Engineering Insect-Resistant by Transgenic Expression of an Insecticidal - Peptide

Md. Shohidul Alam Master of Science in Agricultural Chemistry

A thesis submitted for the degree of Doctor of Philosophy at The University of Queensland in 2014 Institute for Molecular Bioscience

Abstract The insecticidal spider-venom peptide ω-hexatoxin-Hv1a (Hv1a) from the Australian Blue Mountains funnel-web spider versuta is one of the most potent insect-specific isolated to date. Hv1a blocks voltage-gated calcium channels in the insect central nervous system, a mechanism quite distinct from existing chemical . It induces a slow-onset paralysis that precedes death in a taxonomically wide range of insects. Hv1a's broad spectrum of target insects, novel mode of action, and absence of toxicity to makes the Hv1a an attractive tool for generating insect- resistant transgenic crops.

The oral activity of Hv1a can be enhanced by coupling it to the lectin Galanthus nivalis agglutinin (GNA) or with the minor capsid protein of pea enation mosaic virus (CP). Recombinant fusions of Hv1a with GNA were produced using the Pichia pastoris expression system to study the intrinsic insecticidal activity of Hv1a-GNA and GNA-Hv1a fusion proteins. By using injection bioassays with houseflies, we found that the intrinsic insecticidal activity of Hv1a was maintained when it was fused to GNA. Moreover, feeding bioassays with diamondback moth larvae revealed that fusion of Hv1a to GNA, in either orientation, enhances its oral insecticidal activity.

In order to generate transgenic plants expressing Hv1a alone or fused to GNA or CP, transformation vectors were constructed by ligating synthesised in the pAOV binary vector with the constitutive Cauliflower Mosaic Virus 35S promoter (35S) or the phloem tissue-specific SUCROSE TRANSPORTER 2 (SUC2) promoter. Homozygous transgenic Arabidopsis were generated using the floral dip method of Agrobacterium-mediated plant transformation and subsequent herbicide selection. PCR of genomic DNA and western blotting were used to confirm integration of the transgenes and protein expression in the transgenic Arabidopsis respectively. Initial experiments revealed a very high level of mortality of Helicoverpa armigera larvae on wild-type plants due to the presence of endogenous glucosinolates, which masked the insecticidal effects of the transgenes Thus, a new set of transgenic plants was generated using an Arabidopsis cyp79B2 cyp79B myb28 myb29 quadruple mutant that lacks endogenous glucosinolates.

Bioassays revealed that H. armigera larvae had a lower level of survival and retarded growth when fed on leaves of transgenic Arabidopsis expressing Hv1a under 35S promoter control compared with those fed on gluc-null control plants. Moreover, larval

mortality was higher for plants expressing Hv1a/GNA fusions than those expressing Hv1a or GNA alone. The highest larval mortality, lowest larval weight gain, and lowest level of leaf damage were observed for larvae fed on plants expressing GNA-Hv1a. Mortality was extremely high (~90%) for larvae fed on GNA-Hv1a plants for 15 days. The resistance to cotton bollworms conferred by expression of GNA-Hv1a in transgenic Arabidopsis highlights the potential of Hv1a transgenes as an alternative to harmful chemical insecticides. Moreover, Hv1a transgenes might provide a useful adjunct or alternative to Bt crops, and they might be useful for trait stacking with Bt transgenes.

Declaration by author

This thesis is composed of my original work, and contains no material previously published or written by another person except where due reference has been made in the text. I have clearly stated the contribution by others to jointly-authored works that I have included in my thesis.

I have clearly stated the contribution of others to my thesis as a whole, including statistical assistance, survey design, data analysis, significant technical procedures, professional editorial advice, and any other original research work used or reported in my thesis. The content of my thesis is the result of work I have carried out since the commencement of my research higher degree candidature and does not include a substantial part of work that has been submitted to qualify for the award of any other degree or diploma in any university or other tertiary institution. I have clearly stated which parts of my thesis, if any, have been submitted to qualify for another award.

I acknowledge that an electronic copy of my thesis must be lodged with the University Library and, subject to the General Award Rules of The University of Queensland, immediately made available for research and study in accordance with the Copyright Act 1968.

I acknowledge that copyright of all material contained in my thesis resides with the copyright holder(s) of that material. Where appropriate I have obtained copyright permission from the copyright holder to reproduce material in this thesis.

Publications during candidature

Book chapters

Herzig, V.; Bende, N. S.; Alam, M. S.; Tedford, H. W.; Kennedy, R. M. & King, G. F. 2014. Chapter Eight - Methods for Deployment of Spider Venom Peptides as Bioinsecticides. In: Tarlochan, S. D. & Sarjeet, S. G. (eds.) Advances in Insect Physiology. Academic Press.

Conference abstracts

Alam, M.S., Mylne, J. S. and King, G.F. (2013) Spider-venom peptide protects plants from insect pest attack. East Coast Protein Meeting–2013, Coffs Harbour, NSW, Australia.

Publications included in this thesis

No publications have been incorporated into this thesis.

Contributions by others to the thesis

Dr Raveendra Anangi and MSc student Mr Mario Donald Bani contributed in part to the design, standardization, and purification of the recombinant fusion proteins described in Chapter 2. Dr Volker Herzig contributed in injection bioassay of houseflies with recombinant fusion proteins described in Chapter 2.

Statement of parts of the thesis submitted to qualify for the award of another degree

Portions of some of the figures from Chapter 2 were used in an MSc thesis submitted by Mario Donald Bani to The University of Queensland (degree awarded July 2013).

Acknowledgements I would like to extend my heartfelt and sincere gratitude to my principal advisor Professor Glenn F. King (Institute for Molecular Bioscience, The University of Queensland, Australia) for his expert supervision, guidance, support and enthusiastic encouragement throughout the course of my PhD. I also wish to express my gratitude, sincere appreciation and indebtedness to my associate advisor Associate Professor Joshua S. Mylne (The University of Western Australia, Australia) for his valuable guidance and cooperation during the research work.

I would like to thank Professor Myron Zalucki (School of Biological Sciences, UQ), Dr Steven Reid (SCMB and AIBN, UQ) and Dr Mark Jackson (IMB, UQ) for their guidance and encouragement as members of my candidature committee.

I would like to acknowledge those who helped in one way or another during the tenure of my research work and report writing, including Dr Brit Winnen for help with gel electrophoresis and western blotting; Dr Raveendra Anangi for help with recombinant protein expression; Dr Aurelie Chanson for help with recombinant DNA technology and techniques; Dr Lynda Perkins for help with insect bioassays and data analysis; Dr Volker Herzig and Dr Margaret Hardy for help with insect bioassays and data analysis; Dr Elaine Fitches (FERA, York, UK) for providing anti-GNA antibody; and Dr Mark Kinkema (AgBiTech, Queensland, Australia) for providing Helicoverpa armigera eggs.

Thanks to all members of the King Group for creating such a warm, friendly and cooperative atmosphere. Particular thanks to Dr Maria Ikonomopoulou, Dr David Morgenstern, Dr Julie Klint, Dr Sandy Pineda Gonzalez, Dr Eivind Undheim, Dr Naushad Shaikh, Mr Niraj Bende, Mrs Darshani Rupasinghe, Mr Carus Lau, Ms Jessie Er and Mr Sebastian Senff for their help, advice, and guidance.

I would like to acknowledge all staff of the Queensland Bioscience Precinct and the Institute for Molecular Bioscience (IMB), specially Dr Amanda Carozzi for her help with my scholarship applications and candidature, and Mikiko Miyagi for help in the laboratory.

I am also very grateful to the IMB and The University of Queensland (UQ) for funding my RHD candidature via an IMB Postgraduate Award, UQ International Research Tuition Award (UQIRTA), and UQ Research Scholarship (UQRS).

Last, but no means least, I would like to express my heartfelt thanks to my family, who have provided limitless love and support over the years.

Md. Shohidul Alam

Keywords , insecticidal , insect-resistant transgenic crop, spider-venom peptide, Galanthus nivalis agglutinin, capsid protein, Arabidopsis thaliana, glucosinolates, Helicoverpa armigera, Plutella xylostella

Australian and New Zealand Standard Research Classifications (ANZSRC)

ANZSRC Code 100103: Agricultural Molecular Engineering of Nucleic Acids and Proteins, 50% ANZSRC Code 060702: Plant Cell and Molecular Biology, 30% ANZSRC Code 100105: Genetically Modified Field Crops and Pasture, 20%

Fields of Research (FoR) Classification

FoR Code 1001: Agricultural Biotechnology, 70% FoR Code 0607: Plant Biology, 15% FoR Code 0601: Biochemistry and Cell Biology, 15%

Table of Contents

Page No. Chapter 1: Introduction………………………………………………………………… 1 1.1 Transgenic crops and food security…………………………………………..……. 1 1.2 Crop losses due to pests and the current status of insect pest management………………………………………………………………………………. 2 1.3 Insect resistant transgenic crops- scopes and benefits………………………….. 3 1.4 Insecticidal venom peptides from arthropod predators…………………………... 6 1.4.1 Insecticidal spider-venom peptides………………………………………………. 6 1.4.1.1 The insecticidal spider-venom peptide ω-HXTX-Hv1a………………………. 8 1.4.2 Insecticidal peptides from venom……………………………………… 10 1.4.3 Insecticidal peptides from centipede venom…………………………………….. 17 1.4.4 Insecticidal peptides from hymenopteran ……………………………... 19 1.5 Potential of insecticidal arthropod-venom peptides as bioinsecticides…………. 19 1.6 Genetic transformation techniques of crop plants for insect resistance………... 25 1.7 Insect-resistant plants using transgenes encoding insecticidal venom peptides 28 1.8 Minimising non-target effects via tissue-specific expression of insecticidal transgenes…………………………………………………………………………………. 32 1.9 Scope for controlling major insect pests using insecticidal venom peptides…... 34

Chapter 2: Insecticidal activity of recombinant Hv1a fusion proteins………… 36 2.1 Introduction……………………………………………………………………………. 36 2.2 Materials and methods………………………………………………………………. 40 2.2.1 Designing synthetic gene constructs for Hv1a, GNA, and GNA-Hv1a fusion proteins ……………………………………………………………………………………. 40 2.2.2 Transformation of P. pastoris expression host strain X-33……………………. 42 2.2.3 Small-scale protein production and detection of clones……………………….. 42 2.2.4 Large-scale protein production…………………………………………………… 43 2.2.5 Ni-NTA affinity purification………………………………………………………… 43 2.2.6 RP-HPLC purification and mass analysis……………………………………….. 44 2.2.7 SDS-PAGE and western blotting of the purified fusion proteins……………… 44 2.2.8 Injection bioassay to determine the insecticidal properties of recombinant fusion proteins…………………………………………………………………………….. 46 2.2.9 Feeding assay for oral activity of recombinant fusion proteins……………….. 46 2.3 Results………………………………………………………………………………… 47 2.3.1 Small-scale protein production and detection of clones……………………….. 47 2.3.2 Large-scale production of recombinant proteins……………………………….. 49

ix 2.3.3 Insecticidal activity of the recombinant fusion proteins………………………… 52 2.4 Discussion…………………………………………………………………………….. 54 2.5 Summary and conclusion……………………………………………………………. 56

Chapter 3: Genetic engineering of Arabidopsis to express Hv1a proteins…... 57 3.1 Introduction……………………………………………………………………………. 57 3.2 Materials and methods………………………………………………………………. 59 3.2.1 Fusion protein expression constructs……………………………………………. 59 3.2.1.1 Diagnostic restriction digestion of the transformation vectors plasmids…… 62 3.2.2 Transformation of A. tumefaciens by tri-parental mating………………………. 64 3.2.3 Plant materials……………………………………………………………………… 64 3.2.4 Floral-dip transformation of Arabidopsis…………………………………………. 65 3.2.5 Selection of transgenic plants using selectable markers………………………. 65 3.2.6 PCR confirmation of transgene integration…………………….………………... 66 3.2.7 Segregation analysis of transgenic lines…………………….…………………... 66 3.2.8 Detection of in planta expressed proteins…………………….…………………. 67 3.3 Results…………………….…………………….…………………….………………. 68 3.3.1 Detection of the transgenes in Arabidopsis…………………….……………….. 68 3.3.2 Detecting in planta peptide expression…………………….……………………. 68 3.4 Discussion…………………….…………………….…………………….…………... 75 3.5 Summary and conclusion…………………….…………………….……………….. 76

