant material was ground to powder using an electric grinder. Metabolites were extracted by a maceration method using organic solvent methanol at room temperature (Padin et al., 2013). After 2 d the solvent layer was filtered with Watman No.1 filter paper and the process repeated three times. The

40 filtrate was concentrated using a rotary evaporator at 35°C and resulting extracts were stored at 4°C. Dried plant powder and yields of crude methanolic extracts are given in the table 2.2.

Table 2.2: Yields of plant’s crude methanolic extracts

Plant crude extracts (Dry powder) Yields (g)

Daphne mucronata (3.5 Kg) 180

Tagetes minuta (4 Kg) 187

Calotropis procera (3.5 Kg) 133

Boenninghausenia albiflora (300 g) 18.3

Eucalyptus sideroxylon (700 g) 49.3

Cinnamomum camphora (500 g) 32.5

Isodon rugosus (1 Kg) 90

2.1.3. Target insects Insects selected for the study from different insect’s order on the basis of their important economic value and feeding habits. List of selected target insects are shown in the table 2.3.

Table 2.3: Details of target insects used for the study

Scientific name Common name Order Family

Bactrocera zonata Saunders Peach fruit fly Diptera Tephritidae

Shizaphis graminum Rondani Green bug aphid Hemiptera Aphididae

Tribolium castaneum Herbst Red flour beetle Coleoptera Tenebrionidae

Spodoptera exigua Hubner Beet army worm Lepidoptera Noctuidae

Drosophila melanogasterMeigen Common fruit fly Diptera Drosophilidae

Acyrthosiphon pisum Harris Pea aphid Homoptera Aphididae

2.2. Bioassays mediated pesticidal potential of selected plants

2.2.1. Bactrocera zonata Saunders, 1842

2.2.1.1. Rearing of Bactrocera zonata Pupae of fruit fly, B. zonata were collected from the laboratory colony maintained on an artificial larval diet at the Nuclear Institute of Agriculture, Tandojam, Pakistan. Three to 4 d before eclosion of pupae, the pupae were received at the Insect Pest Management Program, National Agricultural Research Centre Islamabad, Pakistan. After emergence the flies were kept in 30 x 30 x 45 cm screened cages and maintained at 26±1°C and 60±5% RH with a 10L: 14D h photoperiod, and fed ad libitum with a protein diet containing hydrolyzed yeast (MP Biomedicals Inc.; www.mpbio.com) and sugar in 1:3 ratio by weight, and water. On the first day of emergence flies were sexed and kept in separated cages having different food regimes. Male and female flies were identified on the basis of morphological characteristics; the female flies have long pointed ovipositor at the end of their abdomen. Female flies were separated into two groups. One group of females was reared on protein diet that contained both yeast and sugar and the other group was reared on sugar only until they were transferred to experimental cages. Females reared on protein diet were used to assess oviposition deterrence and those on sugar only to assess toxicity. The reason for keeping flies deprived of protein

42 was that protein is critical for producing fertile eggs and protein deprived females will show attraction to protein. Therefore, for assessment of toxicity, females were initially maintained on sugar only and switched to a diet containing protein at the onset of their sexual maturity. Males were in one group and reared on protein diet from emergence onwards because males showed strong attraction to methyl eugenol and there was no need to have different food regimes for their attraction purpose.

2.2.1.2. Adult male fruit fly toxicity bioassay After 14 d of emergence B. zonata males from large cage were shifted to experimental screened cages 15 x 15 x 20 cm and kept them for 1 h before bioassay. Laboratory adopted males reached sexual maturity at 14 d. Studies of B. zonata male age response to methyl eugenl (ME) have not been undertaken, however, Shelly et al. (2010) reported that many of the ME responsive males are responsive at the beginning of their sexual maturity. Therefore, sexually mature males were selected for toxicity bioassay by mixing plant extracts with ME. For this bioassay, crude methanolic plant extracts were tested against adult male fruit fly. Four mg each of plant extract was mixed with 200 µL ME and 50 µL added to single filter papers in three replicate Petri dishes. These Petri dishes (without lids) were then placed in experimental cages having male flies at 10:00 h and removed after 24 h. Thirty males (10 males in each replication) were exposed to each treatment. Three controls were included; ME (a negative control), untreated filter paper (a negative control) and organophosphate synthetic insecticide i.e. 2, 2- dichlorovinyl dimethyl phosphate (DDVP; the positive control). In the positive control, 4 µL of DDVP was mixed with 200 µL of ME. Mortality was observed after 24 and 48 h (Fig. 2.1).

Figure 2.1: Adult male fruit fly toxicity bioassay (a) flies exposed to untreated sample containing only methyl eugenol on filter paper (b) flies exposed to filter paper containing methyl eugenol mixed with crude methanolic extract of Calotropis procera. 2.2.1.3. Adul female fruitfly toxicity bioassay From emergence, female flies were provided with sugar only and after 14 d they were transfer to experimental cages, starved for 8 h and switched to protein diet. Feeding toxicity bioassay was used to analyze the toxicity of plant extracts by mixing each plant extract into the diet of flies (Shakunthala and Thomas, 2001b). For this purpose, each crude methanolic plant extract was mixed at 2% into the adult diet and placed on filter paper in Petri dishes. Four mg of each plant extract was first dissolved in 200 µL of methanol and then mixed with diet containing 2 mg of sugar and 2 mg of hydrolyzed

44 yeast and placed in experimental cages. In each treatment total of 30 flies were exposed in three replicates of 10 flies. Three control treatments were included; methanol only (a negative control), food containing hydrolyzed yeast and sugar (a negative control) and commercial protein bait containing Spinosad (GF 120; positive control). In the positive control, 12 mg of GF 120 was mixed with 90 µL of water as recommended by the manufacture. Mortality was observed after 24 and 48 h (Fig. 2.2).

Figure 2.2: Adult female fruit fly toxicity bioassay (a) flies exposed to treated diet mixed with crude methanolic extract of Cinnamomum camphora on filter papers (b) flies exposed to treated diet mixed with crude methanolic extract of Tagetes minuta on filter papers

2.2.1.4. Repellence and oviposition deterrence effect bioassay Fifty each of virgin male and female fruit flies, maintained on protein diet for 14 d, were combined for copulation 90 min before sunset in a 45 x 45 x 45 cm plexiglass screen cage. Fruit fly couples were collected in plastic vials, transferred to separate cages and left to continue copulation. Next morning female flies were transferred to experimental cages and provided a protein diet and water ad libitum. Fifteen flies (five per replicate) were taken for each treatment for evaluation. Next day at 10:00 h, the females were provided access to guava fruit treated with 2% crude methanolic plant extracts. The fruit used were of uniform size, cold treated at 4°C for 22 d in order to eliminate any larvae from wild flies and kept at room temperature for 24 h before exposure for oviposition. In each treatment, 60 mg of plant extract was mixed with 3 mL of methanol. An aliquot of 1 mL was applied on each guava by pipette while continuously rotating the guava to ensure uniform distribution over the fruit. Three guavas were used for each treatment (1 guava per replication). After treatment the guavas were allowed to dry for 2 h, and then exposed to flies for 48 h. For repellence bioassay, settled or repelled females from treated guava in each treatment were counted every 2 h. For oviposition deterrent effect bioassay, female flies were removed after 48 h and guavas were placed in sawdust for 15 d so that larvae could pupate in sawdust. Number of pupae and emerged adults were counted. Two negative control treatments were included; methanol only and untreated guavas (Fig. 2.3).

Figure 2.3: Repellence and oviposition deterrence effect bioassay (a) flies exposed to plant extract treated guavas placed on petri plates inside the cages (b) guavas placed on saw dust for adult emergence inside the cages(c) pupae obtained from guavas in three replicates of Calotropis procera extract’s treatment, placed to further observe the adult emergence. 2.2.1.5. Data analysis Percent repellence was calculated by using the formula (Rehman et al., 2009):

%R= [1/2 (A-B)/A] ×100

Where R represents repellence, A represent half of the number of flies settled on both treated and untreated guavas and B represents number of flies settled on treated guava. Differences in mortality, repellence and oviposition deterrence caused by different plant extracts were analyzed by one-way analysis of variance (ANOVA). Complementary pairwise comparisons of means were performed by Tukey’s test. All analyses were performed with SPSS version 16.

2.2.2. Schizaphis graminum Rondani, 1852

2.2.2.1. Rearing of Schizaphis graminum The green bug aphid, Schizaphis graminum were reared on wheat plants in glass house conditions. Before the start of experiment uniform sized colony was taken from the plants and brought to the laboratory. This aphid colony was maintained on clean wheat leaves in the laboratory until the aphids become healthy and reached to adult stage, required for the bioassay. Two bioassays were used to analyze the toxicity and repellence of aphids after treatment with plant extracts.

2.2.2.2. Toxicity bioassay In this bioassay, no choice method was used. For the bioassay, 6 cm healthy leaf discs were used. Leaf discs were washed with distilled water and dried. 2% solution of each crude methanolic plant extract was prepared in methanol. 400 µL from each 2% plant extract’s solution was applied equally on both sides of each leaf with the help of micropipette. After treatment with plant extract solutions, leaves were kept for drying. After drying, each leaf petiole was tied with cotton soaked in water to prevent early drying of the leaf. Then each treated leaf was placed on filter paper in Petri plates. 10 healthy aphids of same age were placed on leaf in each treatment. Five replications were run for each treatment. Total 50 aphids were used for each treatment. Aphids were observed for mortality after every 12 h from the start of experiment till 48 h. Three controls were used including two negative controls, untreated leaf and leaf treated with methanol and one positive control, leaf treated with synthetic pesticide, lambda cyhalothrin (Karate).

2.2.2.3. Repellence bioassay In this bioassay, choice method was used. As described in toxicity bioassay in section 2.2.2.2, 6 cm healthy leaf discs were prepared and treated with each 2% plant extract. Unlike in toxicity bioassay, in this bioassay two leaf discs were utilized, one was treated with plant extract and other was untreated in each replication of treatment. 10 adult aphids were placed on filter paper in the middle of both leaves to give them choice to move either on the treated or untreated leaf in each replication of each treatment. Five replications were run for each treatment. Aphids were observed for the repellent behavior after every 12 h from the start of experiment until 48 h. Methanol treated leaf discs (a negative control) and Karate treated leaf discs (a positive control) were used as controls (Fig. 2.4).

Figure 2.4: Shizaphis graminum exposed to leaves treated with Calotropis procera crude methanolic extract along with untreated leaves to observed their repellence behaviour 2.2.2.4. Data analysis Percent repellence was calculated by using the formula (Giner et al., 2013). [(C - T)/C + T)]×100 Where, C= Number of aphids on untreated leaf and T= Number of aphids on treated leaf. Differences in mortality and repellence caused by different plant extracts were analyzed by one-way analysis of variance (ANOVA). Complementary pairwise

49 comparisons of means were performed by Tukey’s test. All analyses were performed with SPSS version 16.

2.2.3. Tribollium castaneum Herbst 1797

2.2.3.1. Rearing of Tribolium castaneum The red flour beetle, Tribolium castaneum were reared on wheat flour, supplementing with 5% brewer’s yeast in an incubator at temperature 30°C and 60% relative humidity in darkness (Walski et al., 2016). For the bioassays, adult beetles were stained from flour with the help of stainless steel strainer. Two bioassays were used to analyze the bioactivity of plant extracts.

2.2.3.2. Impregnation bioassay In this bioassay, filter papers were impregnated with each plant extract solution. Impregnation of filter papers was performed with 700 µL of 2% of each extract solution in one replication. Filter papers were dried for 10 minutes at room temperature under flow. After drying, 10 beetles were added along with their diet in each petri plate. Two replications were used for each treatment. All the petri plates with insects were kept at temperature 30°C and 60% relative humidity in an incubator. The mortality was analyzed after 24, 48 and 72 h of the treatment. Two controls were used i.e. methanol treated filter paper and untreated filter paper (Fig. 2.5).

Figure 2.5: Impregnation bioassay against Tribolium castaneum (a) Indiviual petri plate exhibiting the impregnation bioassay in which beetles exposed to plant extract treated filter paper (b) whole experimental setup for beetles exposed to plant extracts treated filter papers 2.2.3.3. Flour discs bioassay In this bioassay, adult beetles were exposed to flour discs (Xie et al., 1996; Yu et al., 2013) containing the plant extracts in them. The flour discs were prepared by dissolving 8.4 mg of each plant extract in 420 µL of methanol and then mixed with 120 mg of corn flour. Aliquots (35 µL) of the mixture containing each of the plant extracts were placed in wells of a 96 well plate (Greiner CELLSTAR)), for making flour discs, and dried overnight under the flow hood to form solid flour diet discs for the beetles. Two control flour discs were prepared (methanol and water). Five flour discs were placed together with10 beetles into each falcon tube per treatment. A total of 10 flour

51 discs were used in two replications for each treatment. Mortality was analyzed at 24, 48 and 72 h after exposure to the treated flour discs.

2.2.4. Drosophila melanogaster Meigen, 1830

2.2.4.1. Rearing of Drosophila melanogaster The common fruit fly, Drosophila melanogaster were reared on agar-yeast-corn meal artificial diet that was prepared as described by Taning et al. (2016), at 25°C, light regime of 16 h light and 8 h dark, and 65% relative humidity. For the bioassay, adult flies were selected from insect cultures from the incubator.

2.2.4.2. Diet preparation For the preparation of diet, 600 mL of water was boiled with 8 g agar. Then 60 g polenta, 60 g torula yeast and 25 g sucrose were added to this solution. In the final step after cooling, 2.5 g nipagine and 25 mL ethanol were added. Then this prepared diet was poured into the falcon tubes up to two inches and after solidifying and caping; tubes were placed at 0°C until used.

2.2.4.3. Contact toxicity For contact toxicity bioassay, 200 µL of 2% (20 mg of the extract in 1 mL of methanol) of each plant extract was layered with the help of micropipette on the surface of a yeast-corn meal artificial diet in 50 mL tubes. The added plant extracts were dried on the surface of the artificial diet (under a flow chamber for about 2 h) before the flies were transferred in the tubes. 10 adult flies selected at random were then exposed to each of the treatment in the tubes. The experiment was replicated three times. Methanol and water without any plant extract were used as negative controls for the experiment. Mortality was assesed at 24, 48 and 72 h (Fig. 2.6).

Figure 2.6: Contact toxicity of different selected plant crude methanolic extracts against Drosophila melanogaster 2.2.4.4. Data analysis Corrected mortalities were calculated by using Abbott’s formula (Abbott, 1925). Differences among treatments were analyzed by One-way analysis of variance (ANOVA) followed byTukey’s Test. All analyses were performed with SPSS, version 16.

2.2.5. Spodoptera exigua Hubner, 1808

2.2.5.1. Rearing of Spodoptera exigua An established colony of beet armyworms (S. exigua) was reared in a growth chamber at 25 °C, 65% relative humidity and a 16 h light: 8 h dark photoperiod. Pupae of S. exigua were placed in 40 × 25 × 25 cm Plexiglass cages, lined on the inner wall with white A4 paper as a substrate for oviposition. The emerging adults were fed with a 10% solution of honey in water. Eggs laid on the paper were transferred to a plastic container and the hatching larvae were fed on an agar-based artificial diet (Smagghe et al., 1998). Second-instar larvae were used for the bioassays.

2.2.5.2. Diet preparation The solution containg 2600 mL of water and 38 g of agar was boiled for 5 minutes. Then 300 g of polenta, 120 g of wheat germs, 100 g of torula yeast, 20 g of casein and 14 g of wesson salt mix were added to this water solution. 18 g of vitamin C/ascorbic acid and 80 mg of vitamin mix*

53 were added in 100 mL water to make separate solution. This 100 mL solution was poured in the mixture already prepared. Finally, 8 g of sorbic acid, 4 g of nipagin and 600 mg streptomycine (antibiotic) were added to the mixture. This mixture of diet was poured in 20 well plates and placed at 0 °C until used.

*Vitamin mix: 1. Nicotin acid 250 mg

2. VB2 riboflavine 125 mg

3. VB1thiamin 57.5 mg

4. VB6pyridoxine 57.5 mg 5. Panthotenic acid (B5) 250 mg 6. Folic-acid 5 mg 7. Biotin 5 mg

2.2.5.3. Contact toxicity For the bioassay, second instar larvae were used. The artificial diet was prepared and poured in 24 wells plates and allowed to settle it for 30 min. After diet settled in the wells, 50 µL of 2% solution (20 mg of the extract in 1 mL of methanol) of each plant extract was layered on diet each well with the help of micropipette. The well plates with treated diet were placed under flow hood to dry the methanol for 1 h. After letting the methanol dry off the surface of the diet, one insect was added in each well. There were 20 insects used for each treatment. Two controls were used, methanol and untreated diet. After adding insects the plates were kept in the incubator. Mortality was analyzed at 24, 48 and 72 h (Fig. 2.7).