Chapter 4: Transgenic expression of Hv1a protects plants from lepidopteran pests…………………….…………………….…………………….…………………….. 77 4.1 Introduction……………………………………………………………………………. 77 4.2 Materials and methods………………………………………………………………. 78 4.2.1 Test plants……………………………………………………….………………….. 78 4.2.2 Test insect species………………………………………….……………………... 79 4.2.3 Insect bioassay with H. armigera feeding on transgenic Arabidopsis………... 79 4.2.3.1 Whole-plant bioassays………………………………………….……………….. 79 4.2.3.2 Detached-leaf bioassays……………………………………………….……….. 79 4.2.4 Western blot analysis of insect hemolymph …………………….……………… 80 4.2.5 Statistical analysis………………………….…………………….………………… 80 4.3 Results………………………………………………………………………………… 81 4.3.1 Resistance of transgenic Arabidopsis to H. armigera………... 81 4.3.1.1 Mortality and development of H. armigera larvae fed on Hv1a transgenic plants constructed on a WT background..…………………….……………………….. 81 4.3.1.2 H. armigera fed leaves from Hv1a transgenics in a gluc-null background… 84

x 4.3.1.3 H. armigera leaf damage on Hv1a transgenics in gluc-null background…... 86 4.3.1.4 Extended feeding assays……………………………….…………………….… 86 4.3.2 Detection of insecticidal peptides in insect hemolymph……………………….. 88 4.4 Discussion…………………………………………………………………………….. 89 4.4.1 Effect of endogenous glucosinolates on H. armigera survival………………… 89 4.4.2 In planta expression of Hv1a/GNA fusions confers resistance to cotton bollworms…………………….…………………….…………………….………………... 89 4.4.3 Comparison with other plants engineered to express Hv1a…………………... 90 4.5 Summary and conclusions…………………………………………………………... 91

Chapter 5: Summary, general discussion and future directions……………….. 92 5.1 Summary of findings…………………………………………………………………. 92 5.1.1 Recombinant production of Hv1a/GNA fusion proteins………………………... 92 5.1.2 Insecticidal activity of recombinant Hv1a/GNA fusion proteins……………….. 92 5.1.3 Transformation vector for Agrobacterium mediated plant transformation…… 92 5.1.4 Generation and characterization of transgenic Arabidopsis plants…………… 92 5.1.5 Insecticidal activity of in planta expressed toxins against H. armigera……… 93 5.2 General discussion…………………………………………………………………… 93 5.3 Future work…………………………………………………….……………………… 96

List of References…………………….…………………….…………………….…….. 98

Appendices…………………….…………………….…………………….…………….. 121

xi List of Figures & Tables

Page No. Chapter 1: Introduction Table 1.1: Development of resistance in major insect pests against Bt toxins……. 5 Figure 1.1: Schematic of an insect synapse showing the molecular targets of spider-venom components (green boxes) and chemical insecticides (red boxes).……………….. 7 Figure 1.2: Schematic of the solution structure of Hv1a (PDB accession code 1AXH)………………………………………………………………………………………. 9 Table 1.2: Insecticidal peptides isolated from spider venoms together with their molecular target and effective dose…………………………………………………….. 11 Table 1.3: Insecticidal peptides isolated from scorpion venoms together with their molecular target and effective dose…………………………………………………….. 14 Table 1.4: Insecticidal peptides isolated from centipede venoms together with their molecular target and effective dose………………………………………………. 18 Figure 1.3: Structure of GNA tetramer (PDB accession code 1MSA)……………… 21 Figure 1.4: Lectin-mediated delivery of peptide neurotoxins………………………... 22 Figure 1.5: The circulative route of luteoviruses in aphids.………………………….. 24 Table 1.5: Insect-resistant transgenic plants developed so far with insecticidal venom peptides…………………………………………………………………………… 30 Figure 1.6: Distribution of fluorescence in source leaves when GFP was expressed under the control of CaMV 35S or SUC2 promoters…………………….. 33

Chapter 2: Insecticidal activity of recombinant Hv1a fusion proteins Figure 2.1: Sequences of Hv1a, GNA, and fusions thereof that were used to design synthetic genes for expression in P. pastoris…………………………………. 40 Figure 2.2: pPICZα Expression vector for P. pastoris expression system (Invitrogen, 2010)…………………………………………………………………………. 41 Table 2.1: Recipe for separating and staking gels for 16.5% polyacrylamide gel… 45 Figure 2.3: Western blot (probed with rabbit anti-GNA polyclonal antibody) showing expression of GNA and Hv1a-GNA in P. pastoris………………………….. 48 Figure 2.4: Western blot (probed with rabbit anti-GNA polyclonal antibody) showing expression of GNA-HV1a……………………………………………………… 48 Figure 2.5: Purification of recombinant Hv1a expressed in P. pastoris……………. 49 Figure 2.6: Purification and detection of recombinant GNA expressed in P. pastoris…………………………………………………………………………………. 50 Figure 2.7: Purification and detection of recombinant Hv1a-GNA expressed in P. pastoris…………………………………………………………………………………. 51 Figure 2.8: Purification and detection of recombinant GNA-Hv1a expressed in P. pastoris…………………………………………………………………………………. 52

xii Table 2.2: Insecticidal activity of recombinant Hv1a, GNA, GNA-Hv1a, and Hv1a- GNA after injection into houseflies (Musca domestica)………………………………. 53 Figure 2.9: Feeding damage by P. xylostella larvae on treated Arabidopsis leaves………………………………………………………………………………………. 54

Chapter 3: Genetic engineering of Arabidopsis to express Hv1a proteins Figure. 3.1: Amino acid sequences encoding Hv1a (red), GNA (green), CP (purple), and fusions of Hv1a to either the N- or C-terminus of GNA (GNA-Hv1a and Hv1a-GNA) or CP (CP-Hv1a and Hv1a-CP)……………………………………… 60 Figure 3.2: Synthetic genes encoding the designed peptide sequences………….. 61 Figure 3.3: Schematic representation of pAOV (Mylne and Botella, 1998)……….. 61 Figure 3.4: Diagnostic restriction digestion strategies for (A) vectors containing the 35S promoter, and (B) vectors containing the SUC2 promoter…………………. 63 Table 3.1: Expected fragment size after double digestion of the constructs………. 63 Figure 3.5: Standard floral dip method to transform Arabidopsis. …………………. 65 Figure 3.6: PCR analysis of transgenic Arabidopsis lines generated on a WT background………………………………………………………………………………... 69 Figure 3.7: PCR analysis of transgenic Arabidopsis lines in WT background…….. 70 Figure 3.8: PCR analysis of transgenic Arabidopsis lines generated in the gluc- null background…...………………………………………………………………………. 70 Figure 3.9: PCR analysis of transgenic Arabidopsis lines generated in WT background.……………………………………………………………………………….. 71 Figure 3.10: Western blot analysis of 35S-Hv1a homozygous transgenic Arabidopsis lines, probed with rabbit polyclonal anti-Hv1a antibody (1:2000 dilution)…………………………………………………………………………………….. 72 Figure 3.11: Western blot analysis of 35S-GNA homozygous transgenic Arabidopsis lines, probed with rabbit polyclonal anti-GNA antibody (1:2000 dilution)………………...... 72 Figure 3.12: Western blot analysis of 35S-Hv1a-GNA homozygous transgenic Arabidopsis lines probed with rabbit polyclonal anti-Hv1a antibody (1:2000 dilution).……………………………………………………………………………………. 73 Figure 3.13: Western blot analysis of 35S-GNA-Hv1a homozygous transgenic Arabidopsis lines, probed with rabbit polyclonal anti-Hv1a antibody (1:2000 dilution).……………………………………………………………………………………. 73 Figure 3.14: Western blot analysis of (A) 35S-Hv1a-CP and (B) 35S-CP-Hv1a homozygous transgenic Arabidopsis lines, probed with rabbit polyclonal anti-Hv1a antibody (1:2000 dilution)………………………………………………………………… 73 Figure 3.15: Western blot analysis of proteins in 35S homozygous transgenic Arabidopsis lines in the gluc-null background, probed with either (A) rabbit polyclonal anti-Hv1a antibody (1:2000 dilution), or (B) rabbit polyclonal anti-GNA antibody (1:2000 dilution)………………………………………………………………... 74

xiii Chapter 4: Transgenic expression of Hv1a protects plants from lepidopteran pests

Figure 4.1: The life cycle of the cotton bollworm (H. armigera) (Stevens et al., 2012)……………………………………………………………………………………….. 77 Figure 4.2: Set up of whole-plant insect bioassay……………………………………. 79 Figure 4.3: Set up of detached-leaf insect bioassay with H. armigera……………... 80 Figure 4.4: Insect bioassay with H. armigera neonates feeding on leaves of WT or transgenic Arabidopsis plants expressing Hv1a, GNA, GNA-Hv1a, or Hv1a-GNA in a WT background………………………………………………………… 82 Figure 4.5: (A) Mortality and (B) weight-gain observed for H. armigera neonates fed on WT and gluc-null Arabidopsis for 9 days. Data are mean ± SEM. (C) Photograph of larvae after feeding on WT and gluc-null Arabidopsis for 9 days….. 83 Figure 4.6: (A) Kaplan-Meier survival curves for H. armigera neonates feeding on leaves of gluc-null plants or transgenic plants expressing Hv1a, GNA, Hv1a-GNA or GNA-Hv1a generated in a gluc-null Arabidopsis background. (B) Weight of larvae (Mean ± SEM) after 9 days of feeding on leaves of gluc-null or transgenic Arabidopsis lines………………………………………………………………………….. 85 Figure 4.7: Consumption of leaf tissue by H. armigera larvae during five days of feeding on detached leaves of gluc-null control or different transgenic plants in gluc-null background……………………………………………………………………… 86 Figure 4.8: Kaplan-Meier survival curves for H. armigera neonates fed for 15 days with leaves from gluc-null, gluc-null/Hv1a or gluc-null/GNA-Hv1a plants……. 86 Figure 4.9: Severity of leaf tissue damage caused by H. armigera larvae supplied continuously for 11 days with leaves from gluc-null plants or gluc-null plants expressing Hv1a or GNA-Hv1a…………………………………………………………. 88 Figure 4.10: Western blot analysis of hemolymph from dead larvae fed on different transgenic leaves. Proteins were detected using an anti-Hv1a polyclonal antibody……………………………………………………………………………………. 88

xiv List of Abbreviations used in the thesis

Nucleic acid abbreviations: A: Adenine T: Thymine G: Guanine C: Cytosine Amino acid abbreviations: Single letter Three letter Single letter Three letter Amino Acid Amino Acid code code code code Alanine A Ala Isoleucine I Ile Arginine R Arg Leucine L Leu Asparagine N Asn Lysine K Lys Aspartic acid D Asp Methionine M Met Phenyl- Cystiene C Cys F Phe alanine Glutamic E Glu Proline P Pro acid Glutamine Gln S Ser Q Serine

Glycine G Gly Threonine T Thr Histidine H His Tryptophan W Trp Valine V Val Tyrosine Y Tyr

Other abbreviations: %: per cent 35S: Cauliflower Mosaic Virus 35S promoter AChE: acetylcholinesterase ANOVA: Analysis of variance AOX: Alcohol oxidase enzyme

BKCa: calcium-activated big potassium channel bp: base pairs Bt / Bt: ButaIT: Indian red scorpion (Mesobuthus tamulus) toxin CaMV: Cauliflower mosaic virus

CaV: Voltage gated calcium channel cm: centimetre CNS: Central nervous system CPU: Contraction paralysis unit d.f.: Degrees of freedom DNA: Deoxy-ribonucleic acid E. coli: Escherichia coli

xv EC50: The half maximal effective concentration,

ED50: the amount of a toxin, which produces an effect on 50% of test EDTA: Ethylene diamine tetra-acetic acid ESI-MS: Electrospray ionization mass spectrometry Fig.: Figure GABA: γ-aminobutyric acid receptor gDNA: Genomic deoxy-ribonucleic acid GM: Genetically modified GNA: Galanthus nivalis agglutinin gluc-null: Glucosinolates null GluR: ionotrophic glutamate receptor

His6: HHHHHH (hexa-histidine) Hv1a: ω-HXTX-Hv1a from Australian Blue Mountains funnel-web spider venom HVA: High voltage activated

IC50: The half maximal inhibitory concentration ISVP: Insecticidal spider venom peptide kD / kDa: KiloDalton Kd: The equilibrium dissociation constant

KV: Voltage gated potassium channel L: Litre LB: Luria-Bertani broth

LD50: amount of a toxin that causes the death of 50% of test animals LVA: Low-voltage-activated M: Molar MALDI: Matrix-assisted laser desorption ionization mass spectrometry mg: Milligram min: Minutes mL: Millilitre mM: Millimolar mRNA: Messenger ribonucleic acid MS: Mass spectrometry Mut–: Methanol utilization minus Mut+: Methanol utilization plus Muts: Methanol utilization slow