Figure 2.7: Contact toxicity against Spodoptera exigua (a) 20 well plates with diet (b) 20 well plates covered with lids after adding second instar larvae of Spodoptera exigua 2.2.5.4. Data analysis Mortalities were corrected by using Abbott’s formula (Abbott, 1925). Differences among treatments were analyzed by One-way analysis of variance (ANOVA) followed by Tukey’s Test. All analyses were performed with SPSS, version 16.

2.2.6. Acyrthosiphon pisum Harris, 1776

2.2.6.1. Rearing of Acyrthosiphon pisum A continuous colony of the A. pisum was maintained on young plants of Vicia faba L. (Fabales: Fabaceae) at 23–25°C and 65±5% relative humidity (RH) under a 16:8 h light: dark photoperiod (Nachman et al., 2011). For the bioassays, adult aphids were transferred from fresh leaves into separate boxes from which neonates were collected after 24 h. The collected neonates were then used for the bioassays.

2.2.6.2. Diet preparation For the preparation of diet, 30 mL solution of amino acids*, 600 µL solution of minerals*, 3 mL solution of vitamins*, and 18 mL solution of sucrose mixture* and 6 mL of phosphate were mixed in a beaker and stirred for 15 minutes at room temperature. This mixture was filtered through whattman filter paper and transferred in eppendorf tubes.This diet was kept at -20°C until used. *Amino acids: The amino acids solution was prepared by adding 0.2080 g alanine, 0.8636 g asparagine monohydrate, 0.7640 g aspartate, 0.1400 g cysteine, 0.5040 g glutamate,

1.0120 g glutamine, 0.0480 g glycine, 0.2600 g proline, 0.2440 g serine, 0.0480 g tyrosine, 1.0080 g arginine, 0.7296 g histidine monohydrochloride monohydrate, 0.4560 g icoleucine,0.4560 g leucine,0.6347 g lysine monohydrochloride, 0.1680 g methionine, 0.1920 g phenylalanine, 0.4120 g threonine, 0.2360 g tryptophan and0.1560 g valinein 200 mL of distilled water.

*Minerals:

The mineral solution was prepared by adding 0.0055 g FeCl3.6H2O, 0.0008 g CuCl2.2H2O, 0.0020 g MnCl2.4H2O and 0.0094 g of ZnSO4.H2O in 5 mL of distilled water.

*Vitamins The solution of vitamins was prepared by adding 0.0004 g d-biotin, 0.0435 g D- calcium pantothenate, 0.0080 g folic acid, 0.0400 g niacin, 0.0122 g pyridoxine hydrochloride, 0.0112 g thiamine hydrochloride, 0.2681g choline chloride and 0.2000 g myo-Inositol in 20 mL of distilled water.

*Surose solution Sucrose solution was prepapre by adding 0.0600 g ascorbic acid, 0.0060 g citric acid, 0.1200 g MgSO4.7H2O and 10.2000 g sucrose in 18 mL of distilled water.

*Phosphate

To prepare phosphate solution 0.9040g of K2HPO4.3H2O was mixed with 6 mL of distilled water.

2.2.6.3. Feeding toxicity bioassay Different concentrations were used for each plant extract against aphids. The stock solution of 1% was prepared by adding 1 mg of each plant crude extract in 100 µL of methanol and then five concentrations were prepared i.e. 1000 ppm, 500 ppm, 200 ppm, 100 ppm and 50 ppm from stock solution by diluting it in artificial diet for aphids to make 300 µL final volume for three replications of each treatment, 100 µL for each replication. For the bioassay, artificial diet test cages were prepared by stretching a layer of parafilm over a hollow plexiglass tube. 100 µL of artificial diet was pipette on this first layer and sealed by stretching a second layer of parafilm on it (Nachman et al., 2011). 10 neonate aphids were placed on second layer of parafilm and covered with hollow plastic ring with ventilated lid to prevent the escape of aphids (Fig. 2.8). These

56 cages were placed in inverted position in six well plates. Two controls were used i.e. untreated artificial diet and methanol treated artificial diet. Three replications were used for each treatment. Mortality was observed after 24 h of exposure by gentle probing of the aphids with a brush and observing post-mortem color change of the body (Sadeghi et al., 2009).

Figure 2.8: Feeding bioassay against A. pisum (a) plant extract treated diet was pipette on parafilm layered on plexiglass (b) Treated diet fixed between two parafilm layers (c) neonate aphids after 24 h of adult collected on fresh leaves (d) hollow ring placed on parafilm layers containing treated diet (e) 10 neonate aphids were placed on each parafimed tube (f) hollow ring covered from top by placing ventilated ring (g) plexiglass placed invertly on well plate after adding aphids (h) whole bioassay setup.

2.2.6.4. Data analysis Probit analysis of mortality vs concentration was conducted to estimate lethal concentrations (LC50 and LC90) with their corresponding 95% confidence intervals (95% CI) by POLO Plus V 2.0 (LeOra Software Inc., Berkeley, CA). LC values were considered to be significantly different when their respective 95% CI did not overlap.

2.3. Isolation of active compounds from the selected plants responsible for pesticidal activity On the basis of previous bioassays with seven plant crude methanolic extracts with all target insects, two most active plants i.e. Daphne mucronata and Isodon rugosuswere selected to evaluate the toxicity of different fractions of their respective extracts against most susceptible insect, Acyrthosiphon pisum among all the target insects analyzed.

2.3.1. Daphne mucronata fractionation The 180 g of crude methanolic extract was suspended in water (900 mL), then liquid-liquid partitioned successively by n-hexane four times (150 mL), dichloromethane four times (150 mL), ethyl acetate four times (150 mL) and n-butanol four times (150 mL), was performed (Fig. 2.9). The extracts were filtered and concentrated by evaporation under reduced pressure (around 20 mbar) with a rotavapor at 40°C to afford a black green n-hexane residue (41.84 g), a shiny dark green dichloromethane residue (2.94 g), a brownish green ethyl acetate residue (4.95 g), and a brown n-butanol residue (44.79g).

Figure 2.9: Daphne mucronata fractionation (a) Step 1, crude methanolic plant extract (b) Step 2, crude methanolic plant extract suspended in water (c) Step 3, liquid-liquid extraction in separating flask (d) Step 4, drying solvent from solvent fraction (e) four different solvent fractions obtained 2.3.2. Isodon rogusus fractionation The 90 g of crude methanolic extract was suspended in water (450 mL), then liquid-liquid partitioned successively by n-hexane three times (150 mL), dichloromethane three times (150 ml), ethyl acetate three times (150 mL) and n-butanol three times (150 mL), was performed (Fig. 2.10). The extracts were filtered and concentrated by evaporation under reduced pressure with a rotavapor at 40°C to afford a black green n- hexane residue (8.82 g), a shiny green dichloromethane residue (3.1 g), a light green ethyl acetate residue (1.33 g), and a yellow n-butanol residue (6 g).

Figure 2.10: Four different solvent fractions obtained from crude methanolic extract of Isodon rugosus 2.3.3. Bioactivity of fractions of Dhaphne mucronata and Isodon rugosus For the bioassay with four fractions i.e. hexane, dichloromethane, ethylacetate and butanol fractions of both D. mucronata and I. rugosus, 1% stock solution for each fraction was prepared by adding 1 mg of each fraction in 100µL of their respective solvents. Then from 1% stock solution five concentrations were prepared i.e. 500 ppm, 200 ppm, 100 ppm, 50 ppm and 25 ppm by diluting in artificial diet of aphids. 100 µL of each solution was layered on parafilm and cages were prepared as described in section 2.2.6.3. 10 neonate aphids were exposed to treated diet in each cage for 24 h. Three replications were used for each treatment. Five controls were used i.e hexane, dichloromethane, ethylacetate and butanol treated artificial diet, and untreated artificial diet. Mortality was observed after 24 h.

2.3.4. Sub-fractionation of 500 mg butanol fraction of I. rugous through first reverse phase automatic flash chromatography On the basis of bioactivity, the most bioactive butanol fraction of I. rugosus was selected for further fractionation and isolation of bioactive compounds as compared to other fractions analyzed. Automatic reverse phase flash chromatography was performed with the Reveleris Flash System from GRACE, United States. 500 mg of butanol fraction was loaded on 1 g reverse phase silica gel and fractionated through reverse phase automatic flash chromatography into a total of 95 fractions by using three solvents in which solvent A was water, solvent B was methanol and solvent C was acetonitrile. A dry column was prepared with the loaded silica and a pre-packed 12 g C18 reverse phase

60 column was used to fractionate it, using a flow rate of 30 mL per minute with a run length of 67 column volumes; volume per vial was set as 25 mL per vial (Table 2.4). A solvent gradient was selected by analyzing 1 mg of sample on HPLC-MS by using different solvent gradients, the gradient that gave the best separation as detected with UV was selected for reverse phase automatic flash chromatography. The gradient, starting with 100% water and 0% methanol for 1 column volume, went to 100% methanol in next 60 column volumes, then for next column volume it went to 100% acetonitrile and stayed at 100% acetonitrile for next last 5 column volumes. The 95 fractions were combined based on UV spectra at 220 nm into a total of 14 fractions (1A-14A). These combined fractions were dried under reduced pressure at 45˚C and finally dried under high vacuum.

Table 2.4: First reverse phase automatic flash chromatography conditions of butanol fraction of Isodon rugosus

Run Conditions

Cartridge Reveleris 12 g C18 40 µm

Solvent A Water

Solvent B Methanol

Sovent C Acetonitrile

Flow rate 30 mL/min

Injection type Dry sample

ELSD Carrier Isopropanol

Per vial volume 25 mL

UV1 Wavelength 220 nm

UV2 Wavelength 254 nm

2.3.4.1. Bioactivity of sub fractions of butanol fraction from first reverse phase automatic flash chromatography For the bioassay, 1% stock solution of each fraction was prepared with different ratios of water and methanol. Different concentrations i.e. 50 ppm, 25 ppm, 12.5 ppm, 6.25 ppm and 3.125 ppm were prepared from stock solution by diluting with artificial diet. Cages were prepared as described in section 2.2.6.3 and 10 neonate aphids were exposed to100 µL of artificial diet in each cage for each replication of treatment. Three replications were used for each fraction. Two controls were used i.e. methanol treated artificial diet and untreated artificial diet. Mortality was analyzed after 24 h.

2.3.5. Sub-fractionation of most active fraction 3A from first reverse phase automatic flash chromatography through preparative high performance liquid chromatography (prep-LC) Out of a total of 14 fractions from first reverse phase automatic flash chromatography, the most bioactive fraction 3A was selected for further fractionation through prep-LC on the basis of best bioactivity. The prep-LC system consisted of Agilent 1100 series equipped with VWD detector and automatic fraction collector. A 10% solution of fraction 3A was prepared in methanol. Two solvents were used, solvent A was water and solvent B was acetonitrile. Prep-LC conditions were: Ascentis C18 column with guard cartridge, 21.2 mm ID × 150 mm (5 µm); flow rate 6 mL/min; UV detection at 220 nm; room temperature; injection volume of 50 µL; running time 132 minutes per injection; automatic fraction collector mode time-based. The gradient started with 100% water and went to 18% acetonitrile during 100 min; subsequently, the acetonitrile percentage rose to 100 during the next 10 minutes. It was kept at 100% until 128 minutes and then the acetonitrile went to 0% at 128.10 min and stayed at 0% until 132.10 min. Three fractions were collected, 3A-1, 3A-2 and 3A-3 after drying under reduced pressure with a rotary evaporator and finally under high vacuum.

2.3.5.1. Bioactivity of sub-fractions collected from prep-LC The bioactivity of all the three fractions 3A-1, 3A-2 and 3A-3, was analyzed against aphids. For bioassay, 1% stock solution was prepared by dissolving 1 mg of fraction in HPLC grade water. Five concentrations were prepared, 5 ppm, 2.5 ppm, 1.25

62 ppm, 0.625 ppm and 0.3125 ppm by diluting with diet of aphids. 300 µL of solution was prepared for each treatment. Three replications were used for each treatment. Bioassay was same as described in section 2.2.6.3. 10 neonate aphids were exposed to each cage containing 100 µL solution of each treatment. One control was used, untreated diet. Mortality was observed after 24 h.

2.3.5.2. Spectroscopic analysis of prep-LC most bioactive fraction, 3A-3 Out of three fractions collected from prep-LC (3A-1, 3A-2 and 3A-3), 1H NMR was recorded for most bioactive fraction, 3A-3. Different gradients were used to purify the compound but during different Prep-LC runs, the chromatographic behavior, i.e. peak shape and position, of this fraction was inconsistent. Therefore, reverse phase automatic flash chromatography was repeated with 5 g of butanol fraction from I. rugosus in order to get the most bioactive compound in pure form.

2.3.6. Sub-fractionation of 5 g butanol fraction of I. rugous through second reverse phase automatic flash chromatography A sample for the second reverse phase automatic flash chromatography was prepared by loading 5 g butanol fraction of I. rugosus on 10 g reverse phase silica gel. The loaded sample was packed in a column in dry form and a 120 g pre-packed C18 reverse phase column was used to fractionate this dry sample. Based on previous knowledge of the first reverse phase automatic flash chromatography, two solvents were used, solvent A was water and solvent B was methanol (Table 2.5). The gradient was set as starting with 100% water for up to 5 column volumes, after 5 column volumes water went to 0% until 60 column volumes, then for last 5 column volumes methanol stayed at 100%. The flow rate was set as 85 ml per min, column length was 55.7 and per vial volume was 25 mL. A total of 354 fractions were collected. On the basis of UV spectra at 220 nm, 354 fractions were combined into 6 fractions (1B-6B).

Table 2.5: Second reverse phase automatic flash chromatography conditions of butanol fraction of Isodon rugosus

Run Conditions

Cartridge Reveleris 120 g C18 40µm

Solvent A Water

Solvent B Methanol

Flow rate 85mL/min

Injection type Dry sample

ELSD Carrier Isopropanol

Per vial volume 25 mL

UV1 Wavelength 220 nm

UV2 Wavelength 254 nm

2.3.6.1. Bioactivity of sub fractions of butanol fraction from second reverse phase automatic flash chromatography Bioactivity of six fractions (1B-6B) collected from second reverse phase automatic flash was analyzed by making 1% stock solution by adding 1 mg of each fraction in different ratios of water and methanol. Five concentrations, 50 ppm, 25 ppm, 12.5 ppm, 6.25 ppm and 3.125 ppm were prepared from 1% stock solution by diluting in diet. Three replications were used for each concentration. 10 neonate aphids were exposed to treatment in each replication. Bioassay procedure was same as descrbed in section 2.2.6.3. Two controls were used, methanol treated diet and untreated diet. Mortality was analyzed after 24 h of treatment.

2.3.7. Acidic extraction of most bioactive fraction 1B from second reverse phase automatic flash chromatography The most bioactive fraction 1B was selected for further isolation on the basis of best activity among all 6 fractions (1B-6B) from the second reverse phase automatic flash chromatography. Using the knowledge from 1H NMR spectra of first reverse phase automatic flash chromatography fraction 3A-3, acidic extraction was done to isolate a pure insecticidal compound from fraction 1B of second reverse phase automatic flash chromatography. A 200 mg of fraction 1B was dissolved in 10 mL of distilled water and acidified with 4 drops of hydrochloric acid. Then this solution was extracted four times with 15 mL of ethyl acetate in a separating funnel. Both the organic and water phase were evaporated. The organic phase was treated with toluene to remove the remaining ethyl acetate azeotropically from the obtained fraction. After drying under high vacuum; 60 mg of material was obtained out of the ethyl acetate phase fraction and 60 mg out of the water phase fraction.

2.3.7.1. Bioactivity of ethyl acetate and water phase obtained through acidic extraction 1% stock solution was prepared by adding 1 mg of both ethyl acetate and water phase fractions in water. Five concentrations, 5 ppm, 2.5 ppm, 1.25 ppm, 0.625 ppm and 0.3125 ppm were prepared from 1% stock solution by diluting them in diet. Three replications were used for each concentration. 10 neonate aphids were exposed to treatment in each replication. Bioassay procedure was same as described in section 2.2.6.3. One control was used, untreated diet. Mortality was analyzed after 24 h of treatment.

2.3.8. Identification of most bioactive compound, rosmarinic acid The ethyl acetate fraction appeared to bemore bioactive than the water fraction after biological evaluation. Different techniques were used to identify isolated pure insecticidal compound, i.e. high-performance liquid chromatography-mass spectrometry (HPLC-MS), nuclear magnetic resonance (NMR) spectroscopy and optical rotation measurement.

2.3.8.1. HPLC-MS of isolated bioactive compound, rosmarinic acid A HPLC-MS analysis of the ethyl acetate phase fraction showed the presence of a single compound, which was further confirmed by using the MS in negative mode. HPLC-MS was programmed with two solvents, water and methanol, starting with 100% water and 0% methanol and went to 100% methanol and 0% water in 16 minutes. The HPLC-MS consisted of Agilent LC/MSD 1100 series.

2.3.8.2. Specific optical rotation of isolated bioactive compound, rosmarinic acid For optical rotation determination of rosmarinic acid two samples were prepared by dissolving 6.114 mg and 6.995 mg in 3 mL of methanol separately. Ten observations were taken to get the average value.

2.3.8.3. 1H and 13C NMR spectra of isolated bioactive compound, rosmarinic acid 1H and 13C NMR spectra of isolated pure compound were recorded on a Bruker Avance-III 400 MHz NMR Spectrometer.

2.4. Characterization of the isolated compounds for their pesticidal activity using different fractions

2.4.1. Bioactivity of isolated rosmarinic acid and commercial rosmarinic acid Bioactivity of rosmarinic acid from Isodon rugosus and commercially available rosmarinic acid were analyzed. For bioassay, 1% stock solution was prepared by dissolving 1 mg of both compounds in HPLC grade water separately. Eight concentrations were prepared, 50 ppm, 25 ppm, 12.5 ppm, 6.25 ppm, 3.125 ppm, 1.5625 ppm, 0.78125 ppm and 0.390625 ppm by diluting with diet of aphids. 300 µL of solution was prepared for each treatment. Three replications were used for each treatment. Bioassay procedure was same as described in section 2.2.6.3. 10 neonate aphids were exposed to each cage containing 100 µL solution of each treatment. One control was used, untreated diet. Mortality was assessed after 24 h of the treatment.

2.4.2. Comparison between growth of live aphids exposed to plant extract treated and untreated diet after 24 h of bioassay After bioassay with isolated rosmarinic acid from I. rugosus, the growth of live A. pisum was observed until 9 days to determine the effect of compound on the growth of treated aphids as compared to aphids exposed to untreated diet.

2.5. Identification of genes involved in the synthesis of selected active compound, rosmarinic acid

2.5.1. Plant material Leaves of the plant, Isodon rugosus were collected in liquid nitrogen and brought to the laboratory where they were washed with RNAase free water and after drying they were kept at -80°C in laboratory for 2 h. After freezing at -80°C, leaves were immediately transferred to sterilized zipper bags and were placed in lyophilizer where they were freeze dried for 72 h.

2.5.2. RNA extraction RNA from leaves of I. rugosus was extracted by using commercial kit (Qiagen). For RNA extraction, 40 mg of freeze dried leaves were taken and crushed in liquid nitrogen through pestle mortar that were already chilled with liquid nitrogen. Plant material along with liquid nitrogen was transferred to 2 mL eppendorf tubes and immediately added 450 µL of RLT buffer in each tube. This solution was vortexed at high speed for 2 min. Lysate was transferred to akit supplied spin columns already placed in 2 mL eppendorf tubes and centrifuged at 16000 rpm for 2 min. Flow through was collected carefully from each tube and transfer to 1.5 mL eppendorf tubes. Ethanol was added in 0.5 volumes in each sample in each tube with continued pipetting. This solution from each tube was transferred to kit supplied spin columns and centrifuged for 2 min at 16000 rpm. Flow-through was discarded and 700 µL of RW1 buffer was added in each tube and centrifuged at 12000 rpm for 15 sec. After discarding flow-through, 500 µL of RPE buffer was added and centrifuge at 1600 rpm. After discarding flow-through spin columns were again centrifuged to eliminate the remaining RPE buffer. Spin columns were placed in 1.5 mL eppendorff tubes and 50 µL of RNAase free water was

67 pipetted on the top of the each spin column. These spin columns were centrifuged at 12000 rpm to collect the extracted RNA from each spin column membrane into RNAase free water. The extracted RNA was immediately stored at -80°C in a freezer.

2.5.2.1. Gel electrophoresis and quantification of RNA Quality of extracted RNA was checked by running 2 µL of RNA on a gel in 1.5 X TE buffer at 100 volts for 25 to 30 min and gel was observed and photographed by Bio- Rad’s gel documentation system after soaking in ethidium bromide solution for 15 min and washing in distilled water. The RNA quantity was checked by Nanodrop DeNovix DS-11 Spectrophotometer, by using RNAase free water as a blank and 260/230 and 260/280 absorbance ratios were also analyzed.

2.5.3. cDNA synthesis cDNA was synthesized from 1 µg of RNA using SuperScriptTM II Reverse Transcriptase by Thermo Fisher Scientific. A total of 20 µL reaction volume was prepared. In PCR tubes, 10 µL of RNA solution, 1 µL of Oligo dt and 1 µL of dNTPs were added and incubated at 65°C for 5 min. Then this mixture was chilled on ice and spined. 4 µL of 5X first strand buffer, 2 µL of 0.1 M DTT and 1 µL of nuclease free water were added to the chilled mixture and after spinning, it was incubated at 42°C for 2 min. After spinning 1 µL of Superscript II reverse transcriptase was mixed by pipetting gently. This mixture was spined and incubated at 42°C for 50 min, at 70°C for 15 min. After incubation, mixture was spined and the resulting synthesized cDNA was stored at - 20°C. DNA quantity was checked by Nanodrop DeNovix DS- 11 Spectrophotometer, by using nuclease free water as a blank and 260/230 and 260/280 absorbance ratios were also observed.

2.5.4. Target genes In the biosynthesis of rosmarinic acid, eight genes have been involved as described in Coleus blumei by many researchers (Petersen et al., 1993, 1994; Kim et al., 2004; Eberle et al., 2009). Two key genes out of total eight genes were targeted for cloning from Isodon rugosus. These two targeted key genes were rosmarinic acid synthase gene (RAS) and hydrooxyphenylpyruvate reductase (HPPR) gene. (Fig. 2.11)

Figure 2.11: Biosynthetic pathway of isolated pure compound, rosmarinic acid indicating two keys genes i.e. RAS (rosmarinic acid synthase gene) and HPPR (Hydrooxyphenylpyruvate reductase) 2.5.4.1. PCR amplification of rosmarinic acid synthase gene (RAS) Three sets of primers were used for amplification of rosmarinic acid synthase gene, their names, sequence and the plant species from which these primes were designed are listed in the table 2.6. Degenerate primers were designed by chosing highly conserved region by aligning sequences for RAS from closely related plant species, Melissa officinalis, Perilla frutescens, Salvia miltiorrhiza and Solenostemon scutellarioides obtained from NCBI Gene Bank (http://www.ncbi.nlm.nih.gov). Gradient PCR amplification was carried out for 33 cycles in 20 µL reaction mixture. This mixture contained 0.9 µL cDNA, 0.2 µL Taq DNA polymerase, 0.6 µL 10mM dNTPs, 0.6 µL each 10 µM forward and reverse primers, 0.6 µL 50 mM MgCl2 and 2 µL 10X PCR buffer. The conditions used for amplification through gradient PCR were; denaturation at 94°C for 2 min, then 33 cycles starting with denaturation at 94°C for 30 sec, anealing temperatures used were 10 temperatures from 50 to 60°C for 30 sec, extension at 72°C for 45 sec and final extension at 72°C for 10 min.

Table 2.6: Primers used in the amplification of rosmarinic acid synthase gene Primers Sequence Plant specie

F-RAS1 5′-ATTACATATGAAGATAGAAGTCAAAGA Coleus blumei

CTC-3′

R-RAS1 5′-TAGGATCCTCATCAAATCTCATAAAACA

ACTTCTC-3′

F-RAS2 5′-AAGGGAATTTCCACGTACCC-3′ Melissa officinalis

R-RAS2 5′-ACCCAGCTAATCACCCACAA-3′

F-RAS3 5′-GACGAAGCTCCACATCCCCTTC-3′ Melissa officinalis

R-RAS3 5′-GCGCTCCATATGCTGCGTGT- 3′

2.5.4.2. PCR amplification of hydrooxyphenylpyruvate reductase (HPPR) One set of primers was used for amplification of HPPR gene from Isodon rugosus. Primer’s names, sequence and the plant species from which these primes were designed are listed in the table 2.7. Degenerate primers were designed by chosing highly conserved region by aligning sequences for HPPR from closely related plant species, Perilla frutescens, Solenostemon scutellarioides, Salvia officinalis and Salvia miltiorrhiza obtained from NCBI Gene Bank (http://www.ncbi.nlm.nih.gov). PCR amplification was carried out in 20 µL reaction mixture. This mixture contained 0.9 µL cDNA, 0.2 µL Taq DNA polymerase, 0.6 µL 10mM dNTPs, 0.6 µL each 10 µM forward and reverse primers, 0.6 µL 50 mM MgCl2 and 2 µL 10X PCR buffer. The conditions used for amplification in PCR were denaturation at 94°C for 2 min, 33 cycles started with denaturation at 94°C for 30 sec, annealing at 56°C for 30 sec, extension at 72°C for 45 sec and final extension at 72°C for 10 min.

Table 2.7: Primers used in amplification of hydrooxy phenylpyruvate reductase gene Primers Sequence Plant Specie

F-HPPR 5′- GCGCTGCCGAAATTGGAGAT- 3′ Perilla frutescens

R-HPPR 5′-CGTTTCTGGAGTCAGCGCACA-3′

2.5.4.3. Gel electrophoresis PCR products were analyzed on gel electrophoresis in 1.5 X TE buffer at 100 volts for 30 to 35 min. After running the complete gels, these were soaked in ethidium bromide solution for 15 min and after washing in distilled water the gels were then observed and photographed by Bio-Rad’s gel documentation system. The amplified bands were cut from the gels and after purification with Wizard® SV Gel and PCR Clean-Up System, sent for sequencing.

2.5.5. 3′ and 5′RACE PCR for sHPPR (HPPR from I. rugosus) After getting the sequence for conserved region of sHPPR from I. rugosus, rapid amplification of cDNA ends (RACE) PCR was performed to get full length sequence for sHPPR from I. rugosus. For 3′ and 5′RACE PCR SMARTer™ RACE cDNA Amplification Kit from Clontech was used. bAll the reagents and chemicals were provided by the kit except RNA that was isolated from Isodon rugosus in the laboratory.

2.5.5.1. 3′ and 5′RACE cDNA synthesis For each 10 µL 3′ and 5′RACE cDNA synthesis reaction, 2 µL of 5X First-Strand Buffer, 1 µL of 20 mM DTT and 1 µL of 10mM dNTPs were mixed in eppendorf tube to make buffer mix and after spinning kept at room temperature. In separate eppendorf tube 1 µL of RNA and 1 µL of 3′-CDS Primer A were combined and for final volume of 4.75 µL, nuclease free water was added in case of 3′cDNA synthesis. In case of 5′ cDNA synthesis, in another eppendorf tube 1 µL of RNA and 1 µL of 5′-CDS Primer A were combined and for final volume of 3.75 µL, nuclease free water was added. RNA mixture was prepared while keeping the tubes on ice. Both mixtures were mixed by spinning. Both the tubes were incubated at 72°C for 3 min and 42°C for 2 min. Then the mixure was mixed and contents were collected at the bottom through spinning. In 5′ cDNA

71 reaction 1 μl of the SMARTer IIA oligo was added by keeping the tube on ice. Master mix was prepared for both 3′ and 5′ cDNA synthesis reaction by mixing 4 µL of previously prepared buffer mix, 0.25 µL of RNase Inhibitor (40 U/μL) and 1 µL of 100 U SMARTScribe Reverse Transcriptase. Master mix in a volume of 5.25 µL was added in each of 3′ and 5′ cDNA synthesis reactions to made final volume of 10 µL. After mixing with pipetting and spinning, tubes were incubated at 42°C for 90 min and at 70°C for 10 min. 20 µL ofTricine-EDTA buffer was added in each synthesized cDNA reaction(s) to dilute the sample. Both synthesized 3′ and 5′RACE cDNAs were kept at -20°C until used.

2.5.5.2. 3′ and 5′RACE PCR For 3′ and 5′RACE PCR, 50 µL reaction was prepared. For each 50 µL reaction, 34.5 µL of PCR-Grade water, 5 µL of 10X Advantage 2 PCR Buffer, 1 µL 10 mM dNTP Mix and 1 µL of 50X Advantage 2 Polymerase Mix were mixed in eppendorf tubes on ice to prepare the master mix. After mixing and spinning, for 3′RACE PCR amplification, 2.5 µL of 3′ RACE synthesized cDNA, 5 µL of 10X UPM (universal primer mix), 1 µL of 10 µM 3′sHPPR gene specific primer (Table 2.8) and 41.5 µL of master mix were added in PCR tubes on ice. For 5′RACE PCR amplification, 2.5 µL of 5′RACE synthesized cDNA, 5 µL of 10X UPM (universal primer mix), 1 µL of 10 µM 5′sHPPR gene specific primer (Table 2.8) and 41.5 µL of master mix were added in PCR tubes on ice. After spinning, each PCR tube was incubated under the following conditions, 5 cycles of 94°C for 30 sec and 72°C for 10 min, 5 cycles of 94°C for 30 sec, 70°C for 30 sec and 72°C for 10 min, and finally 25 cycles of 94°C for 30 sec, 68°C for 30 sec and 72°C for 10 min.

Table 2.8: sHPPR gene specific primers for 3′ and 5′RACE PCR amplfication

Primers Sequence

3′RACE sHPPR 5′-GATTTGGCGATCGGGTTGATGTTG-3′

5′RACE sHPPR 5′-CCCAATTCTGCCCAATCCTATGATGC-3′

2.5.5.3. Gel electrophoresis Both 3′ and 5′RACE PCR products were run in 10X TE buffer gel electrophoresis at 100 volts for 30 to 35 min. After running the complete gels, these were soaked in ethidium bromide solution for 15 min and after washing in distilled water gels were then observed and photographed by Bio-Rad’s gel documentation system. The amplified bands were cut from the gel and after purification with Wizard® SV Gel and PCR Clean- Up System, sent for sequencing.

2.5.6. Expression of HPPR in leaves, roots and stems of I. rugosus

2.5.6.1. RNA extraction Lyophilized leaves, roots and stems were used for the extraction of RNA. RNA was extracted three times on three different days from leaves; roots and stems in order to accomplished three biological repetitions. Qiagen kit was used to extract RNA from all the samples as described in section 2.5.2. Extracted RNA was analyzed through gel electrophoresis before DNAase treatment. For DNAase treatment of the extracted RNA, TURBO DNA-FreeTM Kit by Thermo fisher Scientific was used. For this treatment, 45 µL of RNA extracted from roots, leaves and stems, was used separately. 0.1 volume of 10X TURBO DNAase buffer and 1 µL of TURBO DNAse were added to each RNA solution while keeping eppendorf tubes on ice and mixed gently by pipetting. After spinning, this mixture was incubated at 37°C for 30 min. After mixing, 0.1 volume of the DNAse inactivation reagent was added to the RNA mixture. This mixture was incubated at room temperature for 5 min with occasional shaking. After incubation it was centrifuged at 10,000 rpm for 1.5 min and supernatant (RNA) was collected carefully and kept at -80°C. After DNAse treatment each RNA sample was analyzed through gel electrophoresis before cDNA synthesis.

2.5.6.2. cDNA synthesis cDNA was synthesized from 1 µg of each RNA using SuperScriptTM II Reverse Transcriptase by Thermo Fisher Scientific. The same procedure was used as described in section 2.5.3. The synthesized cDNA was store at -20°C.

2.5.6.3. RNA and cDNA quantification RNA and DNA quantity was checked by Nanodrop DeNovix DS- 11 Spectrophotometer, by using nuclease free water as a blank and 260/230 and 260/280 absorbance ratios were also observed.

2.5.6.4. PCR amplification For PCR amplification 20 µL of reaction mixture was used for each reaction of leaves, roots and stems, separately. For each reaction, 0.9 µL cDNA, 0.2 µL Taq DNA polymerase, 0.6 µL 10mM dNTPs, 0.6 µL each 10 µM forward and reverse primers for

HPPR and for actin gene respectively, 0.6 µL 50 mM MgCl2 and 2 µL 10X PCR buffer were added in PCR tubes. Actin gene was used as reference gene in each leaves, roots and stem reaction. Actin and HPPR gene primers used in the reactions are listed in the table 2.9. The conditions used for amplification in PCR were denatuaration at 94°C for 2 min, 33 cycles of denaturation at 94°C for 30 sec, primer annealing at 56°C for 30 sec, extension at 72°C for 45 sec and final extension at 72°C for 10 min.