Nav: Voltage gated sodium channel nAChR: nicotinic acetylcholine receptor ND: Not determined

xvi ng: Nanogram Ni-NTA: Nickel-nitrilotriacetic acid nm: Nanometer NMDA receptor: N-methyl-D-aspartate receptor NMJ: neuromuscular junction NMR: Nuclear magnetic resonance oC: Degree Celsius OD: Optical density ORF: Open reading frame PAGE: Polyacrylamide gel electrophoresis PBS: Phosphate buffered saline PCR: Polymerase chain reaction

PD50: the amount of a toxin that causes the paralysis of 50% of test animals. pH: decimal logarithm of the reciprocal of the hydrogen ion activity in a solution PIs: inhibitors pSlo: Cockroach (Periplaneta americana) Slowpoke (dSlo) calcium-activated potassium channels rcf: Relative centrifugal force Rif50: Rifampicin 50 µg/mL RNA: Ribonucleic acid RP-HPLC: Reverse phase high performance liquid chromatography rpm: Revolutions per minute s: Second SDS-PAGE: Sodium dodecyl sulphate polyacrylamide gel electrophoresis SDS: Sodium dodecyl sulphate SS-rich: disulfide-rich Strep25: Streptomycin 25 µg/mL SUC2: Arabidopsis thaliana SUCROSE TRANSPORTER 2 promoter (At1g22710) SVP: Spider venom peptide T-DNA: Transfer DNA

T0: Untransformed plants

T1: Primary transformed plants

T2: Second generation transgenic plants

T3: Third generation transgenic plants

Ti: Tumor inducing plasmid TAE: Tris acetate EDTA buffer TBE: Tris borate EDTA buffer

xvii TE: Tris-EDTA Tet2: Tetracycline 2 µg/mL TEV: Tobacco etch virus w/w: weight / weight WT: Columbia 1 ecotype (wild type) Arabidopsis thaliana YNB: Yeast nitrogen base YPD: Yeast peptone dextrose µg: Microgram µL: Microliter µm: Micrometre µM: Micromolar

xviii

Chapter 1: Introduction

1.1 Transgenic crops and food security

Transgenic, or genetically modified (GM), crops are those in which their genetic make-up has been modified by incorporation of one or more heterologous genes from the same or another species. Contemporary molecular biology techniques can be used to insert transgenes into plant genomes in order to create or enhance desirable characteristics such as resistance or tolerance to biotic or abiotic stresses such as pest insects, disease, drought, salinity and temperature. GM crops represent the most widespread application of biotechnology in agriculture (James, 2012).

The ever-increasing human population is creating a continuous demand to increase food production, particularly in the developing regions of Asia, Africa, and Latin America (Boulter et al., 1990; Sharma et al., 2004). Developed countries are also facing a big challenge to maintain food security. Yet, at the same time, urbanisation and industrialisation are hijacking arable lands day by day. Global climate change is also threatening global food security by facilitating the emergence and spread of crop pests and pathogens and also by allowing their establishment in previously unsuitable regions (Bebber et al., 2013). Thus, there is a major incentive to increase yields from major crops grown on existing cultivated land (Hilder and Boulter, 1999). As stated by the Food and Agricultural Organization of the United Nations (FAO): “To feed a growing world population, we have no option but to intensify crop production. But farmers face unprecedented constraints. In order to grow, agriculture must learn to save” (FAO, 2011). The threat of a food security crisis has given fresh stimulus to the long-standing debate about the potential contribution of agricultural biotechnology to food security (Dibden et al., 2013).

Despite some controversies about adoption of GM crops (Mendelsohn et al., 2003; Qaim and Kouser, 2013), the area devoted to their cultivation reached more than 170 million hectares within 16 years of their introduction in 1996 (James, 2012). As of 2012, 17.3 million farmers in about 30 countries were growing GM crops on 170.3 million hectares of land (James, 2012). Four GM crops dominate global agriculture, with GM cotton, soybean, maize, and canola accounting for 81%, 81%, 35% and 30%, respectively, of global production (James, 2012). The most important traits introduced into GM crops so far are resistance to herbicide and insect pests (Hilder and Boulter, 1999). In the following sections, we review the potential of existing insect-resistant transgenic plants and the scope for development of new GM crops expressing insecticidal venom peptides.

1

1.2 Crop losses due to arthropod pests and the current status of insect pest management

Arthropod pests, primarily insects, are the major cause of reductions in crop yield and quality (Oerke, 2006). Consequently, their control is critical for achieving optimal crop yields. Around 10% of over 10,000 species of arthropod pests are responsible for global pre- and post-harvest crop losses of approximately 20–50% of potential production (Thacker, 2002). These losses occur despite current pest control strategies, including application of chemical insecticides and biological control methods. Phytophagous (plant-eating) insects are the major cause of this crop loss. These include insect species from the orders Lepidoptera (moths and butterflies), Orthoptera (locusts and grasshoppers), and Coleoptera (beetles) (Novotny et al., 2002). The lepidopteran larvae are considered the most destructive and about 40% of total chemical insecticides being used to control them. (McCaffery, 1998). Insect species from the orders Diptera (flies), Hemiptera (plant sap sucking bugs), Thysanoptera (thrips) and Acarina (mites) are also recognised as important crop pests (McCaffery, 1998; Nicholson, 2007). In addition to directly damaging crops, insects also vector plant viruses (Nicholson, 2007; King and Hardy, 2013). In addition to the economic cost of direct and indirect losses caused by insect pests, there are additional costs in the form of insecticides applied for insect pest control, which amount to ~US $11 billion annually (2007 estimate) (Grube et al., 2011).

Arthropod pest control relies heavily on synthetic chemical insecticides (Whetstone and Hammock, 2007) Commercially available chemical insecticides mostly act on one of just six molecular targets in insect body and nervous systems: voltage-gated sodium (NaV) channels (e.g., DDT, dihydropyrazoles, oxadiazines and pyrethroids); the nicotinic acetylcholine receptor (e.g., spinosad and imidacloprid); acetylcholinesterase (organophosphates and carbamates); glutamate receptor (avermectins); GABA-gated chloride channels (e.g., cyclodienes and fipronil); and ryanodine receptors (e.g., Rynaxypyr and Cyazypyr) (Tedford et al., 2004b; Nicholson, 2007; Sattelle et al., 2008)). Indiscriminate use of some of these insecticides over many decades has provided intense selection pressure for the development of insecticide resistance (Feyereisen, 1995; Brogdon and McAllister, 1998). More than 600 major arthropod pests have developed resistance to one or more classes of chemical insecticide (King and Hardy, 2013). Insecticide resistance can result from (i) enhanced metabolic detoxification of insecticides (e.g., from elevated levels of esterases, glutathione S-transferase, and/or monooxygenases); (ii) decreased target sensitivity through subtle point mutations;

2

(iii) increased sequestration or lowered insecticide availability (Feyereisen, 1995; Brogdon and McAllister, 1998; Hemingway and Ranson, 2000).

Excessive use of chemical insecticides can expose farmers to serious health risks or have adverse ecological consequences by affecting non-target organisms. Often less than 1% of insecticides actually reach the targeted pest organism; the remainder contaminates the air, soil and water (Pimentel and Levitan, 1986). Prenatal exposures to organophosphate insecticides, which were widely used until recently, as well as pyrethroid insecticides, can have negative effects on fetal development, resulting in adverse birth outcomes (Rauch et al., 2012; Greenop et al., 2013; Van Maele-Fabry et al., 2013), and male reproduction (reduction in sperm quality, damaged sperm DNA, and reproductive hormone disorders) (Koureas et al., 2012). Elevated serum levels of dichlorodiphenyldichloroethylene, a metabolite of the insecticide dichlorodiphenyltrichloroethane (DDT), are reported to increase the risk for Alzheimer's disease (Richardson et al., 2014). The potential adverse effect of chemical insecticides on human health also makes consumers more concerned about insecticide residues in food (FAO, 2011).

Although the immense contribution of chemical insecticides to increases in global agricultural production over the past half-century cannot be overlooked, the scope for use of chemical insecticides to control arthropod pests is declining, largely because of the cancellation or withdrawal of chemical insecticides from the market due to the development of insect resistances, environmental legislation, lack of discovery of new chemical insecticides, environmental activism, and public pressure (King and Hardy, 2013; Smith et al., 2013). Thus, a major challenge for the future is to increase crop productivity in a sustainable manner with less reliance on chemical insecticides. This has necessitated the development of target-specific compounds with low persistence and a greater emphasis on integrated pest management based on host-plant resistance to insect pests. However, there is still a great need to develop alternative or additional technologies, which can minimise dependency on chemical insecticides. The development of GM crops that are resistant to insect pests is one potential strategy for addressing this challenge.

1.3 Insect resistant transgenic crops - scopes and benefits

From the beginning of agriculture, humans have searched for crop plants that can tolerate and survive attack by insect pests. However, conventional host-plant resistance to insects involves quantitative traits at several loci, and as a result, progress has been slow and

3

difficult to achieve. But, with the development of genetic engineering techniques, genes for insect resistance now can be engineered into plants more quickly and deliberately. The ability to isolate and manipulate single genes through recombinant DNA technology together with the ability to insert specific genes into a chosen plant species has opened a new era of targeted plant breeding. Significant progress has been made over the past three decades in introducing foreign genes into plants, and this has provided opportunities to modify crops to obtain desired traits. With the advent of genetic transformation techniques based on recombinant DNA technology, it is now possible to insert insect-resistance transgenes into plant genomes. Genetic engineering opened up the possibility of using insecticidal genes from different sources for breeding insect-resistant plants.

The advantages of genetically-engineered endogenous insect-resistance traits relative to exogenous chemical control are: (1) crops are provided with season-long protection, thus preventing pest populations from building up; (2) the protection is independent of weather; (3) protection is afforded to plant tissues that are difficult to reach with foliar sprays; (4) insects are affected at their most sensitive stage; (5) only crop-eating insects are exposed; (6) the protectant is confined to plant tissues; and (7) the active factor is biodegradable and usually non-toxic to man and animals (Gatehouse, 2008).

The first commercially available insect-resistant GM contained transgenes encoding insecticidal toxins from Bacillus thuringiensis (Bt). Bt is a naturally occurring spore-forming bacterium from the family Bacillaceae. B. thuringiensis is widespread in soil and lethal to a range of insects. Sporulation results in formation of insecticidal crystal protein to form protoxin crystals (Bt crystals). Bt crystals contain Cry toxins (Bravo et al., 2011). Cry toxins are pore-forming toxins responsible for the insecticidal activity of Bt. Cry toxins are toxic to certain species of insects, especially larval stages, and they have proved to be safe to humans and other vertebrates. Bt crystals have to be ingested to result in insecticidal activity and insect death. After ingestion, Bt crystals dissolve within the insect gut and form the active toxin (Cry toxin). The Cry toxin binds to receptors in the midgut to form pores, which results in insect paralysis and/or bacterial septicaemia (Bravo et al., 2007; Gatehouse, 2008; Gatehouse et al., 2011). resistant to lepidopterans and Bt corn resistant to both lepidopterans and coleopterans have become widely used in global agriculture and have led to significant reductions in insecticide use on these crops, as well as lower production costs (Toenniessen et al., 2003).

The insect-resistant trait based on Bt toxins has become an indispensable tool in modern agriculture. There are many advantages of Bt crops including reduction in harmful

4

chemical insecticide usage, insect pest suppression, reduced production costs, conservation of beneficial natural enemies, increased yield, and higher farmer profits that make them a valuable and commercially adaptable pest management tool (Tabashnik et al., 2013). The Bt toxins used in GM crops target the digestive system of lepidopterans and coleopterans, and hence they have no effect on sap-sucking insects. As a result, sap-suckers that used to be secondary pests have become major pests on some GM crops, resulting in increased use of insecticides to control them (Zhao et al., 2011). Moreover, there are now five confirmed cases of insects evolving resistance to Bt toxins in the field: Bt cotton in India (2009) and the USA (2002), moth pests in maize in Puerto Rico (2008) and South Africa (2006), and a beetle pest in maize in the USA (2009) (Table 1.1) (Tabashnik et al., 2013). In each of these cases, resistance developed despite deployment schemes to prevent resistance, including the use of Bt lines expressing high levels of toxin grown alongside non-Bt plant refuges for susceptible insects. As a consequence of this increased resistance, insecticide costs on infested Bt crops have risen by nearly a third (Tabashnik et al., 2013).