Table 2.9: Actin and HPPR primers used in PCR amplification of HPPR and actin gene from roots, leaves and stem of Isodon rugosus

Primers Sequence Plant Specie

F-Actin 5′-GAGGTTGGATCTTGCTGGTC-3′ Melissa officinalis

R-Actin 5′-GCTCGGCAGTTGTGGTAA-3′

F-HPPR 5′-GCGCTGCCGAAATTGGAGAT-3′ Perilla frutescens

R-HPPR 5′-CGTTTCTGGAGTCAGCGCACA-3′

Chapter 3

Results

3.1. Pesticidal potential of selected plants against target pests The seven plant species, Cinnamomum camphora (L.) J. Presl (Lauraceae), Eucalyptus sideroxylon A. Cunn. ex Woolls (Myrtaceae), Isodon rugosus Wall. ex Benth (Labiatae), Boenninghausenia albiflora (Hook.) Rchb.ex Meisn. (Rutaceae), Calotropis procera Aiton (Dryand).(Apocynaceae), Daphne mucronata Royle (Thymelaeaceae), Tagetes minuta L.(Asteraceae) were analyzed against target insects belonging to different insect orders i.e. Bactrocera zonata Saunders, 1842, Schizaphis graminum Rondani, 1852, Drosophila melanogaster Meigen, 1830, Tribolium castaneum Herbst, 1797, Spodoptera exigua Hubner,1808 and Acyrthosiphon pisum Harris, 1776.

3.1.1. Bactrocera zonata Saunders, 1842

3.1.1.1. Adult mortality Male mortality between treatments was significantly different (F= 5.79, P>0.001). Five treatments, DDVP and T. minuta, I. rugosus, E. sideroxylon and D. mucronata extracts, gave similar but significantly higher mortality than all other treatments (Table 3.1). Female mortality was significantly higher with GF 120 treatment than all other treatments (Table3.1).

Table 3.1: Mean percentage mortality of males and females of Bactrocera zonata exposed to methanolic extracts of different plants in female protein baits and male lures under laboratory conditions. Mortality (female) Mortality (male) Percentage Mean Percentage Mean Pesticide/Plant extracts Mortality Mortality Mortality Mortality GF 120* 100 a 10±0 - DDVP* - 100 a 10±0.0 Tagetes minuta 6.6 b 0.7±0.6 73.3 a 7.3±2.5 Isodon rugosus 16.6 b 1.7±1.5 53.3 a 5.3±1.2 Daphne mucronata 13.3 b 1.3±0.6 50 a 5.0±1.7 Eucalyptus sideroxylon 3.3 b 0.3±0.6 50 a 5.0±2.6 Calotropis procera 0 b 0.0±0.0 43.3 b 4.3±2.5 Cinnamonum camphora 16.6 b 1.7±0.6 40 b 4.0±1.0 Boenninghausenia albiflora 10 b 1.0±1.0 26.6 b 2.7±1.5 Methanol* 0 b 0.0±0.0 - - Methyl eugenol* - - 13.3 b 1.3±0.6 Untreated 0 b 0.0±0.0 20 b 2.0±1.7

*GF 120, positive control for female toxicity bioassay; DDVP, positive control for male toxicity bioassay; methanol, negative control for female toxicity bioassay; methyl eugenol, negative control for male toxicity bioassay. Means followed by the same letter within a colomn are not significantly different (Tukey’s test, P< 0.05). 3.1.1.2. Effect of treatments on settlement and repellence behavior of females Mean number of females settling on untreated guavas was greater than on treated guavas (F= 5.79, P>0.001). The minimum number of females (3.3 of 15) that settled on any treated guavas was on those treated with D. mucronata extract, followed by those treated with B. albiflora (3.7 of 15), I. rugosus (4 of 15) and C. camphora (4.7 of 15) extracts. Both methanol and C. procera extract treated guavas had 5 of 15 females settle. With E. sideroxylon and T. minuta extracts 5.3 and 6.3 of the 15 females settled, which was also less than the 8.3 females that settled on untreated guavas (Fig. 3.1).

Figure 3.1: Mean percent repellence (%) caused by selected plant extracts at 2% concentration against fruit flies, Bactrocera zonata, under laboratory conditions 3.1.1.3. Oviposition deterrence

3.1.1.3.1. Effect of plant extracts on recovery of pupae There was significant treatment effect on number of pupae recovered from guavas. (F= 5.15, P= 0.001) (Table 3.2). The lowest number of pupae (3.3) was obtained from guavas treated with T. minuta extract, followed by those treated with D. mucronata and E. sideroxylon. Treatment with B. albiflora, C. camphora, C. procera and I. rugosus extracts resulted in recovery of about 30 to 40 pupae. The highest number of pupae was recovered from untreated guavas.

3.1.1.3.2. Effect of plant extracts on adult emergence Paralleling the results for pupae, the lowest number of adults emerged from guavas treated with T. minuta extract, followed by those treated with D. mucronata and E. sideroxylon extracts (Table 3.2). Guavas treated with B. albiflora, C. camphora, C. procera and I. rugosus extracts had greater numbers, but the highest number of adults that emerged was for untreated guavas.

Table 3.2: Mean number of Bactrocera zonata pupae recovered and adults emerging for guava fruit treated with various plant extracts and exposed for oviposition for 48 h.

Plants Extracts Pupae Count Mean ± SD Adult Mean ± SD Emergence Tagetes minuta 3.3 a 2.6 ± 0.6 0.3 a 0.3 ± 0.6

Daphne mucronata 26.7 b 26.7 ± 23.3 7.0 a 26.0 ± 22.6

Eucalyptus sideroxylon 28.3 b 45.0 ± 15.1 13.0 b 7.0 ± 10.4

Boenninghausenia albiflora 31.7 c 31.6 ± 42.5 13.3 b 27.3 ± 45.6

Cinnamomum camphora 35.0 c 35.0 ± 30.8 16.7 b 13.0 ± 11.1

Calotropis procera 35.7 c 35.7 ± 40.7 27.3 c 29.7 ± 42.2

Isodon rugosus 39.7 c 39.7 ± 33.7 29.7 c 10.0 ± 17.3

Untreated 62.7 d 62.7 ± 11.6 45.3 d 1.7 ± 2.9

Means followed by the same letters within each column are not significantly different (Tukey’s test, P< 0.05).

3.1.2. Schizaphis graminum Rondani, 1852 Two bioassays were used to analyze the effect of plant extracts against Schizaphis graminum.

3.1.2.1. Toxicity bioassay In toxicity bioassay, aphids were exposed to leaf discs treated with selected plant crude methanolic extracts at 2%. Five replications were used for each treatment containing 10 aphids per replication. Time course mortality of aphids had been observed after 12, 24, 36 and 48 h. After 12 h, all the treatments had negligible toxic effect except C. procera plant extract and Karate (F=155.4, P=0.000). Calotropis procera depicted 34% mortality and was significantly different from all other treatments, while C. comphora, Tagetes minuta, E. sideroxylon, B. albifora, I. rugosus and D. mucronata gave negligible toxic effects. After 24 h, D. mucronta and C. procera gave similar toxic effects with 36 and 44% mortality respectively, whereas C. comphora, T. minuta, E. sideroxylon, B. albifora, I. rugosus and D. mucronata except Karate had negligible

79 bioactivity (F=59.8, P=0.000). After 36 h all the treatments gave low bioactivity except C. procera, D. mucronata and Karate (F=53.6, P=0.000). The positive control and C. procera showed similar toxic effects with 92 and 100% mortality respectively, followed by D. mucronata with 44% mortality. After 48 h all the treatments gave nigligible toxic effects except C. procera, D. mucronata and Karate (F=45.3, P= 0.000). Calotropis procera exhibited 96% mortality followed by D. mucronata that gave 54% mortality, while C. comphora, T. minuta, E. sideroxylon, B.albifora and I. rugosus had negligible toxic effects. There was nill or negligible mortality observed in aphids exposed to untreated and methanol treated leaf discs. In the whole bioassay only two plants, C. procera and D. mucronata gave significantly different and higher mortalities as compared to all the treatments and in case of positive control, Karate all the aphids exposed were died after 12 h of treatment (Table3.3).

Table 3.3: Time course mean percentage mortality of Schizaphis graminum exposed to methanolic extracts of different plants at 2% for 12, 24, 36 and 48 h under laboratory conditions in toxicity bioassay

Treatments 12 h 24 h 36 h 48 h

Untreated 0 a 0 a 0 a 0 a

Cinnamomum comphora 0 a 2 a 2 a 2 a

Tagetes minuta 0 a 2 a 6 a 8 a

Eucalyptus sideroxylon 0 a 2 a 8 a 10 a

Boenninghausenia albifora 2 a 6 a 10 a 16 a

Isodon rugosus 4 a 6 a 16 a 16 a

Daphne mucronata 12 a 36 b 44 b 54 b

Calotropis procera 34 b 44 b 92 c 96 c

Karate 100 c 100 c 100 c 100 c

Methanol 2 a 4 a 8 a 16 a

Means followed by the same letter within a colomn are not significantly different (Tukey’s test, P< 0.05).

3.1.2.2. Repellence bioassay In this bioassay, repellent effect of plant crude methanolic extracts at 2% were analyzed against S. graminum by exposing aphids to plant extract treated and untreated leaf discs simultaneously in each treatment. Five replications were used for each treatment containing 10 aphids per replication. Aphids were analyzed for their repellent behavior after every 12 h until 48 h. After 12 h, all the treatments had significantly different repellent effects (F=6.2, P=0.000). In Table 3.4, positive values indicate repellency and negative values indicates attraction. After 12 h, methanol control and synthetic pesticide, Karate depicted negative values. Maximum of 76% repellent effect was exhibited by D. mucronata followed by C. camphora, C. procera, B. albifora, T. minuta, I. rugosus and E. sidroxylon depicted 72, 56, 52, 40, 36 and 24 % repellent effect,

81 respectively. After 24 h, all the treatments gave significantly different repellence effect (F=12, P=0.000) except methanol and Karate control. Cinnamomum camphora repellent effect increased from 72% to 76% followed by I. rugosus depicting increase in repellent effect from 36 to 68% and repellency caused by B. albifora increased from 52 to 60%. In the case of B. albifora, repellent effect was increased after every 12 h and after 72 h it gave 68% repellency. After 36 h, there was no significant difference among all the treatments except Karate control (F=10.8, P= 0.000). Cinnamomum camphora caused highest repellent effect of 76% followed by B. albifora that exhibited 64% repellent behavior. Whereas, E. sideroxylon I. rugosus, D. mucronata, T. minuta and C. procera depicted less than 50% repellent effect. bAfter 48 h, there was no significant difference among all the treatments except Karate control (F=12.6, P=0.000). Boenninghausenia albifora, C. camphora, E. sideroxylon and I. rugosus gave more then 50% repellent effect. In case of D. mucronata and C. procera, repellency was reduce after every 12 h until 72 h because aphids died due to their strong toxic effects as described in toxicity bioassay (Table 3.4).

Table 3.4: Mean percentage repellence of Schizaphis graminum exposed to methanolic extracts of different plants for 12, 24, 36 and 48 h under laboratory conditions in repellence bioassay

Treatments 12 hr 24 hr 36 hr 48 hr Methanol -16 ab 4 b 12 b 12 b Karate -55 a -76 a -88 a -88 a Calotropis procera 56 bc 24 bc 20 b 12 b Daphne mucronata 76 c 64 bc 36 b 32 b Cinnamomum camphora 72 c 76 c 76 b 64 b Isodon rugosus 36 bc 68 c 44 b 52 b Tagetes minuta 40 bc 36 bc 32 b 24 b Boenninghausenia albifora 52 bc 60 bc 64 b 68 b Eucalyptus sideroxylon 24 abc 24 bc 48 b 52 b

Means followed by the same letter within a colomn are not significantly different (Tukey’s test, P< 0.05).

3.1.3. Tribollium castaneum Herbst, 1797

3.1.3.1. Impregnation bioassay There was no mortality observed when adults of Tribolium castaneum were exposed to filter papers treated with all plant extracts after 24, 48 and 72 h.

3.1.3.2. Flour discs bioassay There was no mortality observed when adults of Tribolium castaneum were fed on flour discs containing plant extracts for 24, 48 and 72 h in all the treatments.

3.1.4. Drosophila melanogaster Meigen, 1830

3.1.4.1. Contact toxicity In this bioassay, flies were exposed to artificial diet layered with 2% crude methanolic extracts in different treatments. Three replications, each replication with 10 flies were used for each treatment. Time course mortality was observed after 24, 48 and 72 h. Mortalities were corrected with Abbott’s formula (Abbott, 1925). After 24 h, there was no significant differences among all the treatments (F=24.0, P=0.000) except C. camphora that exhibited 30.7% mortality. Daphne mucronata, B. albiflora, E.

83 sideroxylon, I. rugosus, T. minuta and C. procera had no toxic effects and were significantly different from C. camphora. After 48 h, there was no significant difference (F=54.4, P=0.000) among all treatments except C. camphora that gave 34.3% mortality. After 72 h, there was no significant difference (F=64.2, P=0.000) among D. mucronata, B. albiflora, E. sideroxylon, I. rugosus, T. minuta and C. procera, depicting negligible toxic effects except C. camphora that exhibited higher mortality of 41.3% (Table 3.5)

Table 3.5: Time course mean percentage mortality of Drosophila melanogaster exposed to methanolic extracts of different plants at 2% for 24, 48 and 72 h under laboratory conditions

Treatments 24 h 48 h 72 h

Daphne mucronata 0.0 a 0.0 a 0.0 a

Boenninghausenia albiflora 0.0 a 0.0 a 0.0 a

Eucalyptus sideroxylon 0.0 a 0.0 a 0.0 a

Isodon rugosus 0.0 a 0.0 a 0.0 a

Tagetes minuta 0.0 a 0.0 a 3.3 a

Calotropis procera 3.3 a 7.0 a 7.0 a

Cinnamomum camphora 30.7 b 34.3 b 41.3 b

Means followed by the same letter within a colomn are not significantly different (Tukey’s test, P< 0.05).

3.1.5. Spodoptera exigua Hubner, 1808

3.1.5.1. Contact toxicity In this bioassay, second instar larvae of S. exigua were exposed to artificial diet layered with 2% crude methanolic extracts in different treatments. 20 insects were used for each treatment. Time course mortalities were observed after 24, 48 and 72 h. Mortalities were corrected with Abbott’s formula (Abbott, 1925). After 24 h, T. minuta and C. procera gave 6% and 11% mortality respectively, while there was no mortality observed in case of larvae exposed to D. mucronata, B. albiflora, E. sideroxylon, I.

84 rugosus, C. camphora treated diets. After 48 h, T. minuta, E. sideroxylon, I. rugosus, D. mucronata and C. procera gave 6, 6, 6, 6 and 22% mortality respectively. While there was no mortality observed in case of larvae exposed to C. camphora and B. albifora treated diets. After 72 h, all the treatments gave mortalities except B. albifora. Throughout the bioassay until 72 h, no mortality was observed in case B. albifora treated diet. After 72 h of bioassay, T. minuta, E. sideroxylon, I. rugosus, D. mucronata and C. procera depicted 6% mortality while C. procera gave 22% mortality. So, there was no further mortality in all the treatments after 48 h of bioassay (Table 3.6).

Table 3.6: Time course mean percentage mortality of Spodoptera exigua exposed to methanolic extracts of different plants for 24, 48 and 72 h under laboratory conditions

Treatments 24 h 48 h 72 h

Tagetes minuta 6 6 6

Cinnamomum camphora 0 0 6

Eucalyptus sideroxylon 0 6 6

Boenninghausenia albiflora 0 0 0

Calotropis procera 11 22 22

Isodon rugosus 0 6 6

Daphne mucronata 0 6 6

Mean of 20 insects

3.1.6. Acyrthosiphon pisum Harris, 1776

3.1.6.1. Feeding toxicity bioassay In this bioassay, aphids were exposed to artificial diet mixed with different concentrations (1000, 500, 200, 100 and 50 ppm) and mortality was observed after 24 of treatment. Except for T. minuta and C. procera with negligible toxic effects (no LC50 and

LC90 could be calculated with the concentrations tested), all of the other plants analyzed showed strong toxic effects against A. pisum after 24 h of exposure to their respective

85 methanolic extracts (Table 3.7). The crude extract from I. rugosus caused the highest mortality (LC50 36.2 ppm and LC90 102.1 ppm) as compared to the other plant extracts tested. Although the crude extract from D. mucronata caused the second highest mortality (LC50 126.2 ppm and LC90 197.5 ppm), this was not significantly different from

E. sideroxylon (LC50 136.2 ppm and LC90 374.2 ppm) and B. albiflora (LC50 160.3 ppm and LC90 379.5 ppm) since their 95% confidence limit values overlap with each other (Table 3.7). I. rugosus and D. mucronata were selected for further fractionation and evaluation against aphids.

Table 3.7: Toxicity of crude methanolic extracts against newborn (˂ 24 hour old) Acyrthosiphon pisum nymphs following 24 h exposure to artificial diet containing different concentrations of crude extracts

Crude extracts LC50 (95% CI) ppm Ratio LC90 (95% CI) ppm Ratio Slope ± Chi- HF SE Square

Isodon rugosus 36.2 (18-49) a 1.0 102.1 (80.4-155.7) a 1.0 2.8 ± 0.8 8.1 0.6

Daphne mucronata 126.2 (112.5 -141.6) b 3.5 197.5 (171.3-247.6) b 1.9 6.6 ± 1.2 3.9 0.3

Cinnamomum camphora 624.1 (505.3-813.2) c 17.2 2002 (1380-3702.6) c 19.6 2.5 ± 0.4 4.1 0.3

Eucalyptus sideroxylon 136.2 (113-162.8) b 3.8 374.2 (294-528.3) d 3.7 2.9 ± 0.4 4.5 0.3

Boenninghausenia albiflora 160.3 (136.4-190) b 4.4 379.5 (302-530.5) d 3.7 3.4± 0.5 8.2 0.6

Tagetes minuta - - - - 0.7 ± 0.6 10.1 0.8

Calotropis procera - - - - 2.4 ± 1.0 2.2 0.2

Data is presented as 50% (LC50) and 90% (LC90) values of lethal concentration (both in ppm) along with their respective 95% confidence interval (95% CI), the slope ± SE of the toxicity vs concentration curve, and the Chi-Square and heterogeneity factor HF as accuracy of data fitting to probit analysis in POLO-PlusV2. Different letters in the same column indicate significant differences due to non-overlapping of 95% CI. Ratio, LCx, crude extract/LCx, Isodon rugosus

3.2. Isolation of active compounds from the selected plants responsible for pesticidal activity On the basis of previous bioassays with seven plant crude methanolic extracts with all target insects, two most active plants i.e. Daphne mucronata and Isodon rugosus were selected to evaluate the toxicity of different fractions of their respective extracts against most susceptible insect, Acyrthosiphon pisum among all the target insects analyzed.

3.2.1. Daphne mucronata fractionation Through liquid-liquid extraction, four solvent fractions of Daphne mucronataon polarity basis were obtained i.e. black green n-hexane residue (41.84 g), a shiny dark green dichloromethane residue (2.94 g), a brownish green ethyl acetate residue (4.95 g), and a brown n-butanol residue (44.79 g)

3.2.1.1. Bioactivity of fractions When aphids were exposed for 24 h to different concentrations of solvent fractions treated diet, the highest mortality in the bioassays was found for the ethyl acetate (LC50 67.8 ppm and LC90 156.9 ppm) and dichloromethane fractions (LC50 63.1 ppm and LC90 157.8 ppm) (Table

3.8). The second most active fraction was the butanol fraction (LC50 449.5 ppm and LC90

1472.1ppm), while the lowest mortality was observed with the hexane fraction (LC50 685.4 ppm and LC90 3557.3 ppm) (Table 3.8).

Table 3.8: Toxicity of Daphne mucronata solvent fractions against newborn (˂ 24 hour old) Acyrthosiphon pisum nymphs following 24 h exposure to artificial diet containing different concentrations of solvent fractions Fractions LC50 (95% CI) ppm Ratio LC90 (95% CI) ppm Ratio Slope ± SE Chi-Square HF

Hexane 685.4 (451.3-1541) a 10.8 3557.3 (1571.3-21021) a 22.6 1.8 ± 0.4 6.6 0.5

Butanol 449.5 (347.7-667.9) a 7.1 1472.1 (910-3624.1) a 9.3 2.5 ± 0.5 3.4 0.3

Ethyl acetate 67.8 (58-79.3) b 1.1 156.9 (126.9-214.6) b 1.0 3.5± 0.5 8.1 0.6

Dichloromethane 63.1 (53.3-74.4) b 1.0 157.8 (125.8-220.5) b 1.0 3.2 ± 0.5 8.8 0.7

Ratio, LCx, fraction/LCx, dichloromethane

3.2.2. Isodon rogusus fractionation Four solvent fractions were obtained through liquid-liquid extraction of crude methanolic extract of I. rugosus. Theses fractions were black green n-hexane residue (8.82 g), a shiny green dichloromethane residue (3.1 g), a light green ethyl acetate residue (1.33 g) and a yellow n-butanol residue (6 g).

3.2.2.1. Bioactivity of fractions When aphids were exposed to different concentrations of sovent fractions treated diet for 24 h, the butanol fraction caused the highest mortality in aphids at a low concentration (LC50 18 ppm and LC90 48.2 ppm) after 24 h of exposure (Table 3.9). The ethyl acetate fraction (LC50 17.5 ppm and LC90 96 ppm) and dichloromethane fraction

(LC50 54.5 ppm and LC90 146.2 ppm) depicted comparable mortality at their LC90 and not

LC50 (Table 3.9). The lowest mortality was obtained for the hexane fraction (LC50 165.2 ppm and LC90 533.4 ppm) after 24 h of treatment (Table 3.9).

Table 3.9: Toxicity of Isodon rugosus solvent fractions against newborn (˂ 24 hour old) Acyrthosiphon pisum nymphs following 24 h exposure to artificial diet containing different concentrations of solvent fractions

Fractions LC50 (95% CI) ppm Ratio LC90 (95% CI) ppm Ratio Slope ± SE Chi-Square HF

Hexane 165.2 (125.6 -231) a 9.4 533.4 (348.2-1150.5) a 11.1 2.5 ± 0.4 22.5 1.7

Butanol 18 (8.5-23.9) b 1.0 48.2 (38.1-73.5) b 1.0 2.9 ± 0.9 4.7 0.4

Ethylacetate 17.5 (7.1-26.2) b 1.0 96 (69-163.9) bc 2.0 1.7 ± 0.4 5.2 0.4

Dichloromethane 54.5 (44.8-64.3) c 3.1 146.2 (115.2-208.9) c 3.0 3.0± 0.5 5.4 0.4

Ratio, LCx, fraction/LCx, butanol or ethyl acetate

3.2.3. First reverse phase automatic flash chromatography of 500 mg butanol fraction of I. rugosus Butanol fraction of I. rugosus was more active with low LC’s values among all the analyzed eight solvent fractions of both D. mucronata and I. rugosus. Therefore, the butanol fraction of I. rugosus was sub-fractionated to find the bioactive insecticidal compound against A.pisum. The butanol fraction, consisting of 500 mg of material, was sub-fractionated through reverse phase automatic flash chromatography. On the basis of UV chromatogram at 220 nm, 95 collected fractions were combined into 14 fractions (1A- 14A) (Fig. 3.2). The obtained sub-fractions are shown in the table 3.10.

Table 3.10: Sub-fractions collected from first reverse phase automatic flash chromatography

Fractions Weight (mg)

1A 52

2A 10.6

3A 45.5

4A 7.3

5A 19.7

6A 14.9

7A 54.1

8A 57.9

9A 14.2

10A 48.5

11A 18

12A 15.3

13A 20.7

14A 18

Figure 3.2: UV chromatogram of first reverse phase automatic flash chromatography of 500 mg butanol sample. UV1 wavelength: 220 nm, UV2 Wavelength: 254 nm 3.2.3.1. Bioactivity of sub-fractions of butanol fraction of I.rugosus Bioactivity of all of the fourteen sub-fractions (1A-14A) obtained through first reverse phase automatic flash chromatography of 500 mg of butanol fraction of I. rugosus was analyzed for 24 h against A. pisum. Except fractions 8A, 9A, 11A, 13A and 14A, all the other fractions showed considerable toxic effects against A. pisum. Fraction 3A (LC50

2.1 ppm and LC90 29.5 ppm) had highest activity as compared to all other fractions, followed by fraction 5A (LC50 3.3 ppm and LC90 50 ppm). Fraction, 1A (LC50 5.5 ppm and LC90 66 ppm), 2A (LC50 8.9 ppm and LC90 81.1 ppm), 4A (LC50 6.8 ppm and LC90

112.2 ppm) and 6A (LC50 17.8 ppm and LC90 187.3 ppm) gave considerable mortality.

Fraction 7A (LC50 74.1 ppm and LC90 267 ppm) showed lower mortality. Moderate toxicity was observed with fraction, 10A (LC50 36.2 ppm and LC90 52.5 ppm) and 12A

(LC50 51.4 ppm and LC90 109 ppm) (Table 3.11).

Table 3.11: Toxicity of sub-fractions of butanol fraction from first reverse phase automatic flash chromatography against newborn (˂ 24 hour old) Acyrthosiphon pisum nymphs following 24 h exposure to artificial diet containing different concentrations of solvent fractions

Fractions LC50 (95% CI) ppm Ratio LC90 (95% CI) ppm Ratio Slope ± SE Chi-Square HF

1A 5.5 (3-8) a 2.6 66 (37-210.6) a 2.2 1.1 ± 0.3 7.1 0.5 2A 8.9 (6.1-12.1) a 4.2 81.1 (46.5-230.5) a 2.7 1.3 ± 0.3 5.6 0.4 3A 2.1 (0.6-3.8) a 1.0 29.5 (18-85.3) a 1.0 1.1± 0.3 7.5 0.6 4A 6.8 (3.8-10.1) a 3.2 112.2 (54-561.1) a 3.8 1.1 ± 0.3 4.6 0.4 5A 3.3 (1.3-5.4) a 1.6 50 (28-176.4) a 1.7 1.1 ± 0.3 10.1 0.8 6A 17.8 (12.9-26.7) b 8.5 187.3 (89.7-807.7) a 6.3 1.3 ± 0.3 3.8 0.3 7A 74.1 (51.5-169.2) c 35.3 267 (130.5-1651) a 9.1 2.3 ± 0.6 8.1 0.6

8A - - - - 1.7 ± 0.7 7.0 0.5

9A - - - - 2.0 ± 1.3 4.7 0.4

10A 36.2 (32.6-40.3) d 17.2 52.5 (46.4-63.6) a 1.8 8.0 ± 1.4 2.8 0.2

11A - - - - 1.6 ± 0.6 8.5 0.7

12A 51.4 (42.8-71) c 24.5 109 (76.8-240.5) a 3.7 3.9 ± 1.0 2.2 0.2

13A - - - - 1.5 ± 1.2 6.6 0.5

14A - - - - 2 ± 1.3 4.7 0.4

Ratio, LCx, fraction/LCx, 3A

3.2.4. Sub-fractionation of 3A fraction through prep-LC Out of fourteen fractions (1A-14A) of the first reverse phase automatic flash chromatography, fraction 3A was the most bioactive fraction with lower LC values as compared to all other fractions analyzed. Therefore, fraction 3A was further fractionated into three fractions through prep-LC, 3A-1, 3A-2 and 3A-3.

3.2.4.1. Bioactivity of sub-fractions of fraction 3A collected through prep-LC All the collected three sub-fractions (3A-1, 3A-2 and 3A-3) of 3A were analyzed against A. pisum for 24 h. Fraction 3A-1 and fraction 3A-2 gave negligible toxic effects

(no LC50 and LC90). Fraction 3A-3 was the most toxic fraction analyzed against A. pisum with low LC’s (LC50 1 ppm and LC90 13.8 ppm) (Table 3.12).

Table 3.12: Toxicity of sub-fractions of fraction 3A against newborn (˂ 24 hour old) Acyrthosiphon pisum nymphs following 24 h exposure to artificial diet containing different concentrations of solvent fractions

Fractions LC50 (95% CI) ppm Ratio LC90 (95% CI) ppm Ratio Slope ± SE Chi-Square HF

3A-1 - - - - 2 ± 1.3 4.9 0.4

3A-2 - - - - 1.5 ± 1.2 6.6 0.5

3A-3 1 (0.6-1.6) a 1 13.8 (6.1-97) a 1 1.1± 0.3 14.8 1.1

Ratio, LCx, fraction/LCx, 3A-3

3.2.4.2. Spectroscopic analysis of fraction 3A-3 Out of three sub-fractions of 3A (3A-1, 3A-2 and 3A-3), fraction 3A-3 was the most bioactive fraction against A. pisum. This fraction 3A-3 was analyzed through 1H NMR which confirmed that the bioactive fraction 3A-3 was rosmarinic acid. As different gradients were used to purify rosmarinic acid through prep-LC, chromatographic problem of peak shifting in each cycle was observed (Fig. 3.3) which complicated further purification at this stage. Therefore, reverse phase automatic flash chromatography was repeated with 5 g of butanol fraction of I. rugosus.

Intensity

Retention time (a)

Intensity

Retention time (b)

Intensity

Retention time (c)

Figure 3.3: (a), (b), (c) Prep-LC chromatograms of three sub-fractions (3A-1, 3A-2 and 3A-3) of fraction 3A, indicating different peak appearance of rosmarinic acid between 40 to 70 min in different cycles of prep-LC.

3.2.5. Sub-fractionation of 5 g butanol fraction of I. rugosust hrough second reverse phase automatic flash chromatography Through second reverse phase automatic flash chromatography 5 g of butanol fraction of I. rugosus was fractionated into 354 fractions. On the basis of UV spectra at 220 nm, 354 collected fractions were combined into 6 fractions (1B-6B) (Fig. 3.4). These fractions are given in table 3.13.

Table 3.13: Fractions from second reverse phase automatic flash chromatography

Fractions Weight (mg)

1B 530.4

2B 830

3B 1522.6

4B 195

5B 140

6B 128

Figure 3.4: UV chromatogram of second reverse phase automatic flash chromatography of 5 g of butanol fraction. UV1 Wavelength: 220 nm, UV2 Wavelength: 254 nm.

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3.2.5.1. Bioactivity of sub fractions of butanol fraction from second reverse phase automatic flash chromatography Six fractions (1B-6B) obtained through second reverse phase automatic flash chromatography of the butanol fraction of I. rugosus, were analyzed against A. pisum for 24 h. Out of six fractions analyzed, fraction 4B, 5B, and 6B showed negligible toxicity

(no LC50 and LC90). Fraction 1B was more toxic (LC50 2.5 ppm and LC90 28.2 ppm) and moderate toxicity was observed by fraction 2B (LC50 7.5 ppm and LC9071.1 ppm).

Lower toxicity was depicted by fraction 3B (LC50 16.3 ppm and LC90 100.8 ppm) (Table 3.14).

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Table 3.14: Toxicity of sub-fractions of the butanol fraction from second reverse phase automatic flash chromatography against newborn (˂ 24 hour old) Acyrthosiphon pisum nymphs following 24 h exposure to artificial diet containing different concentrations of solvent fractions

Fractions LC50 (95% CI) ppm Ratio LC90 (95% CI) ppm Ratio Slope ± SE Chi-Square HF

1B 2.5 (1-4.1) a 1.0 28.2 (17.8-69.3) a 1 1.2 ± 0.3 11.4 0.9

2B 7.5 (4.3-11) b 3.0 71.1 (38-280.4) a 2.5 1.3 ± 0.3 16.5 1.3

3B 16.3 (11.2-25.5) c 6.5 100.8 (52.3-417.4) a 3.6 1.6± 0.3 22.3 1.7

4B - - - - 1 ± 0.3 25.3 2.0

5B - - - - 1.5 ± 1.2 6.6 0.5

6B - - - - 1.8 ± 0.7 6.5 0.5

Ratio, LCx, fraction/LCx, 1B

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3.2.6. Acidic extraction of most bioactive fraction 1B from second reverse phase automatic flash chromatography The most active fraction 1B was further subjected to acidic extraction. After acidic extraction two phases, i.e. 60 mg of ethyl acetate phase and 60 mg of water phase were collected.

3.2.6.1. Bioactivity of ethyl acetate and water phase of acidic extraction Both collected phases of acidic extraction were analyzed for their pesticidal potential through bioassay against A. pisum for 24 h. Water phase gave negligible toxic effect (no LC50 and LC90) while ethyl acetate phase showed more toxicity (LC50 0.2 ppm and LC90 9.2 ppm) (Table 3.15).

103

Table 3.15: Toxicity of ethyl acetate and water phase of acidic extraction against newborn (˂ 24 hour old) Acyrthosiphon pisum nymphs following 24 h exposure to artificial diet containing different concentrations of both phases

Fractions LC50 (95% CI) ppm Ratio LC90 (95% CI) ppm Ratio Slope ± SE Chi-Square HF

Water - - - - 1.5 ± 1.2 6.6 0.5

Ethyl acetate 0.2 (0.04-0.5) a 1 9.2 (3.9-130.7)a 1 0.8 ± 0.3 4.2 0.3

Ratio, LCx, fraction/LCx, ethyl acetate

104

3.2.7. Identification of most active bioactive compound Out of the two phases of acidic extraction, the ethyl acetate phase fraction was the most bioactive. This fraction was further purified, analyzed and identified as rosmarinic acid through HPLC-MS, optical rotation measurement and 1H and 13C NMR spectroscopy.

3.2.7.1. HPLC-MS Both isolated and commercial rosmarinic acid from Sigma Aldrich had the same peak appearance in the HPLC-MS chromatograms with the same solvent gradient. Both had a pseudomolecular ion with an m/z value of 359 with negative mode electrospray

ionization which confirmed that it was rosmarinic acid (Fig. 3.5).