Table 1.1: Development of resistance in major insect pests against Bt toxins Target insect Year Year (resistance Country Crops Toxin References pest species (commercialized) first reported) (Ali et al., 2006; Helicoverpa zea Cry1Ac 1996 2002 USA Bt-Cotton Ali and Luttrell, (Lepidoptera) Cry1Ab 2003 2005 2007) Spodoptera (Storer et al., frugiperda USA Bt-Corn Cry1F 2003 2008 2010; Storer et (Lepidoptera) al., 2012) (van Rensburg, Busseola fusca South Bt-Corn Cry1Ab 1998 2006 2007; Tabashnik (Lepidoptera) Africa et al., 2009) Pectinophora (Dhurua and gossypiella India Bt-Cotton Cry1Ac 2002 2009 Gujar, 2011) (Lepidoptera) Diabrotica (Gassmann et al., virgifera virgifera USA Bt-Corn Cry3Bb 2003 2009 2011; Devos et (Coleoptera) al., 2013)

Questions about the long-term effectiveness of transgenic Bt crops have now become a major issue. One potential strategy to avoid or delay the evolution of insect resistance is trait-stacking or "pyramiding", whereby transgenes encoding toxins with different modes of action are engineered into the same plant. For example, transgenic plants expressing combinations of Cry toxins that interact with different mid-gut receptors were shown to delay the development of resistance compared to plants expressing a single insecticidal transgene (Gould, 2003; Zhao et al., 2003; Zhao et al., 2005). Trait stacking can also be used to broaden the range of affected insects (i.e., by stacking toxins with different host ranges) and to minimize secondary pest infestations (Tu et al., 2000; Chitkowski et al., 2003). In addition to Bt toxins, there are a host of insecticidal proteins that could be

5

expressed in GM crops, including protease inhibitors, neurotoxins from venoms, chitinase, and plant lectins. These could be used alone or in combination with Bt genes to generate transgenic plants for insect pest control. These alternative insect resistance genes could be particularly useful in cases where lepidopteran and coleopteran insect pests have developed resistance to plants engineered to express Bt toxins. In the following sections, we examine insecticidal venom peptides that have the potential to provide insect-resistance in GM crops.

1.4 Insecticidal venom peptides from arthropod predators

A diverse range of animals produce venoms for prey capture and defence, including arthropod predators such as ( and ), hymenopterans (wasps, bees and ants), and centipedes; aquatic animals such as cone snails and cnidarians (sea anemones and jellyfish), and vertebrates such as snakes and the platypus. Arthropod predators such as spiders, scorpions, and centipedes mainly use their venom to catch insect prey, and consequently their venoms contain many insecticidal peptides (Schwartz et al., 2012; King and Hardy, 2013; Smith et al., 2013). Due to hundreds of millions of years of evolutionary selection pressure, these insecticidal toxins have developed remarkable selectivity and potency for their molecular targets (Smith et al., 2013). In the venoms of spiders alone, which are among the world’s most successful insect predators, it is estimated that millions of insecticidal toxins are yet to be discovered (Schwartz et al., 2012; King and Hardy, 2013; Smith et al., 2013). Venom peptides from these predators can act with high affinity on ion channels, membrane receptors, and neurotransmitter transporters in the nervous system of prey (Schwartz et al., 2012; Smith et al., 2013).

1.4.1 Insecticidal spider-venom peptides

Spiders are the most successful venomous and one of the most abundant terrestrial predators (Windley et al., 2012). The number of extant spider species (~45,000; see http://research.amnh.org/iz/spiders/catalog/counts.html) predicted to be greater than 150,000 (Coddington and Levi, 1991), is likely to be larger than the total number of venomous predators in all other terrestrial phyla (King and Hardy, 2013). Along with their ingenious exploitation of silk, the remarkable evolutionary success of spiders is due to the evolution of a pharmacologically complex venom, the main purpose of which is to ensure rapid subjugation of prey (King and Hardy, 2013).

6

The chemical complexity of spider venoms is extraordinary, ranging from salts and small organic compounds to large presynaptic neurotoxins (Escoubas et al., 2000a; Rash and Hodgson, 2002; Tedford et al., 2004b; Estrada et al., 2007; Vassilevski et al., 2009; Kuhn- Nentwig et al., 2011). These venom compounds can be broadly grouped into five classes on the basis of their chemical structure and mechanism of action (King and Hardy, 2013) namely: (i) salts and small organic compounds; (ii) linear cytolytic peptides; (iii) disulphide-rich (SS-rich) peptide neurotoxins; (iv) enzymes; and (v) large presynaptic neurotoxins. SS-rich peptides are the dominant compounds in most spider venoms and they are the major contributors to the venom’s insecticidal activity; they typically target presynaptic ion channels or postsynaptic receptors either at peripheral neuromuscular junctions or at synapses in the insect central nervous system (CNS) (Fig. 1.1) (King and Hardy, 2013).

Figure 1.1: Schematic of an insect synapse showing the molecular targets of spider-venom components (green boxes) and chemical insecticides (red boxes). Spider venoms contain enzymes that facilitate access of peptide and protein neurotoxins to their molecular targets by degrading the myelin sheath around axons as well as the extracellular matrix of the synaptic cleft. The α- cause massive neurotransmitter release by promoting synaptic vesicle exocytosis. Figure and caption from (King and Hardy, 2013).

7

Insecticidal compounds derived from spider venom are mainly the small SS-rich peptide neurotoxins (King and Hardy, 2013; Smith et al., 2013). These insecticidal peptides either alone or together target the insect nervous system resulting in flaccid paralysis by blocking or convulsive paralysis by stimulating to rapidly debilitate the envenomed insect prey (King and Hardy, 2013).

Most spiders prey on , mainly insects, although other arachnids such as mites, opilionids, and both conspecific and nonconspecific spiders often contribute to their diet (Kuhn-Nentwig et al., 2011). Thus, spider venoms have evolved to target a broad spectrum of insects. Hence, the primary rationale for investigating spider venoms as a potential source of bioinsecticides is that their venoms are expected to contain a wide array of insecticidal peptides that mostly have little or no activity. The past two decades of research on spider venoms has largely validated this hypothesis.

To date, more than 200 SS-rich insecticidal spider-venom peptides (SVPs) have been sequenced (Maggio et al., 2005; Windley et al., 2012). They range in size from 3.3 to 9.0 kDa and contain 3–6 disulfide bonds (King and Hardy, 2013). However, only several −1 dozen of these peptides are sufficiently potent (LD50 < 1500 pmol g ) to deserve serious consideration as bioinsecticides, and even fewer have been shown to be harmless to vertebrates (King and Hardy, 2013). Table 1.2 provides a summary of characterised insecticidal SVPs. The absence of detrimental effects in vertebrates injected with a considerable number of SVPs indicates their selectivity for insects (King and Hardy, 2013).

Many insecticidal SVPs have molecular targets that are distinct from those of extant chemical insecticides, including CaV channels, NMDA receptors, and glutamate transporters (King and Hardy, 2013). Some of these, such as CaV channels, have been validated as insecticide targets by gene knockout and inducible expression of SVP transgenes in Drosophila melanogaster (King, 2007b). Thus, in addition to their potential as bioinsecticides, insecticidal SVPs have helped expand the range of validated insecticide targets (King and Hardy, 2013).

1.4.1.1 The insecticidal spider-venom peptide ω-HXTX-Hv1a

The venom of the Australian Blue Mountains funnel-web spider Hadronyche versuta is a cocktail of around 500 peptides (Escoubas et al., 2006). Almost 70% of these peptides range in size from 3 to 5 kDa (24–45 residues) and contain 3–4 disulfide bonds (Tedford et al., 2004b). Most of these peptides contain a conserved arrangement of three disulfide

8

bonds that form an inhibitor cystine knot (ICK) motif in which two of the disulfides and the intervening sections of the peptide backbone form a loop that is bisected by the third disulfide bond (Fig. 1.2B). The ICK motif typically provides SVPs with a high level of chemical and thermal stability as well as resistance to (Saez et al., 2010). Several families of peptides have been identified from H. versuta venom that are toxic to a wide range of insect pests but harmless to vertebrates (Tedford et al., 2004b).

ω-HXTX-Hv1a (hereafter referred to as Hv1a) is the most potent insecticidal peptide described thus far from the venom of H. versuta (Tedford et al., 2004b). Hv1a specifically targets Cav channels in the insect CNS but is harmless to vertebrates (Fletcher et al., 1997; Tedford et al., 2004a; Tedford et al., 2004b; Chong et al., 2007). This 37-residue peptide has potent insecticidal activity in a wide range of insect orders, including Lepidoptera, Homoptera, and Diptera (Khan et al., 2006; Mukherjee et al., 2006), but it is harmless to honeybees (Nakasu et al., 2014). Thus, Hv1a is considered a strong candidate for development as a novel bioinsecticide (King, 2007a; King and Hardy, 2013; Nakasu et al., 2014).

Figure 1.2: Schematic of the solution structure of Hv1a (PDB accession code 1AXH). (A) The two β strands are represented by gold arrows, while the three SS bonds are shown as blue tubes. (B) The inhibitor cystine knot (ICK) motif of Hv1a in which a closed loop formed by the Cys11–Cys22 and Cys4–Cys18 SS bonds (red) and the intervening sections of the peptide backbone (green) are bisected by the Cys17-Cys37 SS bond (blue). (C) Pharmacophore of Hv1a. Residues Pro10, Asn27, and Arg35 are the three most important residues for modulating the activity of insect

Cav channels. Figure from (King, 2007b).

There are three main regions within the three-dimensional structure of Hv1a (Fig. 1.2A): a structurally disordered N-terminus (residues 1–3), an SS-rich globular core (residues 4-21) and a β-hairpin at the C-terminus (residues 22–37) that protrudes from the SS-rich

9

core (Tedford, 2001; Tedford et al., 2004a). The three disulfide bonds form an ICK motif in which the loop formed by the Cys11–Cys22 and Cys4–Cys18 SS bonds and the intervening sections of the peptide backbone is pierced by the Cys17–Cys37 SS bond (Fig. 1.2B) (Tedford, 2001; Tedford et al., 2004a). The binding site of Hv1a on insect

Cav channels is not known but three residues on the toxin (Pro10, Asn27 and Arg35; see

Fig. 1.2C) mediate its interaction with CaV channels (Tedford, 2001; Tedford et al., 2004a; King, 2007b; King et al., 2008).

1.4.2 Insecticidal peptides from scorpion venom

Scorpions were the first to emerge from the sea to conquer the terrestrial environment more than 350 million years ago (Froy and Gurevitz, 2003). There are ~1500 extant species with conserved morphology (Dehesa-Davila et al., 1994; Chowell et al., 2006). Scorpions can survive in extreme environments; they remain hidden during the day and emerge at night to feed on small insects (Schwartz et al., 2012). As for spiders, scorpion venom is a complex mixture of small molecules such as serotonin and histamine, protease inhibitors, SS-rich neurotoxic peptides, enzymes such as hyaluronidase and phospholipase, and mucopolysaccharides (Goudet et al., 2002; Fernandes-Pedrosa et al., 2013).

The SS-rich peptides in scorpion venom interact with a variety of ion channels but most of the insecticidal peptides isolated to date from scorpion venoms target NaV channels (see Table 1.3). These toxins have been categorised into several classes depending on whether they delay NaV channel inactivation (anti-insect alpha toxins and alpha-like toxins) or have affects on channel activation.