Intensity

Molecular mass (a)

105

Intensity

Molecular mass (b) Figure 3.5: Mass spectra (negative mode electrospray ionization) of rosmarinic acid obtained via HPLC-MS with a pseudo molecular ion at m/z value of 359 (a) Isolated rosmarinic acid (b) Commercial rosmarinic acid 3.2.7.2. 1H and 13C NMR

24 1 Brown crystals; [α]D +78,0° (c 0.233, MeOH); H NMR (400 MHz, CD3OD): δ 3.01 (1H, dd, J = 8.3, 14.3 Hz, H7a), 3.10 (1H, dd, J = 4.4, 14.3 Hz, H7b), 5.19 (1H, dd, J = 4.4, 8.3 Hz, H8), 6.27 (1H, d, J = 15.9, H17), 6.61 (1H, dd, J = 2.0, 8.0 Hz, H6), 6.70 (1H, d, J = 8.0 Hz, H5), 6.75 (1H, d, J = 2.0 Hz, H2), 6.78 (1H, d, J = 8.2 Hz, H14), 6.95 (1H, dd, J = 2.0, 8.2 Hz, H15), 7.04 (1H, d, J = 2.0 Hz, H11), 7.55 (1H, d, J = 15.9 Hz, 16 13 7 8 17 11 H ); C NMR (100 MHz, CD3OD): δ 37.9 (C ), 74.6 (C ), 114.4 (C ), 115.2 (C ), 116.3 (C5), 116.5 (C14), 117.6 (C2), 121.8 (C6), 123.2 (C15), 127.7 (C10), 129.2 (C1), 145.3 (C4), 146.2 (C3), 146.8 (C12), 147.7 (C16), 149.7 (C13), 168.4 (C18), 173.5 (C9); ESI-MS

106

(70 eV): 359.0 (M-H+). NMR signals were in accordance with the literature (Dapkevicius et al., 2002; Hyun et al., 2015) (Fig. 3.6).

Figure 3.6: Structure of rosmarinic acid isolated from I. rugosus 3.3. Characterization of the I. rugosus rosmarinic acid for their pesticidal activity using different fractions

3.3.1. Bioactivity of I. rugosus rosmarinic acid and commercial rosmarinic acid Rosmarinic acid isolated from I. rugosus and commercial rosmarinic acid from Sigma Aldrich were analyzed against A. pisum for their pesticidal activity for 24 h. Both,

I. rugosus RA (LC50 0.2 ppm and LC90 5.4 ppm) and commercial RA (LC50 0.2 ppm and

LC90 14 ppm) gave similar toxic effects (Table 3.16).

107

Table 3.16: Toxicity of isolated rosmarinic acid (RA) and commercial rosmarinic acid (RA) against newborn (˂ 24 hour old) Acyrthosiphon pisum nymphs following 24 h exposure to artificial diet containing different concentrations of isolated rosmarinic acid and commercial rosmarinic acid

Fractions LC50 (95% CI) ppm Ratio LC90 (95% CI) ppm Ratio Slope ± SE Chi-Square HF

Commercial RA 0.2(0.05-0.5) a 1 14 (7.4-41.8) a 2.6 0.7 ± 0.2 15.5 0.7

I. rugosus RA 0.2 (0.04-0.4) a 1 5.4 (3.3-11.5) a 1 0.8 ± 0.2 10.5 0.5

Ratio, LCx, fraction/LCx, Isodon rugosus RA

108

3.3.2. Comparison between growth of live aphids exposed to plant extract treated and untreated diet after 24 h of bioassay After incorporating the rosmarinic acid in diet, its effect on A. pisum was analyzed everyday for up to 9 d. It was confirmed that rosmarinic acid had drastic effect on its growth. Firstly, aphids exposed to treated diet were died but if survived they were not grow further to become adults and were thus not able to reproduce further. Figure 3.8 shows a comparison between treated and untreated aphids. It was observed that there was a clear difference between untreated and treated aphid after day 4 and at day 9 treated aphid was died while untreated was still alive as shown in the figure 3.7.

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Figure 3.7: Comparison between growth of live aphids exposed to plant extract treated and untreated diet after 24 h of bioassay, (a) to (i) comparison observed for upto 9 days, treated aphid died on day 9 3.4. Identification of genes involved in the synthesis of rosmarinic acid in Isodon rugosus In the biosynthesis of rosmarinic acid, eight genes have been involved as described in Coleus blumei by many researchers (Petersen et al., 1993, 1994; Kim et al., 2004; Eberle et al., 2009). In this study, two keys genes; rosmarinic acid synthase gene

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(RAS) and hydrooxyphenylpyruvate reductase gene (HPPR) of this biosynthetic pathway were targeted for cloning from I. rugosus. 3.4.1. PCR amplification of rosmarinic acid synthase gene (RAS) Three sets of primers were used for amplification of rosmarinic acid synthase gene from I. rugosus. Gradient PCR was performed by using 10 different primer annealing temperatures between 50◦C to 60◦C under specified conditions. 3.4.1.1. Amplification with first set of primers With first set of primers, F-RAS15′- ATTACATATGAAGATAGAAGTCAAAGACTC-3′and F-RAS1 5′- TAGGATCCTCATCAAATCTCATAAAACAACTTCTC-3′, there was no amplification observed for rosmarinic acid synthase gene after analyzing gel by Bio-Rad’s gel documentation system.

3.4.1.2. Amplification with second set of primers Amplification with second set of primers, F-RAS2 5′- AAGGGAATTTCCACGTACCC-3′ and R-RAS2 5′-ACCCAGCTAATCACCCACAA- 3′,through gradient PCR from 50◦C to 60◦C primer annealing temperature resulted in multiple bands on gel after observing by Bio-Rad’s gel documentation system. Expected size of the band was 786 bp and this band was sequenced after purification through Wizard® SV Gel and PCR Clean-Up System (Promega). There was no sequence observed for RAS after doing BLAST for the obtained sequence through NCBI Gene bank (Fig. 3.8 and 3.9).

Figure 3.8: Multiple bands appearence for RAS, required band should be at 786 bp, M= Molecular weight marker

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Figure 3.9: Purified band for RAS at 786 bp, M= Molecular weight marker 3.4.1.3. Amplification with third set of primers Amplification with third set of primers, F-RAS35′- GACGAAGCTCCACATCCCCTTC-3′and R-RAS3 5′- GCGCTCCATATGCTGCGTGT-3′, through gradient PCR using 50◦C to 60◦C annealing temperatures results in multiple bands on gel after observing by Bio-Rad’s gel documentation system. All the bands from the gel were purified separately through Wizard® SV Gel and PCR Clean-Up System (Promega), and were sequenced. There was no sequence observed for RAS in case of all purified bands (Fig. 3.10).

Figure 3.10: PCR products (multiple bands) for RAS in gradient PCR using 10 annealing temperatures between 50◦C to 60◦C, M= Molecular weight marker

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3.4.2. PCR amplification of hydrooxyphenylpyruvate reductase (HPPR) Amplification of HPPR was performed by using a set of primers, F-HPPR 5′- GCGCTGCCGAAATTGGAGAT-3′ and R-HPPR 5′- CGTTTCTGGAGTCAGCGCACA-3′ at 56◦C primer annealing temperature. PCR amplification was observed at 438 bp after analyzing the gel by Bio-Rad’s gel documentation system. This band was purified with Wizard® SV Gel and PCR Clean- Up System and sequenced. It was confirmed from the obtained sequence that the amplified product was sequence of HPPR from I. rugosus (sHPPR) (Fig.3.11).

Figure 3.11: Purified PCR product for sHPPR at 438 bp from I. rugosus, M= Molecular weight marker 3.4.2.1. sHPPR sequence (HPPR from I. rugosus) >sHPPR GGTCGACTTAATCAAGTGTAAGGAGAAGGGGATTAGGGTTACCAACACGCCAGATG TGCTGACCGATGACGTCGCGGATTTGGCGATCGGGTTGATGTTGGCGGTTTTGAGGC GGATTTGTGAGTGTGATAAGTATGTGAGGAGGGGGGCGTGGAAACTTGGTGACTTCA AGTTGACGACTAAGTTCAGTGGCAAAAGAGTTGGCATCATAGGATTGGGCAGAATTG GGTTAGCAATTGCTGAGCGCGCAGATGCATTTGATTGTCCCATCAGTTACTACTCAA GATCCGAGAAAACCTACACGAACTACAAGTACTATGACAGCGTTGTTGAGTTGGCAT CAAACAGTGACATCTTAGTGGTAGCATGTGCGCTGACTCCAGAAACG

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3.4.2.2. sHPPR phylogenetic tree NCBI blast analysis and alignment in Clustal omega of sHPPR and HPPR in other plant species confirmed the identity of sHPPR as HPPR. A phylogenetic tree was also generated to show the relatedness of sHPPR to HPPR in other plant species (Fig. 3.12).

gi|448278785|gb|JX566894.1| Salvia officinalis hydroxyphenylpyruvate reductase mRNA complete cds

gi|72256934|gb|DQ099741.1| Salvia miltiorrhiza putative hydroxyphenylpyruvate reductase (hppr) mRNA complete cds

gi|62816283|emb|AJ507733.2| Solenostemon scutellarioides mRNA for hydroxyphenylpyruvate reductase (hppr gene)

gi|305379591|gb|HM587131.1| Perilla frutescens hydroxyphenylpyruvate reductase mRNAcomplete cds

sHPPR

Figure 3.12: Phylogenetic tree for sHPPR (HPPR from I. rugosus) with HPPR in other plant species; Perilla frutescens, Solenostemon scutellarioides, Salvia officinalis and Salvia miltiorrhiza.

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3.4.2.3. Alignment of amplified sHPPR with the conserved region amplified from other reported plant species

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3.4.3. 3′ and 5′ RACE PCR for sHPPR For full length amplification of sHPPR, RACE PCR was performed. For 3′ and 5′RACE PCR, 3′ and 5′RACE cDNAs were synthesized using SMARTer™ RACE cDNA Amplification Kit from Clontech and amplification was done by using the I. rugosus gene specific primers for sHPPR according to the manufacturer’s protocol. After PCR, PCR samples were run on gel in 1.5X TE buffer for 30 min. After the gel electrophoresis the gels were observed by Bio-Rad’s gel documentation system. The both 3′ and 5′PCR products were purified from the gels with Wizard® SV Gel and PCR Clean-Up System and were sequenced (Fig. 3.13).

Figure 3.13: Purified 5′ and 3′ RACEPCR products for sHPPR, M= Molecular weight marker 3.4.3.1. 3′RACE PCR product sequence for sHPPR >3′RACE AAGTATGTGAGGAGGGGGGCGTGGAAACTTGGTGNNTTCAAGTTGACGACTA AGTTCAGTGGCAAAAGAGTTGGCATCATAGGATTGGGCAGAATTGGGTTAGC AATTGCTGAGCGCGCAGATGCATTTGATTGTCCCATCAGTTACTACTCAAGAT CCGAGAAAACCTACACGAACTACAAGTACTATGACAGCGTTGTTGAGTTGGC ATCAAACAGTGACATCTTAGTGGTAGCATGTGCCCTGACTCC

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3.4.3.2. 5′RACE PCR product sequence for sHPPR >5′RACE CTGAACTTAGTCGTCAACTTGAAGTCACCAAGTTTCCACGCCCCCCTCCTCAC ATACTTATCACACTCACAAATCCGCCTCAAAACCGCCAACATCAACCCGATC GCCAAATCCGCGACGTCATCGGTCAGCACATCTGGCGTGTTGGTAACCCTAA TCCCCTTCTCCTTACACTTGATTAAGTCGACCTTATCCAGACCTACGCTATAG CTCGAAACTATCTCCAATTTCGGCAGTGCATCGATCAGCTCGGCGTCGGCGCC GGCGGTGGCGTTTCCGACCACCGCGCGGATGGACTCGGCCTGCCGAGTGAGA AACTCGCGCTGCGCCGGCTGAGCCCAGTAACGGAAGAGCTTGAACCGCTTGT CGAGCTCTTGCTCCAAGTAGTTGCTCATCGGGCACAACATCAGAACACCGAT CGCCTCCATTTTAACTGCAGCAGCGGCGTCGGCGTCGGCGGCGGTGGCGGGG GATGGAGATGGTGTTTGAGTGGTAGTTGAGA

3.4.4. Expression of HPPR in leaves, roots and stems of I. rugosus HPPR gene was amplified from, roots, leaves and stems of I. rugosus to confirm its presence in these parts of the plant through PCR at 56C primers annealing temperature. Actin gene was also amplified from these parts of the plant, used as a control. RNA from all these parts was extracted on three different days and used for this analysis. Amplification of HPPR gene in leaves, roots and stems has confirmed its expression in these parts of the plant, I. rugosus (Fig. 3.14).

Figure 3.14: PCR products of HPPR and actin from I. rugosus leaves, roots and stems, M= Molecular weight marker

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Chapter 4

Discussion

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The management of insect pests has always been and will continue to be a constant challenge to agricultural researchers and producers alike. As insect resistance to commonly used pesticides builds and the elimination of more toxic pesticides from the market continues, controlling insect pest infestations will become increasingly more difficult. Therefore, as farmers struggle to remain profitable in a highly competitive global economy, they will be constantly faced with the dilemma of producing high quality, pest-free crops within economical means, and without endangering the environment and the worker’s safety. This struggle has resulted to increase research into alternative control methods, which are cost-effective, environmentally-friendly and capable of keeping insect pests at bay.

4.1. Pesticidal potential of selected plants against target pests The selected botanicals in this study were C. camphora, E. sideroxylon, I. rugosus, B. albiflora, C. procera, D. mucronata and T. minuta. All the selected plants are indigenous, easily available and have been reported as having insecticidal effects against different insect pests (Negi et al., 2012; Pathak et al., 2014; Otengo et al., 2015; Jiang et al., 2016; Alzogaray et al., 2011) except I. rugosus and D. mucronata. The folk or ethnobotanical (phytoinsecticides) uses of I. rugosus and D. mucronata in the area as insect repellents is common and has been so for decades. This led us to hypothesize that these two plant species may contain compounds that could be exploited as insecticides against a number of insect pests of economic importance. Therefore, the insecticidal effects of I. rugosus and D. mucronata are reported in this study for the first time. Exploring the scientific bases of this important trait and transforming local knowledge into commercial uses was the major and long term goal of this study. Although worldwide several botanical pesticides have been reported up to date but there is still need to explore more potent, effective and eco-friendly plant products that have the ability to replace obnoxious synthetic pesticides.

4.1.1. Bactrocera zonata Saunders, 1842 The plant crude methanolic extracts at 2% concentration were assayed for their effect as toxicants against B. zonata males and females, their repellence effect and oviposition deterrence effect on females. Extracts of T. minuta were found to give the

119 highest male mortality and oviposition deterrence, whereas B. albiflora extract showed the strongest repellence. Female mortality with these plant extracts was less than for males, which may have been due to differences in the mode of application for males and females. The highest mortality of males with T. minuta extract was about 73%, but against females, the highest mortality was only 16% with I. rugosus extract, which was not significantly different from the results with the other plant extracts. The lower mortality in females compared to males may have been due to the plant extracts being mixed into the diet and females may have consumed less toxicants by restricting their intake, whereas the males may have been unable to restrict their intake of plant extracts mixed with ME (Haq et al., 2014).

Currently farmers use synthetic insecticides to control fruit flies. In a summer crop of guava 5 to 7 insecticide sprays are applied, in mango 2 to 3 sprays, in plums, peaches, apricot and pear sprays are applied every 10 to 15 days. Ten percent of total synthetic insecticides applied in Pakistan are for fruit flies (Siddiqi et al., 2006; Stonehouse et al., 1998). Due to unacceptable levels of insecticide residues in fruit and vegetables, exports of these are adversely affected.

As plants contain a rich source of bioactive compounds, they may give an alternative solution to synthetic insecticides for control plant pests and diseases (Pino et al., 2013; Ghosh et al., 2012). According to different reports, plant extracts showed strong pesticidal properties and have additional advantages as these chemicals can be specific for targeted pests, biodegradable to nontoxic products and therefore, considered as appropriate to apply in integrated pest management programs (Tare et al., 2004).

In this study we assayed the insecticidal effect of different plants by applying extracts in ME and protein baits that ensured the ingestion of these extracts. Among the plant extracts assayed, T. minuta extracts had the highest toxicity for males. The studies of Shivendra and Singh (1998), Shakunthala and Thomas, (2001a) and Tewari (2001) revealed the insecticidal properties of A. calamus and A. indica against fruit flies. Assessing the efficacy of neem extracts by applying them along with food, Van Randen and Roitberg (1998) reported an inverse effect on adult survival and on the development of eggs of Western fruit fly. Van Randen and Roitberg (1998) also reported that the

120 artificial diet containing neem based insecticides has negative effect on pupae formation and adult emergence of Western fruit fly. Shakunthala and Thomas (2001b) indicated the significant changes in the appearance of reproductive organs of adult B. cucurbitae when the flies were fed with a diet containing A. calamus extract. However, the plant extracts assayed in this study did not cause high female mortality. In addition to plant extracts causing mortality of adult fruit flies, they can repel fruit fly females and deter their oviposition. These effects encourage their incorporation in integrated pest management strategies against fruit flies. This study recorded the highest repellence with B. albiflora and D. mucronata extracts and was in accordance with Walter (1999) and Jimenez et al. (2000), who reported repellence of a number of botanical pesticides against B. zonata on guava. Similarly, Singh et al. (2007) reported the repellent effect of neem products as biopesticide against B. zonata and B. dorsalis. Later studies by Rehman et al. (2009) indicated the effectiveness of petroleum ether, ethanol and acetone extracts of A. calamus, Citrullus colocynthis (L.) Schrad, C. longa, P. harmala, S. lappa, V. jatamansi for repellence and oviposition deterrence of B. zonata and reported promising effects of these plant based pesticides. Solangi et al. (2011) reported that botanical pesticides, such as neem oil, neem seed powder solution, tobacco leaf solution and solution prepared from Eucalyptus leaves, have repellent effects on B. zonata.