10

Table 1.2: Insecticidal peptides isolated from spider venoms together with their molecular target and effective dose Molecular Venom peptide Source weight Target Effective dose (kDa) µ-agatoxin-Aa1a Agelenopsis 4.273 Nav LD50 6,550 pmol/g (Manduca sexta) (Skinner et al., 1989) aperta µ-agatoxin-Aa1b A. aperta 4.110 Nav LD50 18,250 pmol/g (M. sexta) (Skinner et al., 1989) µ-agatoxin-Aa1c A. aperta 4.197 Nav LD50 6,670 pmol/g (M. sexta) (Skinner et al., 1989) µ-agatoxin-Aa1d A. aperta 4.208 Insect specific Nav LD50 9,510 pmol/g (M. sexta) (Skinner et al., 1989) µ-agatoxin-Aa1e A. aperta 4.208 Nav LD50 11,410 pmol/g (M. sexta) (Skinner et al., 1989) µ-agatoxin-Aa1f A. aperta 4.168 Nav LD50 9,120 pmol/g (M. sexta) (Skinner et al., 1989) µ-agatoxin-Hc1a Hololena curta 4.111 Nav LD50 4,860 pmol/g (Acheta domestica) (Stapleton et al., 1990) µ-agatoxin-Hc1b H. curta 4.197 Nav LD50 953.06 pmol/g (A. domestica) (Stapleton et al., 1990; Quistad et al., 1991) µ-agatoxin--Hc1c H. curta 4.245 Nav LD50 942.29 pmol/g (A. domestica) (Stapleton et al., 1990) U1-CUTX-As1c Apomastus 8.327 ND LD50 2.4 pmol/g by subcutaneous injection (Skinner et al., 1992) schlingeri µ -cyrtautoxin-As1a Apomastus 3.769 Nav LD50 133 pmol/g (M. sexta) (Skinner et al., 1992) schlingeri LD50 > 2.66 pmol/g (Spodoptera exigua) (Skinner et al., 1992) PD50 700 ± 35 pmol/g (Lucilia cuprina) (Bende et al., 2013) δ-ctenitoxin-Pn1a Phoneutria 9.088 Insect specific Nav LD50 0.418 pmol/house fly (Musca domestica) (Figueiredo et al., 1995) nigriventer µ-diguetoxin-Dc1a Diguetia canities 6.975 Insect specific Nav PD50 3,800 pmol/g (Lepidopteran larvae) (Krapcho et al., 1995) U1-NETX-Csp1a Calisoga sp. 8.946 Insect specific PD50 0.265 pmol/g (Heliothis virescens) (Johnson et al., 1997) Paralysis U1-AGTX-Ta1a Tegenaria 7.741 Insect specific PD50 890 pmol/g (Tobacco budworm), PD50 780 pmol/g (cabbage looper), agrestis ND, PD50 900 pmol/g (beet armyworm) and PD50 2,000 pmol/g (southern corn Direct effect rootworm) (Johnson et al., 1998). on CNS

11

Table 1.2: Insecticidal peptides isolated from spider venoms together with their molecular target and effective dose (Continued) Molecular Venom peptide Source weight Target Effective dose (kDa) ω-hexatoxin-Hv1a Hadronyche 4.055 Insect specific M- LD50 89.0 pmol/g (A. domestica) (Wang et al., 1999; Mukherjee et al., versuta LVA Cav 2006) HVA Cav LD50 77.0 pmol/g (M. domestica) (Mukherjee et al., 2006) IC50 279,000 pmol (M-LVA) (Chong et al., 2007) LD50 1080,000 pmol (HVA) (Chong et al., 2007) κ-hexatoxin-Hv1a Hadronyche 3.685 BK Ca LD 303 ± 42 pmol g-1 (A. domestica) (Wang et al., 2000) versuta 50 κ-hexatoxin-Hv1b Hadronyche 3.651 BK Ca LD 214 ± 16 pmol g-1 (A. domestica) (Wang et al., 2000) versuta 50 -1 κ-hexatoxin-Hv1c Hadronyche 3.768 BKCa LD50 167 ± 10 pmol g (A. domestica) (Wang et al., 2000; Gunning et al., versuta 2008) LD50 320 ± 20 pmol/g (M. domestica)(Maggio and King, 2002b) LD50 91 ± 5 pmol/g (M. domestica) (Maggio and King, 2002a). δ-AMATX-PI1a Pireneitega 4.047 Na v LD 2350 pmol/g (S. litura) (Corzo et al., 2000) luctuosa 50 δ-AMATX-PI1b Pireneitega 4.124 Na v LD 5870 pmol/g (S. litura) (Corzo et al., 2000; Corzo et al., 2005) luctuosa 50 δ-AMATX-PI1c Pireneitega 3.934 Na v LD 3130 pmol/g (S. litura) (Corzo et al., 2000) luctuosa 50 δ-AMATX-PI1d Pireneitega 4.062 Nav LD50 > 11,030 pmol/g (S. litura) (Corzo et al., 2000) luctuosa ω-hexatoxin-Hv2a Hadronyche 4.484 HVA Cav PD50 160 ± 9 pmol/g (A. domestica) (Wang et al., 2001) versuta EC50~139 pmol (bee brain neurons) (Wang et al., 2001) µ-hexatoxin-Mg1a 14.188 Na v LD 17,600 pmol/g (Spodoptera litura) (Corzo et al., 2003) gigas 50 µ-hexatoxin-Mg2a Macrothele 5.233 Insect specific LD50 > 32,800 pmol/g (S. litura) (Corzo et al., 2003) gigas Nav

12

Table 1.2: Insecticidal peptides isolated from spider venoms together with their molecular target and effective dose (Continued) Molecular Venom peptide Source weight Target Effective dose (kDa) ω-theraphotoxin- Haplopelma 9.659 HVA Cav ED50 1,650 pmol/g (Migratory manieusis) (Zhang et al., 2003; Deng et al., Hh2a schmidti 2008) IC50 219,000 pmol (cockroach DUM neurons) (Deng et al., 2008) µ-theraphotoxin- Haplopelma 9.270 Na v IC 4300,000 pmol (insect sodium channel para/tipE) (Li et al., 2003) Hhn2b hainanum 50 δ-ctenitoxin-Pn1b Phoneutria 5.210 Insect-specific nigriventer Glutamate LD 3.84 pmol/house fly (M. domestica) (Oliveira et al., 2003) neurotrans- 50 mission γ-ctenitoxin-Pn1a Phoneutria 9.021 NMDA receptor LD 1.03 pmol/house fly (M. domestica) (de Figueiredo et al., 2001) nigriventer 50 ω-hexatoxin-Ar1a robutus 9.010 M-LVA Cav LD50 236 ± 28 pmol/g (A. domestica) (Chong et al., 2007) HVA Cav IC50 692 nM (M-LVA) (Chong et al., 2007) and 644 nM (HVA) (Chong et al., 2007) U1-TRTX-Ba1a Brachypelma 4.412 ND LD 2,450 pmol/g (A. domestica) (Corzo et al., 2009) ruhnaui 50 U1-TRTX-Ba1b Brachypelma 4.446 ND LD 2,069 pmol/g (A. domestica) (Corzo et al., 2009) ruhnaui 50 κ-theraphotoxin- Eurocratoscelus 3.635 Insect-specific IC50 [peak IBK(Ca)] 0.0037 pmol (Gryllus bimaculatus) (Windley et al., 2011) Ec2a constrictus BKCa κ-theraphotoxin- Eurocratoscelus 3.685 Insect-specific IC50 [peak IBK(Ca)] 0.0253 pmol (G. bimaculatus) (Windley et al., 2011) Ec2b constrictus BKCa

Legends Nav: Voltage gated sodium channel Kd: The equilibrium dissociation constant EC50: The half maximal effective concentration, ND: Not determined LD50: The median lethal dose pSlo: Cockroach (Periplaneta americana) Slowpoke (dSlo) calcium-activated PD50: The dose of antiserum or vaccine that protects 50% of the animals challenged potassium channels BKCa : calcium-activated big potassium channel HVA: High voltage activated NMDA receptor: N-methyl-D-aspartate receptor LVA: Low-voltage-activated IC50: The half maximal inhibitory concentration ED50: The half maximal effective dose

13

Table 1.3. Insecticidal peptides isolated from scorpion venoms together with their molecular target and effective dose

Venom Molecular Molecular Source species Effective dose Peptide weight (kDa) target Alpha (α)-anti-insect LqqIII Leiurus quinquestriatus 7.240 NaV LD50 8.29 pmol/g (Blatella germanica), LD50 16.57 pmol/g (Musca quinquestriatus domestica) (Kopeyan et al., 1993) BotIT1 Buthus occitanus 7.343 NaV LD50 81.71 pmol/g (B. germanica) (Borchani et al., 1997) tunetanus BjαIT Buthotus judaicus 9.270 NaV PD50 14.02 pmol/g (Sarcophaga falculata), PD50 53.94 pmol/g (Locusta migratoria) (Arnon et al., 2005) LqhαIT Leiurus quinquestriatus 9.571 NaV LD50 2.51 pmol/g (B. germanica) (Gordon et al., 1996) herbraeus Alpha (α)-Like Bom III Buthus occitanus 6.872 NaV LD50 52.53 pmol/g (B. germanica) (Vargas et al., 1987; Gordon et al., mardochei 1996; Cestele et al., 1999) Bom IV Buthus occitanus 7.296 NaV LD50 19.7 pmol/g (B. germanica) (Vargas et al., 1987; Gordon et al., 1996; mardochei Cestele et al., 1999) Lqh III L. quinquestriatus 7.057 NaV LD50 28 pmol/g (B. germanica) (Krimm et al., 1999) herbraeus Lqh 6 L. quinquestriatus 6.803 NaV LD50 34.3 pmol/g (B. germanica) (Hamon et al., 2002) herbraeus Lqh 7 L. quinquestriatus 6.830 NaV LD50 28.7 pmol/g (B. germanica) (Hamon et al., 2002) herbraeus BmKI Buthus martensii 9.496 NaV CPU 158.41 pmol/g (Gryllus emma), CPU 278.38 pmol/g (Calliphora fly larvae) (Ji et al., 1996; Li and Ji, 2000) BmKII B. martensii 7.226 NaV CPU 345.97 pmol/g (G. emma) CPU 1,110 pmol/g (Calliphora fly larvae) (Ji et al., 1996; Li and Ji, 2000)

14

Table 1.3. Insecticidal peptides isolated from scorpion venoms together with their molecular target and effective dose (continued)

Venom Molecular Molecular Source species Test insect, assays and effective dose Peptide weight (kDa) target Beta (β)-contractive AaHIT1 Androctonus australis 9.852 NaV CPU 0.609 pmol/g (Sarcophaga argyrostoma) (Loret et al., 1990) Bmk IT1 B. martensii Karsch 9.767 NaV ED50 18.43 pmol/g (Gryllus bimaculatus) (Escoubas et al., 2000b) BjxtrIT B. judaicus 10.512 NaV ED50 3.9 pmol/g (Sarcophaga falculata) (Oren et al., 1998; Pelhate et al., 1998; Froy et al., 1999) Beta (β)-depressant LqhIT2 L. quinquestriatus 9.334 NaV LD50 53.57 pmol/g (S. falculata) (Ali et al., 2001) herbraeus LD50 546.39 pmol/g (S. litura) (Corzo et al., 2000) LqqIt2 L. quinquestriatus 9.100 NaV LD50 153.85 pmol/g (S. falculata) (Zlotkin et al., 1985; Kopeyan et al., quinquestriatus 1990; Ali et al., 2001) BotIT2 B. occitanus tunetanus 6.918 NaV LD50 195.14 pmol/g (B. germanica) (Borchani et al., 1996; Cestele et al., 1997) BotIT4 B. occitanus tunetanus 6.845 NaV LD50 160.7 pmol/g (B. germanica) (Borchani et al., 1996; Borchani et al., 1997) BotIT5 B. occitanus tunetanus 6.817 NaV LD50 161.36 pmol/g (B. germanica) (Borchani et al., 1997) BsIT1 Buthus sindicus 6.821 NaV LD50 98.23 pmol/g (S. falculata) and LD50 202.32 pmol/g (B. germanica) (Ali et al., 2001) BsIT2 Buthus sindicus 6.892 NaV LD50 117.53 pmol/g (S. falculata) and LD50 232.15 pmol/g (B. germanica) (Ali et al., 2001) BsIT3 Buthus sindicus 6.715 NaV LD50 153.39 pmol/g (S. falculata) and LD50 242.74 pmol/g (B. germanica) (Ali et al., 2001) BsIT4 Buthus sindicus 6.657 NaV LD50 117.17 pmol/g (S. falculata) and LD50 231.31 pmol/g (B. germanica) (Ali et al., 2001) BaIT2 Buthacus arnicola 6.845 NaV LD50 511.32 pmol/g (B. germanica) (Cestele et al., 1997)

15

Table 1.3. Insecticidal peptides isolated from scorpion venoms together with their molecular target and effective dose (continued)

Venom Molecular Molecular Source species Test insect, assays and effective dose Peptide weight (kDa) target Beta (β)-contractive/depressant (intermediary) TsVII Tityus serrulatus 9.382 NaV CPU 68.22 pmol/g (S. argyrostoma) (De Lima et al., 1986; Lima and Martin-Eauclaire, 1995) TbIt-1 Tityus bahiensis 6.821 NaV LD50 586.42 pmol/g (M. domestica) (Pimenta et al., 2001) Tb2-II T. bahiensis 6.963 NaV LD50 287.23 pmol/g (M. domestica) (Pimenta et al., 2001)

Legends ND: Not determined LD50: The median lethal dose IC50: The half maximal inhibitory concentration ED50: The half maximal effective dose CPU: Contraction paralysis unit PD50: The dose of antiserum or vaccine that protects 50% of the animals challenged Nav: Voltage gated sodium channel

16

1.4.3 Insecticidal peptides from centipede venom

Centipedes (class Chilopoda) are among the oldest extant terrestrial arthropods. They are characterized by the presence of a head and an externally segmented body containing a pair of articulate legs in each segment. The ventral region of the head contains a pair of forcipules (modified legs), whose extremities finish in venom claws. These structures are connected to a short, cylindrical venom gland via a venom duct. There are ~3,300 centipede species worldwide within five extant orders: Scutigeromorpha, Lithobiomorpha, Craterostigmomorpha, Geophilomorpha, and Scolopendromorpha (Undheim and King, 2011). The order Scolopendromorpha contains the largest centipedes. Although centipedes are generalist predators, they appear to eat mainly insects, spiders and other arthropods (Malta et al., 2008).

Centipedes are an ecologically important group of predators that use venom primarily to subdue insect prey. Despite their abundance and frequent, often painful, encounters with humans, little was known about the venom and venom apparatus of centipedes until quite recently (Rates et al., 2007; Undheim and King, 2011). However, there is been much interest in these venomous predators in the past few years. Centipede venoms contain a variety of enzymes, including phosphatases, phospholipases, esterases, glycoside hydrolases, hyaluronidases, metalloproteases, serine proteases etc.), non-enzymatic proteins including pore-forming toxins, protease inhibitors, carditoxins, disintegrins, and mytotoxins, as well as small molecules such as histamine and serotonin (Undheim and King, 2011; Liu et al., 2012; Undheim et al., 2014). Several recent studies have revealed that centipede venoms are also rich in SS-rich neurotoxic peptides, although they differ in sequence and structure from those found in arachnid venoms (Liu et al., 2012; Yang et al., 2012; Undheim et al., 2014).

Centipede venom peptides have the potential to be effective bioinsecticides (Yang et al., 2012). The SS-rich centipede-venom peptides µ-SLPTX-Ssm1a, κ-SLPTX-Ssm1a, κ-SLPTX-Ssm2a and κ-SLPTX-Ssm3a isolated from the venom of Scolopendra subspinipes mutilans are potent insecticides that cause signs of neurotoxicity, including twitching, paralysis, and body contraction, within 10 min to 2 h following injection (Yang et al., 2012). Remarkably, crude venom from a single individual of Scolopendra viridicornis nigra is capable of killing 100,000 houseflies (Rates et al., 2007).

17

Table 1.4 presents a summary of the insecticidal peptides that have been isolated to date from centipede venoms. Historically, insect NaV channels have been the most commonly exploited targets for insecticide development, and this is a common target of spider and scorpion toxins as discussed in Sections 1.4.1 and 1.4.2. Surprisingly, KV channels, which have not been previously exploited for insecticide development, are the most common targets of the few insecticidal centipede-venom peptides that have been characterised. Thus, centipede venoms might prove to be a useful source of insecticidal leads with new modes of action.

Table 1.4. Insecticidal peptides isolated from centipede venoms together with their molecular target and effective dose

Molecular Venom Source Molecular weight Effective dose Peptide species Target (kDa) µ-SLPTX- Scolopendra 3.767 NaV LD50 66.37 pmol/g (adult blowflies) Ssm1a subspinipes and LD50 6290 pmol/g (cockroach) mutilans (Yang et al., 2012) κ-SLPTX- S. sub-spinipes 8.557 KV LD50 8.88 pmol/g (adult blowflies) and Ssm1a mutilans LD50 3.39 pmol/g (mealworms) (Yang et al., 2012) κ-SLPTX- S. sub-spinipes 8.335 KV LD50 2.04 pmol/g (adult blowflies) and Ssm2a mutilans LD50 1.56 pmol/g (cockroach) (Yang et al., 2012) κ-SLPTX- S. sub-spinipes 9.869 KV LD50 4.05 pmol/g (adult blowflies) and Ssm3a mutilans LD50 2.44 pmol/g (mealworms) (Yang et al., 2012)

Legends Nav: Voltage gated sodium channel KV: Voltage gated potassium channel LD50: The median lethal dose

18

1.4.4 Insecticidal peptides from hymenopteran venoms

Hymenopterans (including bees, wasps, and ants) are a highly prevalent and widespread group of insects that evolved more than 300 million years ago (Meinwald and Eisner, 1995). These insects are classified in two groups based on their life history: social and solitary. The great majority of both social and solitary wasps are predators, including many agricultural pests (Schwartz et al., 2012). The venoms of social hymenopterans evolved to be used as defensive tools to protect colonies of these insects from attacks by predators. In contrast, the venom of solitary hymenopterans evolved mainly to induce paralysis of prey in order to permit egg laying on/within the body of the prey; thus, some components of these venoms cause permanent/transient paralysis in prey, while other components seem to prevent infection of the food as well as future progeny.

In general, hymenopteran venoms are composed mainly of proteins, peptides, biogenic amines, and inorganic salts (Nakajima et al., 1986). Among the high molecular weight compounds there are many types of allergens (reviewed in (Schwartz et al., 2012)), such as antigen 5 (Hoffman, 1993; Cascone et al., 1995; Pirpignani et al., 2002), hyaluronidases (Kolarich et al., 2005), and phospholipases (Soldatova et al., 1993; Abe et al., 2000; Costa and Palma, 2000).

The peptide components of hymenopteran venoms have masses ranging from 1.4 to 7 kDa and together they comprise up to 70% of the weight of lyophilised venom (Baptista-Saidemberg et al., 2011; Brigatte et al., 2011). Most of these toxins are linear polycationic amphipathic peptides with a high content of α-helical secondary structure (Palma, 2006). These peptides generally account for cell lysis, hemolysis, antibiosis, and some promote delivery of cellular activators/mediators through interaction with G-protein coupled receptors (Palma, 2006). In addition to these peptides, hymenopteran venoms also contain a neurotoxins that target neuronal ion channels and receptors (Palma, 2006). Ant venoms are less well studied that those of bees and wasps, but two insecticidal ant-venom peptides have been described, namely the ponericins (Orivel et al., 2001) and poneratoxins (Piek et al., 1991).

1.5 Potential of insecticidal arthropod-venom peptides as bioinsecticides

The most potent insecticidal compounds isolated from arthropod venoms are peptides that target ion channels or receptors in the insect nervous system. These neurotoxic peptides are unlikely to be topically active because in order to access their sites of action in the insect nervous system they would have to penetrate the insect exoskeleton, which

19

comprises an outer lipophilic epicuticle and a heavily sclerotized exocuticle (King and Hardy, 2013; Smith et al., 2013). In the only report that describes topical activity for a spider-venom peptide, a fusion of Hv1a to the C terminus of thioredoxin was found topically active to second-instar Helicoverpa armigera and Spodoptera littoralis larvae (Khan et al., 2006) when applied in a solution containing a very high concentration of imidazole. However the contact insecticidal activity of imidazole itself (Pence, 1965), makes it uncertain whether Hv1a is indeed topically active. If the topical route is excluded, then ISVPs must be delivered via a vector such as an entomopathogen or baculovirus or, alternatively, ingested by the targeted insect pests (if they have oral activity) in order to be effective (King and Hardy, 2013; Smith et al., 2013).

Very few studies have explored the oral activity of insecticidal peptides from arthropod venoms, but emerging evidence indicates that some of them, particularly those with an ICK motif and high levels of protease resistance, have a low level of toxicity when fed to insects. Because of their hyper-stability and long residence time in the insect gut, even low rates of intestinal absorption of these peptides would show enhanced toxicity. For example, the ICK-containing insecticidal peptide Hv1a is orally active against the lone star tick, Amblyomma americanum, and its oral potency is only slightly lower compared to when the peptide is injected (Mukherjee et al., 2006). Consistent with this observation, the same peptide (or its ortholog ω-HXTX-Ar1a) was shown to be orally active against lepidopteran pests when expressed in cotton (Gossypium spp.), poplar (Populus spp.), and tobacco (Nicotiana tabacum) plants (Hong et al., 1996; Khan et al., 2006; Cao et al., 2010; Omar and Chatha, 2012).

Low level of oral insecticidal activity is one of the major concerns in the development of new insecticides from insecticidal venom peptides. Most insecticidal venom peptides that showed promising activity in injection-based insect bioassays were almost 100 times less toxic when fed to insects (King and Hardy, 2013) with some exceptions (Mukherjee et al., 2006). The low oral toxicity of insecticidal venom peptides is probably due to their slow rate of absorption in the insect gut as noted previously for disulfide-rich toxins from scorpion and snake venoms (Casartelli et al., 2005). Thus, any approach that improved the rate of venom-peptide absorption in the insect gut would enhance the potential of these insecticidal toxins as candidates for commercial insect pest management (King and Hardy, 2013).

Fusion of venom peptides with a carrier protein to facilitate their transport across the insect gut would be the best possible option. The best-studied fusion protein for this purpose is Galanthus nivalis agglutinin (GNA), a mannose-specific lectin from the

20

snowdrop plant. Lectins are carbohydrate-binding, protease-resistant proteins that are naturally produced in various plant species (Romeis et al., 2003; Sadeghi et al., 2008). Gatehouse (1998) reviewed that, lectins were first identified over a century ago when red blood cells were shown to be agglutinated by an extract of castor bean (Ricinus communis) (Gatehouse, 1998). Lectin compounds have a common characteristic multiple binding sites for free sugars enabling it to cross-link with oligo- or polysaccharides (Gatehouse, 1998). The biological functions of lectins depend on the reversible binding with specific monosaccharides or complex glycans through noncatalytic domains. They play an important role in plant-defence against insect herbivores and consequently a broad spectrum of plant lectins have been tested for insecticidal activity against agriculturally important lepidopteran, coleopteran, dipteran, and hemipteran pests (Vasconcelos and Oliveira, 2004; Michiels et al., 2010; Vandenborre et al., 2011).

Figure 1.3: Structure of GNA tetramer (PDB accession code 1MSA). The four subunits are colour coded (A = green, B = cyan, C = magenta, D = yellow; dimers are formed between subunits A and D, and subunits B and C). One intramolecular disulfide bond (Cys29–Cys52; red tube) is found in each subunit.

Lectins bind to glycoproteins on the insect gut membrane and thus negatively affect multiple physiological processes. In addition, certain plant lectins such as GNA can pass intact into the insect hemolymph following oral delivery (Fitches et al., 2001). GNA is a homotetrameric lectin (50 kDa) which is abundantly produced in bulbs of the snowdrop plant (Hester and Wright, 1996). It is highly stable over a wide range of temperatures and

21

pH (Hester and Wright, 1996), and has insecticidal activity in some insect species (Fitches et al., 2001).

The binding of GNA to aminopeptidase N, a glycoprotein on the insect gut membrane (Fitches et al., 2010), is thought to mediate entry of GNA into the cell by receptor-mediated endocytosis, followed by transcytosis of the endocytosed lectin (Fig. 1.3). The movement of orally delivered GNA from the gut into the hemocoel was confirmed by detection of orally ingested GNA in hemolymph, Malpighian tubules, fat bodies, ovarioles, and the central nerve cord of insects (Fitches et al., 2012). Thus, fusion of insecticidal venom peptides to GNA provides a mechanism for effective oral delivery of these toxins to their site of action, thereby allowing exploitation of venom peptides that show limited insecticidal activity after oral administration.

Figure 1.4: Lectin-mediated delivery of peptide neurotoxins.

When ingested, most neurotoxic peptides have limited insecticidal activity because they are unable to move across the midgut epithelium into the hemocoel to reach their site of action in the insect nervous system. Lectins such as GNA bind receptors in the gut epithelium and translocate into the hemocoel (Fitches et al., 2001). Thus, fusions of insecticidal venom peptides to GNA serve to deliver these toxins to their sites of action.

22

GNA itself has not been used for crop protection due to its weak, species-specific insecticidal activity (Powell et al., 1998). However, the special ability of GNA to transcytose cross the midgut of insects has received considerable attention in past few years. GNA has been used to successfully deliver a number of venom-derived insecticidal peptide toxins, including Hv1a, into insect hemolymph (Fitches et al., 2002; Fitches et al., 2004; Down et al., 2006; Trung et al., 2006; Fitches et al., 2010; Wakefield et al., 2010; Fitches et al., 2012). Moreover, the oral activity of these insecticidal venom peptides was shown to be much higher when they were fused to GNA. A fusion protein comprising ButaIT, a toxin from Indian red scorpion Mesobuthus tamulus, fused to the N-terminus of GNA, showed significant oral activity when fed to larvae of the tomato moth Lacanobia oleracea (Trung et al., 2006). In contrast, GNA and ButalT showed minimal toxicity when they were fed individually (Trung et al., 2006). The ButaIT–GNA fusion is orally active against insects from the orders Lepidoptera, Coleoptera, Hemiptera, and Diptera (Fitches et al., 2010). Delivery of orally ingested ButaIT–GNA into the hemocoel of L. oleracea was demonstrated by immunoblotting (Trung et al., 2006; Fitches et al., 2010). A similar enhancement of oral insecticidal activity was found when SFl1, a neurotoxic peptide from the spider Segestria florentina, was fused to the N-terminus of GNA (Fitches et al., 2004).