Khattak et al. (2006) demonstrated the repellence and growth inhibition caused by petroleum ether, acetone and ethanol extracts of P. harmala, S. lappa and Valariana officinalis L. of B. zonata. These results are in concurrence with the studies of Akhtar et al. (2004), reporting that three plants, sweet flag, neem seed and turmeric rhizomes had repellent effects on B. zonata and that turmeric extract had pronounced effect on suppression of egg laying and emergence of pupae and adults. Siddiqi et al. (2006) indicated the pesticidal effect of acetone, petroleum ether and ethanol extracts of turmeric on B. zonata settling response and fecundity, and reported promising results. Studies on foraging and oviposition behavior of different Bactrocera spp. found that these species have a non-resource based mating system (Kuba and Koyama, 1985; Iwahashi and Majima, 1986) and adult flies engaged in mating during dusk time at the surrounding vegetation of the main host fruits. This behavior of flies suggests that such control strategies should be useful as an area-wide integrated pest management (AW-IPM)

121 approach. The systemic insecticides are not the preferred choice for fruit flies control in guava fruit and, the insecticides having contact action remained insufficient to give successful control of fruit flies, unless targeting the fruit fly adults in abandoned areas and vegetation. Therefore, plant extract formulations giving oviposition deterrence effects have an added advantage over synthetic insecticides and can be included in integrated pest management programs for the control of fruit flies (Khattak et al., 2006).

4.1.2. Schizaphis graminum Rondani, 1852 The crude methanolic extracts of all the selected plants were analyzed at 2% to determine their toxic and repellent effects against S. graminum. In case of toxic effect evaluation, time course mortality was observed after 12, 24, 36 and 48 h. Continuous efforts are being made by the wheat breeders to evolve relatively resistant varieties to aphids and these improved varieties are being released with regular intervals. However, in case of relatively higher infestation of aphids synthetic insecticides are also used. But there are limitations of applying synthetic insecticides on wheat, i.e. residues in the wheat grain can cause severe health risks, damage to the biocontrol agents. In Pakistan, aphid infestation appeared when wheat crop is approaching to maturity stage and due to harvesting of Brassica crop few days earlier the biocontrol agents are shifting from Brassica and Fodder crops to wheat crop. Application of broadspectrum insecticides, therefore, caused severe damage to the non-target beneficial insects (Van Emden and Harrington, 2007). Another limitation of applying the chemical control is that these aphids have been notorious to developed resistance to many of older insecticides including DDT, pyrethroids and parathion-methyl over the past 20-30 years (Gubran et al., 1992; Gao et al., 1992); this is most likely that aphids can acquire resistance against modern pesticides. As plants constitute a rich source of bioactive chemical compounds, botanical insecticides may present attractive alternatives to currently used synthetic chemical insecticides for pest management (Pino et al. 2013; Miresmailli and Isman, 2014; Pavela, 2016; Khan et al., 2016). As compared to synthetic pesticides, botanical pesticides besides their insecticidal potential have been reported to pose little threat to the environment or to human health (Pavela 2015, 2016).

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To determine toxic effects, no choice method was used, in which aphids were exposed to plant extracts treated leaves. Time course mortality was observed after 12, 24, 36 and 48 h of treatment. Among all the analyzed plant extracts, Calotropis procera at 2% concentration depicted significantly higher mortality after every 12 h of treatment. After 48 h, mortality caused by C. procera at 2% concentration, was 96% which was not significantly different from mortality caused by synthetic pesticide control, Karate (lambda cyhalothrin). Previously, when alkaloids from Macleaya microcarpa were analyzed for their insecticidal activity against S. graminum, the LC50 value after 48 h was 0.710 g/L (Feng et al., 2008). There are numerous studies, reporting the insecticidal potential of C. procera against many insects (Bakavathiappan et al., 2012; Khan et al., 2012; Pathak et al., 2014). Begum et al. (2011) evaluated the insecticidal activity of ethanolic extracts of leaves from C. procera against Musca domestica in different concentrations and found that the C. procera showed LC50 value of 282.5 mg/L. In this study, C. procera exhibited higher mortality followed by 54% mortality caused by D. mucronata, 16% depicted by I. rugosus and B. albifora. Other treatments including T. minuta, E. sideroxylon and C. camphora depicted no or negligible toxic effects. These findings showed similar results to the study conducted by Sohail et al. (2012), who observed that tobacco extract gave maximum effect followed by neem and garlic extract when analyzed against aphids. Similarly, insecticidal effect of hexane extract and its isolated compound, xanthoxylin from plant, Ungernia severtzovii bulb was reported against S. graminum (Chermenskaya et al., 2012). Gao et al.(2001) reported the toxicity of 1,5-diphenyl-1-pentanone and 1,5-diphenyl-2-penten-1-one isolated from Stellera chamaejasme, against S. graminum and found that 1,5-diphenyl-1-pentanone gave LC50 of 0.4 g/L, while 1,5-diphenyl-2-penten-1-one gave LC50 of 0.2 g/L.

To evaluate repellent effect, choice method was used in which aphids were exposed to plant extract treated and untreated leaves simultaneously in each treatment. Repellent effect was observed after every 12, 24, 36 and 48 h. All the treatments showed considerable repellent effect. After every 12 h, synthetic pesticide, Karate depicted negative values which mean that they have no repellence. Karate is known to have contact toxicity, and due to attraction aphids died instead of repelling from Karate treated leaf throughout the bioassay until 72 h. Maximum repellence of 76% was observed in

123 case of D. mucronata after 12 h which was reduced after every 12 h and after 72 h it was 32% because D. mucronata exhibited highly toxic effects insteated of repellent effects, therefore mortality observed in insects exposed to this treatment. Insect mortality might be due to entering of toxic odour into insect’s spiracles that blocked oxygen supply (Mekuaninte et al., 2011). Daphne mucronata has high ethnopharmacological value and has been used in conventional medication to treat many diseases (Afzal et al., 2009; Mosaddegh et al., 2012; Hussain et al., 2012; Ghasemi et al., 2013). Similar effects were showen by C. procera that exhibited 56% repellence after 12 h whereas; effect was reduced to 12% after 72 h. In the whole bioassay, repellent effect depicted by B. albifora and E. sideroxylon was inceased after every 12 h. These plants might contain effective insecticidal compounds that have the ability to repell the insects and thus insects were unable to feed on leaf treated with these extracts. The repellenct effect of isolated compounds, luciamin from Solanum laxum and indole alkaloid graminefrom barley were reported against S. graminum which exhibited effective repellent effects (Soulé et al., 2000; Ryan, 2002). Similarly, choice method was conducted by Chermenskaya et al. 2010, who reported the deterrence effect of 20 plant extracts against S. graminum and found that Ungernia severtzovii have maximum deterrence index of 54.2%.

4.1.3. Tribollium castaneum Herbst, 1797 In this study all the selected plant crude methanolic extracts were analyzed at 2% concentration against aduts of T. castaneum and mortality was observed after 24, 48 and 72 h. Two methods were used to analyze the effect of crude methanolic extracts of plants against adults of Tribolium castaneum. Due to continuous use of chemical insecticides, T. castaneum become resistant worldwide (Dyte and Blackman, 1970; Champ and Dyte, 1976; Rossi et al., 2010; Singh and Prakash, 2013). A number of plant based pesticides as alternative to synthetic pesticides, known to posses effective insecticidal properties that can control insect pests during storage (Joel, 2015).

In this study, one of the methods used was impregnation bioassay, in which plant extracts treated filter papers after drying were exposed to adults of T. castaneum. In the second method, beetles were fed on flour discs containing plant extracts. In both methods mortality was assessed after 24, 48 and 72 h. There was no mortality observed

124 in case of any plant extract even after 72 h in both methods. The reason may be the concentration used in this study was not enough high to cause mortality in the beetles or these plant extracts might not contain such compounds that could be effective against these beetles. In another study, no toxic effects was observed when the aqueous and methanolic extracts of T. minuta were evaluated for their toxic and repellence effects against T. castaneum through topical application and application of extracts on grains, it was found that T. minuta showed only strong repellent effect against T. castaneum (Padin et al., 2013). Powders from the aerial parts of the C. camphora were analyzed against T. castaneum and Trogoderma granarium. Trogoderma granarium was more susceptible to C. camphora plant powders than T. castaneum (Nenaah and Ibrahim, 2011). In this study different methodology was used to analyze the toxic effect of selected plant extracts.

Previously, C. procera showed pronounced insecticidal activity against T. castaneum (Khan et al., 2012) because they used 80% of the extract and observation was taken after 3 months as compared to the this study where low concentration was used by utilising different mode of application of selected plant extracts for up to 72 h of treatment. In this study, C. procera showed no effect, the reason is lower concentration (2%) of extract used and observation was taken until 72 h. Mostafa et al. (2012) also reported that when T. castaneum were exposed to Eucalyptus species extract, there was no mortality observed after 72 h. Khan et al. (2013) used the same impregnation methodology for 24, 48 and 72 h, reported that E. globules extract was used at 5, 10 and 15% and gave 14.5% mortality at 15%, which means that even at higher concentration mortality was low.

4.1.4. Drosophila melanogaster Meigen, 1830 All the selected crude methanolic plant extracts at 2% concentration were analyzed for their toxic effects against adults of Drosophila melanogaster for 24, 48 and 72 h. Plant extracts were layered on the diet and flies were exposed to treated diet after drying of extracts on the diet surface. Among all the plant extract analyzed, C. camphora gave maximum mortality of 41.3% after 72 h. Khan et al. (2015) demonstrated the toxic effect of Otostegia limbata extract against D. melanogaster for 24 h and reported that at 2% concentration of the extract, the mortality was 12%. They also observed that with the

125 increase of concentration mortality also increased, thus at 6% concentration, mortality reached to 90%. In this study, after higher mortality of 41.3% caused by C. camphora, C. procera gave 7% mortality followed by T. minuta that exhibited 3.3% mortality after 72 h of treatment. Begum et al. (2011) evaluated the insecticidal activity of ethanolic extracts of leaves from C. procera against Musca domestica in different concentrations and found that the C. procera gave LC50 value of 282.5 mg/L. Barati et al. (2013) reported the insecticidal activity of C. procera against Bemisia tabaci grown on tomato plants under controlled conditions in glass house. The aerial parts of T. minuta were analyzed for insecticidal activity against sand fly, Phlebotomus duboscgi and considerable mortalities were observed (Njeru, 2011).

Miyazawa et al. (2004) reported the insecticidal effect of Angelica acutiloba extract and its isolated compound against D. melanogaster. Bioactivity guided isolation of choloroform extract, resulted in four compounds (Z)-ligustilide, furanocoumarins, xanthotoxin, (Z)-butylidenephthalide and isopimpinellin. (Z)-butylidenephthalide gave more toxic effect against adults with LD50 of 3.7 µg/adult.

Tsukamoto et al. (2005) demonstrated the insecticidal activity of Cnidium officinale and its isolated compounds against adults of D. melanogaster. Four compounds isolated through bioassay guided isolation procedure, (Z)-ligustilide, neocnidilide, cnidilide and (3S)-butylphthalide. The compound, (3S)-butylphthalide depicted more toxicity with LD50 3.7 µg/adult as compared to other analyzed compounds.

Anjum et al. (2010) evaluated the toxic effect of Neem extract against D. melanogaster larvae. It was found that Neem extract gave high lethality with LD50 of 2.4%. In this study I. rugosus, E. sideroxylon, B. albifora and D. mucronata depicted no mortality even after 72 h. The reason may be the insecticidal compounds that could be effective against D. melanogaster might not be present in these analyzed plant extracts.

4.1.5. Spodoptera exigua Hubner, 1808 Crude methanolic extracts at 2% concentration were analyzed for their toxic effects against seccond instar larvae of Spodoptera exigua and time course mortality was observed after 24, 48 and 72 h of treatment. The second instar larvae were selected for

126 the study because at this stage insects are able to move from plant to plant. Among all the treatments, C. procera extract exhibited maximum of 22% mortality after 48 and 72 h of treatment. Upadhyay, (2013) reported the insecticidal effect of C. procera extract against third instar larvae of Spodoptera litura and obseved that after 24 h exposure to methanolic extract, being topically applied on insect body, the LD50 was 3 μg/gm body weights of insect larvae. In another study, chloroform, hexane, ethanol, ethyl acetate and methanol extracts from the leaves of C. procera were evaluated for their antifeedent activity against S. litura and found that choloroform extract had more antifeedent effect as compared to all other analyzed extracts (Bakavathiappan et al., 2012). From leaves and latex of C. procera, important pharmaceutical compounds have been isolated like calotropon, uscharine, calotropin, calctin, calotropagenin and uscharidin (Chopra et al., 1956; Pathak et al., 2014); these may be responsible for such insecticidal activity.

In this study, Tagetes minuta, C. camphora, E. sideroxylon, I. rugosus and D. mucronata depicted 6% mortality after 72 h, while B. albifora exhibited no mortality. Similar weak toxicity was observed by Feng et al. (2012), who analyzed the 30 plant species against S. exigua for 24, 48 and 72 h and found that after 72 h, Fritillatia thunbergii extract gave maximum of only 33.3% mortality and there was no significant difference between all the treated plant extract.

Bullangpoti et al. (2011) reported the toxic effects of ethyl acetate extract and isolated compound ricinine from senescent leaves of Jatropha gossypifolia against second instar larvae of S. exigua and observed LC50 of 8.644 ppm and 3.215 ppm in case of ethyl acetate extract and ricinine, respectively. In another study, Ntalli et al. (2014) analyzed extract of Melia azedarach fruits and its fraction of limonoids against S. exigua and observed that there was low toxic effect but insect activity was reduced considerably.

4.1.6. Acyrthosiphon pisum Harris, 1776 In this study, all the selected crude methanolic plant extracts were analyzed in different concentrations for their toxic effects against A. pisum and mortality was observed after 24 h of the treatment. Among all of the analyzed crude extracts, the most active plants were I. rugosus and D. mucronata, with the lowest LC values as compared

127 to the other plants evaluated. On the basis of the lowest LC values, I. rugosus and D. mucronata crude methanolic extracts were chosen for fractionation into different solvent fractions. The highest bioactivity against aphids for the fractions obtained from D. mucronata was observed with the ethyl acetate and dichloromethane fractions, which are moderately polar solvents. This observation is similar to the findings of Zewdu, (2010) where toxic effects of different solvent extracts of Birbira, Millettia ferruginea, were evaluated against A. pisum. It was reported that deionized water extract was more toxic causing 98% mortality, followed by acetic acid extract causing 89% mortality, whereas, the chloroform, toluene and hexane extracts were the least toxic. As compared to D. mucronata fractions, I. rugosus fractions gave the highest mortality even at lower concentrations. The highest bioactivity for the fractions obtained from I. rugosus was observed with the butanol and ethyl acetate fractions, with the smallest LC90 values obtained for the butanol fraction. It can be said that the butanol fraction is more active at lower concentration as compared to ethyl acetate. In a similar study, Baek et al. (2013) found that the butanol fraction had more activity as compared to other tested fractions when dried seeds of Macleaya cordata extracted with methanol and successively partitioned into hexane, butanol and water fractions, were analyzed against the cotton aphid, Aphis gossypii. The presence of alkaloids, saponins, tannins, flavonoids, phenols, terpenes, glycosides and steroids have been reported in the butanol fraction for some plant species such as Sapindus saponaria (Gangula et al., 2013). Since some of these compounds are known to have insecticidal properties, this could explain the high toxicity against aphids observed with the butanol fraction. Baek et al. (2013) reported the presence of alkaloids, 8-hydroxydihydrochelerythrine and 8-methoxydihydrosanguinarine in the butanol fraction of Macleaya cordata seeds, which were very active against Aphis gossypii.