Similarly, fusion of Hv1a to GNA results in oral delivery of the toxin to its site of action, the CNS, in Mamestra brassicae (Fitches et al., 2012). GNA-mediated delivery of Hv1a into the hemolymph and CNS was demonstrated by immunoblotting as well as fluorescence microscopy (Fitches et al., 2012). Feeding second instar larvae with cabbage leaf discs coated with 0.2% Hv1a–GNA caused 85% mortality after 10 days (Fitches et al., 2012). Thus, studies with GNA fusions to the venom peptides ButaIT, SFI1, and Hv1a have confirmed the potential of this approach for oral delivery of insecticidal venom peptides.

An alternative strategy that has been successfully used for oral delivery of insecticidal venom peptides is fusion to the coat protein of aphid-vectored viruses. Persistently transmitted plant viruses (i.e., viruses that enter and persist in the hemocoel of the insect vector) are ingested during vector feeding on plant sap; they then move from the gut of the vector into the hemocoel before being transmitted to other plants via the salivary glands (Tamborindeguy et al., 2010) (Fig. 1.5). The ability of the virus to move from the gut into the hemocoel by transcytosis is of particular interest for oral delivery of insect-specific toxins (Bonning and Chougule, 2014; Bonning et al., 2014).

In the Fig. 1.5 showing the circulative route of luteoviruses in aphids, the ingested virus moves up the food canal, through the foregut, and then accumulates in the midgut or hindgut. Virus is then acquired into the hemocoel. Virus may accumulate in the hemocoel

23

and remain viable for weeks. Transmissible virus (white hexagons) is transported into the accessory salivary gland, but does not associate with the principal salivary gland. Transmissible virus is then injected into the plant through the salivary duct when the aphid feeds on a plant.

Figure 1.5: The circulative route of luteoviruses in aphids.

Movement of luteoviruses across the gut of aphid vectors is mediated by clathrin-coated vesicles (Gray and Gildow, 2003), which form tubular transport structures that release virus into the hemocoel. A similar clathrin-coated-vesicle-mediated process occurs for virus movement from the hemocoel across the accessory salivary gland into the duct of the aphid salivary gland. The viral coat protein (CP) mediates the recognition of luteovirus by epithelial receptors in the aphid hindgut. Bonning and co-workers have taken advantage of this property of the coat protein to deliver insecticidal venom peptides into the hemocoel of aphids (Miller and Bonning, 2003; Bonning et al., 2014). For example, they showed that fusion of AaIT, a 70-residue insecticidal scorpion-venom peptide, to an N-terminal portion of the CP from barley yellow dwarf luteovirus substantially enhanced its oral activity against the aphids Myzus persicae and Rhopalosiphum padi (Miller and Bonning, 2003).

More recently, the CP from pea enation mosaic virus (PEMV) was also shown to deliver peptides to the aphid hemocoel. Bonning et al. (2014) observed green fluorescence in the pericardial cells of the pea aphid after feeding aphids with a fusion protein consisting of GFP fused to a portion of the CP of PEMV (Bonning et al., 2014) but not with GFP alone,, indicating that the PEMV CP transported GFP across the aphid gut epithelium. Significant mortality was observed in four species of aphid when they were fed a PEMV CP–Hv1a

24

fusion protein, but not when they were fed Hv1a alone (Bonning et al., 2014).

CP-mediated delivery should be widely applicable to insecticidal venom peptides, but since luteoviruses are specifically vectored by aphids, it is unlikely that luteoviral coat proteins will facilitate transport of these peptides across the gut of insects other than aphids. However, PEMV CP is able to cross the gut epithelium of the aphid Aphis glycines, which is not vector for PEMV (Bonning et al., 2014), and therefore this approach is likely to be widely applicable to aphids and possibly other sap suckers.

A significant advantage of the fusion-protein approach is that fusion-protein transgenes could be used to engineer insect-resistance traits in crop plants. There has been some controversy about expressing carbohydrate-binding lectins in transgenic plants due to their putative anti-nutritional properties (Vasconcelos and Oliveira, 2004); however, no adverse effects were found on rats that consumed transgenic rice expressing GNA for 90 days (Poulsen et al., 2007).

1.6 Genetic transformation techniques of crop plants for insect resistance

Genetic transformation of crop plants involves the integration of genetic material (single or multiple genes) into the recipient plant genome. Genetic engineering offers the potential to transfer characteristics freely into plant species, if the genes that determine those characteristics can be identified and isolated. The ability to genetically transform a plant is useful for studying gene function, producing heterologous proteins, or conferring new properties to the plant that are difficult or impossible to introduce by conventional breeding techniques. The existence of a natural transformation system for plants (the bacterium Agrobacterium tumefaciens) (Schell and Van Montagu, 1977; Horsch et al., 1985; Hooykaas, 1989); the totipotency of plant tissue (a transformed portion of leaf tissue could regenerate a whole plant); and sophisticated techniques for plant transformation and regeneration makes genetic engineering easier for a variety of plants. Plant genetic transformation techniques generally require the construction of a vector (genetic vehicle) that transports the genes of interest and contains essential control sequences, such as a promoter and terminator, as well as a selectable marker.

The method for transferring foreign genes into plants can be categorized as indirect (vector-mediated) or direct (vector-less) gene transfer. Biological methods using bacteria are referred to as indirect, while direct methods are physical and based on penetration of the cell wall. A good comparison of these genetic transformation methods along with a

25

summary of their advantages and limitations is reviewed in Rivera et al. (2012) (Rivera et al., 2012).

Popular techniques for direct gene transfer into plants include microinjection, polyethylene glycol-mediated transfer into protoplasts, electroporation of protoplasts, and particle gun/microprojectile bombardment (biolistics) (Klein et al., 1987). While these techniques can be used for certain transformations, their application is limited. For example, in microprojectile bombardment, DNA-coated gold particles are introduced into target cells via electric discharge particle acceleration or helium gas; the disadvantages are high copy number and rearrangement of the transgene. Also, a tissue-culture stage is necessary with unavoidable risk of somaclonal variation (discussed later).

Indirect transformation methods involve introducing plasmids into the target cell by means of bacteria that are capable of transferring genes to higher plant species (Broothaerts et al., 2005). The most commonly used microorganism is Agrobacterium tumefaciens, a soil bacterium. The size of a plasmid employed for transformation varies between 5 and 12 kilobase pairs (kbp); the plasmid is replicated in the same way as the bacterial chromosome, and replicates autonomously within the host. A single cell may have up to 50 or more plasmids. Agrobacterium is a plant-pathogenic bacterium, capable of transferring a Ti plasmid to its host, and consequently these Ti plasmids are used as vectors to transfer heterologous genes into the plant genome. This technique was introduced in 1977 and it is now widely used (Chilton et al., 1977; Schell and Van Montagu, 1977; Gelvin, 2003). Agrobacterium is known as a “natural genetic engineer” of plants because these bacteria can transfer T-DNA of their plasmids into the plant genome upon infection of cells at a wound site, thereby leading to disorganized growth of cell mass known as crown gall. The tumour-inducing genes are removed (disarmed) in the vectors used for plant transformation. A foreign gene is cloned into the T-DNA region of a Ti plasmid in place of unwanted sequences, and then the Ti-plasmid is used to deliver that gene into target plant cells and tissues. As the transformation success rate is low, transformed plant cells are distinguished from untransformed cells by including a selectable marker gene (e.g. for antibiotic or herbicide resistance) in the Ti plasmid.

Agrobacterium-mediated gene transfer has been widely used with many dicot and monocot crops. To transform plants, leaf discs (dicots) or embryogenic callus (monocots) are collected and infected with Agrobacterium carrying recombinant disarmed Ti-plasmid vector. The infected tissue is then cultured (co-cultivation) on shoot regeneration medium for 2–3 days during which time transfer of T-DNA takes place. After this, the transformed tissues (leaf discs/calli) are transferred onto plant regeneration medium supplemented

26

with a concentration of antibiotic or herbicide that is lethal to non-transformed tissues. After 3–5 weeks, the regenerated shoots (from leaf discs) are transferred to root-inducing medium, and after another 3–4 weeks, complete plants are transferred to soil following the hardening of regenerated plants. PCR of genomic DNA and southern hybridization can be used to detect the presence of foreign genes in the transgenic plants.

There are some complications with Agrobacterium-mediated transformation, including difficulties with transforming some monocots (grasses, etc.) and difficulties in regeneration of plants from tissue culture of leaf discs. Plant transformation by tissue culture is generally time-consuming and requires significant technical skill. Furthermore, there are several variables that must be considered with this method, such as explant availability, identification of a large population of regenerable cells, accessibility of regenerable cells to Agrobacterium inoculation, and appropriate media and hormones that induce shoot and root regeneration. Since the regeneration of a plant from tissue culture relies upon a few transformed cells, the resulting plants will likely have somaclonal variation, the sum of genetic and epigenetic changes in the transgenic plant that was inherited from the parental cells (Larkin and Scowcroft, 1981; Karp, 1995). This problem was solved by in planta Agrobacterium-mediated plant transformation using vacuum infiltration or floral dip, which does not require regeneration of plants from tissue culture of leaf discs (Bechtold et al., 1993; Clough and Bent, 1998; Narusaka et al., 2010). Moreover in planta methods do not require performance by a specialist, and less equipment, labour and reagents are needed to obtain transformed plants. Also, in a given T1 hemizygous transformants, all cells are transgenic. Thus, there is minimal somaclonal variation as compared to that typically encountered with tissue culture (Labra et al., 2004). In planta transformation was first shown with Arabidopsis by imbibing germinating seeds with Agrobacterium (Feldmann and David Marks, 1987). Later, vacuum infiltration was used as a means to increase the likelihood of getting Agrobacterium penetration into whole Arabidopsis plants (Chang et al., 1994; Bechtold and Pelletier, 1998; Ye et al., 1999; Bechtold et al., 2000). Vacuum infiltration methods have been used successfully in transforming pakchoi (Brassica rapa L. ssp. chinensis) (Liu et al., 1998; Qing et al., 2000), alfalfa (Medicago truncatula) (Trieu et al., 2000), Camelina sativa (Lu and Kang, 2008) and Brassica napus (Wang et al., 2003).

The floral dip technique is a recent advance in in planta Agrobacterium-mediated transformation of flowering plants like Arabidopsis (Clough and Bent, 1998; Bent, 2006). This technique involves transformation of plant germ cells, in which inflorescences are dipped into a suspension of Agrobacterium carrying activated vir genes (Clough and Bent,

27

1998). In the typical floral dip method, a vacuum is no longer required for efficient infiltration of Agrobacterium into the plant. However, frequent multiple applications of dipping solution comprising Agrobacterium is detrimental to Arabidopsis, particularly if the dip intervals are less than every fourth day. Arabidopsis, radish (Raphanus sativus L. longipinnatus Bailey) and canola have been successfully transformed by use of a floral dip method (Clough and Bent, 1998; Curtis and Nam, 2001; Curtis, 2005). Note, however, that a single transformation technique is not always suitable for all plant species (Gelvin, 2003); some plant varieties may be successfully transformed with one technique but not another.

1.7 Insect-resistant plants using transgenes encoding insecticidal venom peptides

Despite the global success of Bt crops, which have dominated the market for insect resistant transgenic crops for the last two decades (Tabashnik et al., 2013), there has been significant research on alternative insecticidal genes for generating insect-resistant transgenic crops during this time. In particular, there has been considerable work on the introduction of transgenes encoding insecticidal arachnid toxins into plants. So far, all transgenic plants expressing insecticidal peptides from arachnids have shown significantly increased resistance to insect pests (Table 1.5). For example, Barton and Miller (1993) patented the first insect-resistant transgenic plant expressing an insect-specific toxin from scorpion in 1993 (Barton and Miller, 1993). They showed that transgenic plants expressing the insect-specific peptide toxin AaIT from the venom of the Buthid scorpion Androctonus australis Hector (Zlotkin et al., 2000) is lethal to Heliothis zea upon ingestion

(Barton and Miller, 1993). AaHIT1 is a specific modulator of insect NaV channels (see Table 1.3). Subsequently, Yao and coworkers (1996) used Agrobacterium-mediated gene transfer to generate transgenic tobacco plants expressing AaHIT1 under the control of two linked Cauliflower mosaic virus (CaMV) 35S promoters. Insect bioassays revealed that some transgenic plants with the AaIT gene had notable resistance against insects (Yao et al., 1996). Transgenic poplar plants expressing AaHIT1 were shown to have increased resistance against first instar larvae of Lymantria dispar as evidenced by a decrease in leaf consumption by the larvae, lower larval weight gain, and higher larval mortality (Wu et al., 2000). Transgenic cotton engineered to express AaHIT1 under the control of a CaMV 35S promoter was found have increased resistant to cotton bollworms (larvae of Heliothis armigera) (Wu et al., 2008).