Since for various pharmacological activities I. rugosus and D. mucronata are used as a source of natural compounds, the extracts obtained are expected to be of low or no hazard to human beings or other animals. The butanol fraction from I. rugosus gave lowest LC90 value of 48.2 ppm. The genus Daphne has been known for the production of valuable bioactive natural products including flavonoids, coumarins, lignan, coumarinolignans and triterpenoids (Rasool et al., 2009). D. mucronata is considered as

128 an important medicinal plant in northern areas of Pakistan as well as in several regions of Iran. It has high ethnopharmacological value and has been used in conventional medication to treat many diseases (Afzal et al., 2009; Mosaddegh et al., 2012; Hussain et al., 2012; Ghasemi et al., 2013). I. rugosus is known to contain some important bioactive compounds including flavonoids, terpenoids, saponins, tannins, cardiac glycosides, coumarins and steroids (Janbaz et al., 2014). It is used in traditional medicine in Pakistan to treat many diseases (Ajmal et al., 2012; Sher et al., 2011; Sabeen and Ahmad, 2009; Akhtar et al., 2013). Extracts and different solvent fractions of this plant are known to exhibit antifungal (Rauf et al., 2012a), antibacterial, phytotoxic (Rauf et al., 2012b) and antioxidant effects (Rauf et al., 2013). As both of these plants are already known to have considerable medical importance, it is of interest that these plants were found to be the best sources of potential pesticides against A. Pisum aphids in our study.

4.2. Isolation of active compound (rosmarinic acid) from butanol fraction of Isodon rugosus On the basis of all the bioassays of seven crude methanolic plant extracts against all the target insects, I. rugosus and D. mucronata extracts were selected with the best bioactivity against A. pisum. Different solvent fractions of these both plants were analyzed in different concentration against A. pisum. Then a butanol fraction of I. rugosus was further selected on the basis of maximum bioactivity against A. pisum. The butanol fraction of I. rugosus was further fractionated to isolate the active principle against A. pisum through bioactivity guided isolation procedure. Similarly, in another study, through bioactivity guided isolation, the active principle, ailanthone was isolated from the aqueous fraction of Ailanthus altissima against A. pisum (De Feo et al., 2009). Previously, the butanol fraction from Citrullus colocynthis was more active against Aphis craccivora, and bioactivity guided isolation led to the isolation of the active principle, 2- O-ß-D-glucopyranosylcucurbitacin E (Torkey et al., 2009).

In this study, the butanol fraction was sub-fractionated through reverse phase automatic flash chromatography. After bioactivity testing of all the resulting sub- fractions (1A-14A) against A. pisum, fraction 3A with lower LC values was selected for further fractionation. Through prep-LC, fraction 3A was sub-fractionated and resulting

129 sub-fractions (3A-1, 3A-2 and 3A-3) were analyzed for their bioactivity. Fraction 3A-3 with lower LC values was subjected to spectroscopic analysis. 1H NMR spectroscopy confirmed that the isolated fraction was rosmarinic acid but due to inconsistent chromatographic behavior during prep-LC, not enough compound was collected to record 13C NMR. The inconsistent chromatographic behavior with peak splitting can have several causes, contamination on guard or column inlet, part, blocked frit or a small void at the column inlet (~wear). The problem of peak shifting (variable retention times) can be due to small changes in mobile composition, temperature fluctuations, column overloading and a combination of these two problems can lead to different UV patterns for each run. Due to this problem, the reverse phase automatic flash chromatography was repeated with a higher amount of butanol fraction. Out of all resulting sub-fractions (1B- 6B), 1B was selected with lower LC values on the basis of bioactivity against A. pisum. Fraction 1B was subjected to acidic extraction to get two phases, water and ethyl acetate. The ethyl acetate phase fraction was more active with lower LC values. After removing ehyl acetate azeotropically with toluene and drying, active principle was purified and identified through different spectroscopic techniques as rosmarinic acid. Similarly in another study, Chakraborty et al. (2007) reported the isolation of caffeic acid and rosmarinic acid from Basilicum polystachyon through acidic extraction with HCl followed by treatment with ethyl acetate and analyzed their antimicrobial activities.

4.3. Active principle, rosmarinic acid This study for the first time reports the isolation and purification of rosmarinic acid (RA) from I. rugosus and its bioactivity against A. pisum. Rosmarinic acid is a phenolic acid soluble in water. RA is a caffeic ester derived from caffeic acid, 3,4- dihydroxycinnamic acid, and 3,4-dihydroxyphenyllactic acid and reported to be present in 39 plant families including Boraginaceae, Choranthaceae, some monocotyledonous plant orders and eudicotyledoneae but it is reported to be mainly present in Lamiaceae plant family (Sahraroo et al., 2014; Khojasteh et al., 2016; Peng et al., 2016). For the very first time, RA was isolated from the plant, Rosmarinus officinalis (Ellis and Towers, 1970). Its structure was first identified by Scarpati and Oriente (1958).

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In this study, rosmarinic acid isolated from I. rugosus (IR-RA) and commercial rosmarinic acid were evaluated in different concentrations for their toxicity against A. pisum for 24 h. There was no significant difference observed between the bioactivity depicted by both rosmarinic acids. IR-RA gave LC values of LC50 0.2 ppm and LC90 5.4 ppm. These are very low LC values depicted after 24 h of bioassay, such lower LC values have never been reported previously in any study against A. pisum using the same feeding bioassay methodology (Sadeghi et al., 2009; Carrillo et al., 2011; Smagghe et al., 2010; De Geyter et al., 2012; Nachman et al., 2012; Giner et al., 2013; Zapata et al., 2016). This means that a very low amount of rosmarinic acid can cause significant toxic effects against A. pisum in 24 h. RA have been known to possess various important biological activities including anti-inflammatory, antimutagenic, antiviral, antibacterial, antioxidant activity, and is used to treat various illnesses such as prevention of Alzheimer’s disease development, improve cognitive performance, reduce kidney disease severity, prevent cancer, rheumatoid arthritis, peptic ulcer, cataracts, bronchial asthma and arthritis (Petersen and Simmonds, 2003; Bulgakov et al., 2012; Sahraroo et al., 2014; Khojasteh et al., 2016). Infact, the awareness to use RA as pharmaceutical and as a dietary supplement to improve human health has been increasing (Bhatt et al., 2013; Khojasteh et al., 2014). Recently, pure RA is used commercially as a drug although many products containing RA are available in the market. It has also been used as preservative in food (Sahraroo et al., 2014).

In another study, it was confirmed that rosmarinic acid could reduced genotoxic effects induced by other chemicals. For this purpose, micronucleus and comet assay were performed by using V79 cells. Cultures were treated with different concentrations of rosmarinic acid alone or along with the DNA damaging agent, doxorubicin. It was confirmed that no genotoxic effects were induced by RA but it could significantly reduced the DNA damage and micronuclei induced by doxorubicin. It was suggested that the antioxidant property of RA could be responsible for this reduction of damage (Furtado et al., 2010).

Very few insecticidal activities have been reported for rosmarinic acid. Regnault- Roger et al. (2004) investigated the insecticidal activities of polyphenolic compounds,

131 isolated from five plants belonging to Lamiaceae family against Acanthoscelides obtectus (Say) and observed that among all the polyphenolic compounds, rosmarinic acid and luteolin-7-glucoside were more toxic. In another study rosmarinic acid isolated from the ethyl acetate fraction of Zostera marina was analyzed for its toxic effects against the nematode, Bursaphelenchusxy lophilus and after 24, 48 and 72 h the LD50 values were 1.2, 1.1 and 1 mg/mL, respectively (Wang et al., 2012).

In this study, comparison between growth of live aphids exposed to plant extract treated and untreated diet after 24 h of bioassay was analyzed. It was clearly observed that the growth of A. pisum nymphs was stopped after 48 h of exposure to rosmarinic acid treated diet and thus the size of the aphids was reduced and ultimately they died as compared to aphids exposed to untreated diet. Similar observation was made by Sadeghi et al. (2009), who observed that aphid size was reduced after 48 h of exposure to novel biorational insecticides, flonicamid and pymetrozine and mortality was observed after 72 h.

4.4. Identification of genes involved in the synthesis of rosmarinic acid in Isodon rugosus For the utilization and industrialization of rosmarinic acid as a botanical pesticide product, the research on biosynthetic pathway of rosmarinic acid might contribute significantly. Incomplete and poor knowledge towards the biosynthetic pathway leads to limited biosynthetic pathway engineering. Biotechnological rosmarinic acid production has been achieved through cell cultures of C. blumei (Razzaque and Ellis, 1977). Suspension cultures of S. officinalis depicted that 36% of RA contents can be achieved (Kim et al., 2015). Molecular knowledge of the biosynthesis of the RA can help to maximize its production in I. rugosus through gene overexpresson studies. This information can also be utilized for RA production in microbes by engineering RA biosynthesis pathway, as described by Bulgakov et al. (2012), Jiang et al. (2016) and Ru et al. (2016). Therefore, research on rosmarinic acid biosynthetic pathway in Isodon rugosus will be helpful for industrial exploitation of rosmarinic acid.

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In the biosynthesis of rosmarinic acid, eight genes have been involved as described in Coleus blumei by many researchers (Petersen et al., 1993, 1994; Kim et al., 2004; Eberle et al., 2009). In this study two key genes were targeted to amplify in I. rugosus. The targeted genes are working specifically in biosynthesis of RA. Other genes of the pathway are involved in many other alternate pathways as well. One of the targeted genes was rosmarinic acid synthase (RAS) and other was hydrooxyphenylpyruvate reductase (HPPR). Rosmarinic acid biosynthetic pathway in C. blumei, has been thoroughly explained by Petersen et al. (1993), (2009) and Petersen (1997). During last years, the enzymes involved in this biosynthetic pathway have been investigated in cell cultures of many plant species belonging to Lamiaceae and Boraginaceae families (Weitzel and Petersen, 2011). The genes involved in biosynthesis of rosmarinic acid has been cloned from many species including Melissa officinalis (Weitzel and Petersen, 2010; 2011), Salvia miltiorrhiza (Huang et al., 2008; Song and Wang, 2009; Di et al., 2013), Prunella vulgaris (Kim et al., 2014; Ru et al., 2016) and Lithospermum erythrorhizon (Yamamura et al., 2001) and Coleus blumei (Kim et al., 2004; Berger et al., 2006). The precursors of the pathway are two amino acids, L-phenylalanine and L- tyrosine.

For HPPR cloning from I. rugosus, degenerate primers were designed by chosing highly conserved region by aligning sequences for HPPR from closely related plant species, Perilla frutescens, Solenostemon scutellarioides, Salvia officinalis and Salvia miltiorrhiza and with the help of these primers conserved region for HPPR from I. rugosus was amplified and sequence obtained was 388 bp in length. Then RACE PCR was performed to get full length sequence for HPPR from I. rugosus, by using gene specific primers based on the already amplified conserved region of HPPR in I. rugosus. RACE PCR results in amplification of 3′ RACE product with sequence length of 251 bp and 5′ RACE amplified product with sequence length of 502 bp. This is the first study reporting the cloning of HPPR gene from I. rugosus. HPPR, firstly defined by Grant (1989) belongs to the family of D-isomer specific 2-hydroxyacid dehydrogenase. HPPR responsible for the biosynthesis of rosmarinic acid and is considered first key enzyme in the biosynthetic pathway (Kim et al., 2004; Janiak et al., 2010). From suspension cultured cells of C. blumei in cell free extracts HPPR was characterized for the first time

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(Petersen and Alfermann, 1988; Hausler et al., 1991), afterwards purified and then sequenced (Kim et al., 2004). Its structure was solved by Janiak et al, 2010. It is active in the form of homodimer. In the biosynthetic pathway of rosmarinic acid it is involved in the reduction of 4-hydroxy phenylpyruvate with the use of NAD(P)H, which is obtained through transamination of L-tyrosine, to D-4-hydroxy phenyllactate (Petersen et al., 1993; Petersen, 1997). As indicated in the dendrogram in Fig. 3.13, the phylogenetic observation showed high levels of similarity of HPPR genes among the different plant species belongs to Lamiaceae family. The phylogenetic tree complements the sequence alignment results for the HPPR gene. It shows the similarity of the gene to its closely related counterparts especially to P. frutescens.

For cloning of RAS gene from I. rugosus, degenerate primers were designed by chosing highly conserved region by aligning sequences for RAS from closely related plant species, Melissa officinalis, Perilla frutescens, Salvia miltiorrhiza and Solenostemon scutellarioides. Three sets of primers were designed but no amplification of RAS from I. rugosus was observed or sometimes multiple bands appeared. It is possible that the RAS gene in I. rugosus does not share high similarities to the RAS gene in the other related plant species. This will make it difficult to amplify a target region in the RAS gene using degenerate primers designed from the RAS gene of closely related plant species. However we can not make strong conclusions based on this. Further tests will be required to fully investigate if the RAS gene is present, absent or different. Nevertheless, the HPPR gene also plays a key role in RA biosynthesis; hence identifying the presence of this gene (HPPR) equally proves that the RA biosynthetic pathway is present in I. rugosus. In biosynthetic pathway, RAS transestrified D-4- hydroxyphenyllacted with 4-coumaroyl moiety obtained from 4-coumaroyl-CoA, to 4- coumaroyl-40-hydroxy phenyllactate and then by cytochrome P450 monooxygenase reactions hydroxylated to rosmarinic acid (Petersen, 1997; Petersen and Simmonds, 2003). It acts in the form of monomer, belongs to the family of BAHD acyltransferase (Berger et al., 2006).

HPPR and RAS were reported to be identified and described their involvement in biosynthesis of rosmarinic acid in Coleus blumei Benth (Petersen and Alfermann 1988;

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Hausler et al., 1991; Petersen, 1991; 1997; Petersen et al., 1993, 2009), Melissa officinalis (Weitzel and Petersen, 2011) and Prunella vulgaris (Kim et al., 2014; Ru et al., 2016).

In order to analyze the involvement of HPPR and RAS in the biosynthesis of rosmarinic acid, Hucherig and Petersen (2012) conducted a study in which they confirmed that both of these enzymes participate in biosynthesis of RA. For this purpose, they established the hairy root lines of C. blumei having constructs of HPPR or RAS RNAi overexpression and suppression. Hairy roots lines showed low RA content up to 92% reduction as compared to control when levels of HPPR or RAS mRNA were lowered. When HPPR levels were increased, RA content increased up to 176% in hairy root lines with overexpression, while overexpression of RAS resulted in reduced level of RA due to co-suppression effects.

In this study it was confirmed that HPPR expressed in all parts of I. rugosus, including roots, stems and leaves. As, HPPR involved in the biosynthesis of rosmarinic acid is confirmed by many previous studies as stated above, this plant can be utilize as a potential source for rosmarnic acid. This is a premier study reporting insecticidal potential of I. rugosus, identifying rosmarnic acid as potent aphicide and expression and sequence analysis of hydroxylphenylpyruvate reductase gene responsible for synthesis of rosmarinic acid in I. rugosus.

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4.5. Conclusion The aim of this study was to evaluate insecticidal potential of selected plants and to isolate and characterize plant based pesticidal compounds along with their biocemical and molecular chracaterization from lower northern areas of Pakistan. For this purpose, seven plant species namely; Cinnamomum camphora, Eucalyptus sideroxylon, Isodon rugosus, Boenninghausenia albiflora, Calotropis procera, Daphne mucronata, Tagetes minutawere selected. Insecticidal activities of selected plants were determined against Bactrocera zonata, Schizaphis graminum, Drosophila melanogaster, Tribolium castaneum, Spodoptera exigua and Acyrthosiphon pisum. In conclusion, Isodon rugosus and Daphne mucronata were identified as the most potent plants against Acyrthosiphon pisum. Following bioactivity guided selection; rosmarinic acid was isolated and identified through spectroscopic analysis as the bioactive compound in I. rugosus. Molecular analysis further confirmed the presence of the biosynthetic pathway of rosmarinic acid in different plant tissues. Based on the bioassay results, either the extracts from I. rugosus or the isolated insecticidal compound, rosmarinic acid, could be exploited to develop potent aphicides, because of the high mortality caused at very low concentrations to aphids. This potential botanical insecticide may fit well in IPM programs designed to control aphids. Considering that I. rugosus is already used for medicinal purposes, it will be safer compared to the current conventional pesticides used to control aphids. Also, rosmarinic acid is known to reduce genotoxic effects induced by chemicals, which is contrary to the currently used toxic synthetic pesticides that are responsible for genotoxic effect induction in consumers. While this study highlights the potential of I. rugosus as a possible biopesticide against a notorious insect pest such as A. pisum, it also provides the basis for further exploration and development of a formulation for effective field application and because of solubility of rosmarinic acid in water; farmers can easily use water extract of the plant I. rugosus against insects.

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4.6. Recommendations • Daphne mucronata exhibited considerable aphicidal activities, can be further explored to isolate active compounds. • In depth molecular characterization of genes involved in rosmarinic acid biosynthesis is required to enhance its quantity within I. rugosus for commercial rosmarinic acid production and to engineer in other suitable plants. • Purified rosmarinic acid may also be analyzed for its toxic effects against other ecomically important insect pests. • Analysis of rosmarinic acid effects towards non-target organisms is required to investigate.

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Chapter 5

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