Transgenes encoding insecticidal spider-venom peptides have also been used to engineer insect-resistant plants. Transgenic tobacco expressing the insecticidal peptide ω-HXTX-Ar1a (Ar1a) from the Australian funnel-web spider Atrax robustus were found to

28

have enhanced resistance against cotton bollworms (Hong et al., 1996). This toxin is a specific blocker of insect CaV channels. Transgenic poplar expressing a fusion protein comprised of Ar1a fused to the Bt toxin C-peptide were found to have enhanced resistance against the Asian gypsy moth Lymantria dispar (Cao et al., 2010). Transgenic tobacco plants expressing Hv1a (a paralog of Ar1a, with the same mode of action; see Table 1.2) have enhanced resistance to cotton bollworms (Khan et al., 2006; Shah et al., 2011). The mortality of second-instar H. armigera fed on transgenic tobacco expressing Hv1a was 75–100% after 72 h compared to 0% for larvae fed on untransformed plants, regardless of whether the peptide was expressed under the constitutive CaMV 35S promoter (Khan et al., 2006) or phloem tissue-specific promoters (Shah et al., 2011). It has been mentioned that transgenic cotton expressing Hv1a is as effective as Monsanto’s pyramided Bollgard II® cotton in controlling major cotton pests (Omar and Chatha, 2012).

Huang et al. (2001) transformed two rice varieties with a transgene encoding an insecticidal spider-venom peptide (details of the toxin were not reported in the paper) and observed enhanced resistance against the leaf folder Cnaphalocrasis medinalis and the striped stem borer Chilo suppressalis (Huang et al., 2001). Transgenic tobacco plants expressing Magi 6, a venom peptide produced by the spider Macrothele gigas, were found to be significantly more resistant than wild-type plants to the fall armyworm Spodoptera frugiperda (Hernandez-Campuzano et al., 2009).

Recently, Bonning and co-workers developed transgenic Arabidopsis plants expressing a fusion protein comprised of Hv1a fused to the PEMV CP carrier protein that was described in Section 1.5 (Bonning et al., 2014). These plants had enhanced significantly resistance to the green peach aphid Myzus persicae (Bonning et al., 2014).

Thus, transgenes encoding insecticidal venom peptides seem to hold great potential either as standalone insect-resistant plant traits or for trait stacking with Bt in order to minimise resistance development in targeted insect pests and to expand the range of susceptible insects.

29

Table 1.5. Insect-resistant transgenic plants developed so far with insecticidal venom peptides

Year Toxin Construct Host plant Target insect Insect resistance Reference Insect specific toxin AaIT from the Transgenic plants expressing sufficient (Barton and pTV4AMVST1- 1993 venom of the Buthid scorpion Dicot plant Heliothis zea amounts of AaIT was claimed lethal upon Miller, 1993) 35S-AaIT Androctonus australis Hector ingestion by Heliothis zea. 1996 ω-HXTX-Ar1a from the venom of pET2-35S-ω-HXTX- Tobacco Heliothis First reported the expression of (Hong et al., Australian funnel-web spider Atrax Ar1a armigera insecticidal spider venom peptide gene in 1996) robustus transgenic plant with enhanced resistance against target insect. 1996 AaIT from the venom of the Buthid pNGY-2-35S-TMV- Tobacco Transgenic plants with AaIT gene had (Yao et al., scorpion Androctonus australis AaIT notable resistance against insect. 1996) Hector 2000 AaIT from the venom of the Buthid Poplar Lymantria Decreased leaf consumption, lower (Wu et al., scorpion Androctonus australis dispar weight gain and higher mortality rate of 2000) Hector larvae feeding on transgenic plant. 2001 Spider insecticidal gene SpI pExT-35S-SpI Rice Leaf folder Transformation of rice with the spider (Huang et (Cnaphalocrasis insecticidal gene confer resistance to leaf al., 2001) medinalis) and folder and striped stem borer the striped stem borer (Chilo suppressalis) 2005 The novel insect-resistant gene pBI101-35S-chi- Brassica Diamondback Some of the transgenic plants were high- (Wang et combination, containing an insect- 35S-35S-Bmk-Nos napus cultivar moth (Plutella level expression for both chitinase and al., 2005) specific chitinase gene T maculipenis) scorpion toxin proteins and performed chitinase(chi) and a scorpion insect (DBM) larvae high resistance against the diamondback toxin gene BmkIT(Bmk). moth (Plutella maculipenis) (DBM) larvae infestation. 2006 The fused BGT gene consisting of Poplar P. Lymantria High mortality and significant low larvae (LIN Tong, the insecticidal toxin gene from the simonii×P. dispar weight was found in larvae fed with the 2006) spider, Atrax robustus, and the C nigra transformed poplars indicating negatively terminal of CryⅠA(b)gene from affected growth rate. Bacillus thuringiensis

30

Table 1.5. Insect resistant transgenic plants developed so far with insecticidal venom peptides (continued)

Year Toxin Construct Host plant Target insect Insect resistance Reference 2006 ω-ACTX-Hv1a toxin (Hvt) from the pGreen-35s-35s- Tobacco Helicoverpa Transgenic expression of Hvt in tobacco (Khan et al., venom of the Australian funnel web Hvt-CaMV term. armigera and effectively protected the plants from H. 2006) spider (Hadronyche versuta) Spodoptera armigera and S. littoralis larvae, with littoralis 100% mortality within 48 hour. 2008 Synthetic scorpion Hector Insect pBAaHIT-NPTII- Cotton Cotton Transgenic cotton expressing synthetic (Wu et al., Toxin (AaHIT) gene CaM35S-AaIT- bollworm scorpion Hector Insect Toxin (AaHIT) 2008) NosT (Heliothis gene, was found resistant to cotton armigera) bollworm (Heliothis armigera) killing 44- 98% of the larvae feeding on the leaves. 2009 Magi 6 (a peptide toxin produced Spodoptera Transgenic tobacco plants expressing (Hernandez- by the Macrothele gigas spider) frugiperda were found significantly more resistant Campuzano than the wild type plants against et al., 2009) herbivorous insect. 2010 Fusion gene consisting of the pYHY- ω-ACTX- (Cao et al., spider, Atrax robustus Simon ω- Ar1-C terminal of 2010) ACTX-Ar1 sequence coding for an Cry I A ω-atracotoxin and a sequence coding for the Bt-toxin C-peptide 2011 ω-ACTX-Hv1a toxin (Hvt) from the pGreen0029- Tobacco Heliothis 93-100% mortality of H. armigera larvae (Shah et al., venom of the Australian funnel web RSs1/RolC-Hvt- armigera within 72 h on the leaves of transgenic 2011) spider (Hadronyche versuta) CaMVterminatot plants. 2014 Fusion gene consisting of the coat pBITG-35S-CP-P- Arabidopsis Myzus persicae Suppressed population and reduced (Bonning et protein and a small portion of the Hv1a-Nos thaliana (Col-0 infestation were observed in Myzus al., 2014) read through domain of Pea wild type) persicae feeding on transgenic enation mosaic virus (PEMV) to the Arabidopsis plants expressing CP-P- highly insect-specific, spider- Hv1a. The dead insects that died after derived, peptide ω-hexatoxin-Hv1a feeding on transgenic plants showed the (Hv1a) sign of paralysis.

31

1.8 Minimising non-target effects via tissue-specific expression of insecticidal transgenes

Beneficial insects include natural predators such as wasps and beetles that prey on pest insects, and pollinators such as bees. These beneficial insects might be unintentionally subjected to the effect of insecticidal toxins expressed by transgenic plants. Confining expression of the insecticidal toxin to specific plant tissues via the use of tissue-specific promoters is one way of potentially avoiding detrimental effects on beneficial insects.

In genetic transformation of any organism, including plants, promoters determine the level of transcription and the tissues in which transcription will take place (Juergensen et al., 2003). Constitutive promoters that target gene expression throughout the plant are commonly used in plant transformation studies (Dutt et al., 2012). These promoters can be obtained from numerous sources such as viruses (e.g., the CaMV35S (Odell et al., 1985) and figwort mosaic virus (FMV) promoters (Maiti et al., 1997)), bacteria (e.g., the Agrobacterium tumefaciens Ti plasmid mannopine synthetase (mas) (DiRita and Gelvin, 1987) and nopaline synthase (NOS) promoters (Bevan et al., 1983)), or plants (e.g., the Arabidopsis thaliana ACT2 promoter (An et al., 1996) and the Medicago truncatula MtHP promoter (Xiao et al., 2005)).

The promoter that has been most widely used in transgenic plants is the CaMV35S promoter (Sunilkumar et al., 2002; Potenza et al., 2004), which has the advantage of driving constitutive expression in most tissues in most plant species, including monocots and dicots (Wilkinson et al., 1997; de Mesa et al., 2004). This promoter is derived from a double-stranded DNA viral genome, but it is able to use the nuclear RNA polymerase of host cells to initiate transcription and it is not dependent on any trans-acting viral gene products (Potenza et al., 2004). The CaMV35S promoter has been used to drive Bt toxin expression in rice, potato, sugar beet, and soybean crops (Wilkinson et al., 1997). Expression of l Bt toxin in all plant tissues raises the potential of harmful effects on non-target species. Moreover, in some cases, strong constitutive expression of a foreign gene can be harmful to the host plant, causing sterility, retarded development, abnormal morphology, yield penalty, and/or altered grain composition (Cai et al., 2007).

Constitutive expression of a transgene may not be necessary if expression in a particular tissue is sufficient to obtain desired result (Dutt et al., 2012). For example, many of the most important crop pests are sap-sucking hemipterans such as aphids that suck sap from the phloem tissue. Targeting expression of insecticidal transgenes to the phloem using phloem-specific promoters might be a good option for developing transgenic plants

32

that are resistant specifically to sap-sucking insects. Limiting expression of the insecticidal transgene to the phloem would minimise the potential for unwanted side effects on beneficial insects such as pollinators.

Several phloem-specific promoters have been described. Some are derived from plant pathogens, such as the Agrobacterium rhizogenes rolC promoter (Schmulling et al., 1989), which has been used to express Hv1a toxin in tobacco plants (Shah et al., 2011) and an insecticidal Allium sativum leaf agglutinin gene in both monocot and dicots (Saha et al., 2007). A large number of phloem-specific plant promoters are associated with the sucrose synthase protein, which is localized in phloem cells (Nolte and Koch, 1993) and its expression has been closely linked with vascular bundles (Hawker and Hatch, 1965). The sucrose synthase-1 promoters from rice (RSs1) (Wang et al., 1992) and maize (Yang and Russell, 1990) have been shown to be active in heterologous systems (Yang and Russell, 1990; Shi et al., 1994) and they were used to express the insecticidal Allium sativum leaf agglutinin gene in both monocot and dicot plants (Saha et al., 2007).

Figure 1.6: Distribution of fluorescence in source leaves when GFP was expressed under the control of CaMV 35S or SUC2 promoters. (A) GFP was expressed throughout the leaves of 35S:GFP plants. When transgene expression was directed by SUC2, both free GFP (B) and tmGFP (GFP fused to a transmembrane tether) (C) were found only in the vascular system of the leaves. (D) No GFP fluorescence was seen in leaves of wild-type plants. Scale bar = 0.85 µm/px. Photograph adapted from (Pham, 2010).

The AtSUC2 promoter that drives expression of the sucrose-H+ symporter in Arabidopsis (Sauer and Stolz, 1994) targets gene expression to the phloem in Arabidopsis (Fig. 1.2), tobacco and strawberry (Truernit and Sauer, 1995; Imlau et al., 1999; Zhao et al., 2004). The sucrose-H+ symporter catalyzes uptake of sucrose into phloem (Imlau et al., 1999).

33

Sucrose is a main photosynthetic product synthesized in source tissues that is used for long-distance carbon transport in the phloem system (Truernit and Sauer, 1995). It has been reported that sucrose transporters locate to the phloem and specifically to the companion cells (Potenza et al., 2004).

Plant-based AtSUC2 and RSs1 promoters are weaker in directing vascular-specific gene expression in vegetative parts of the plant than the bacterial rolC promoter (Dutt et al., 2012). However, a weaker plant-based promoter that is able to drive adequate levels of gene expression may be preferable than stronger non-plant-based promoters.

1.9 Scope for controlling major insect pests using insecticidal venom peptides

Control of insect pests is one of the major challenges in modern agriculture as conventional chemical approaches are becoming insufficient to combat key pests. Genetically modified plants engineered to express insecticidal transgenes provide an alternative approach for controlling insect pests, and their utility has been demonstrated via successful adoption of Bt crops in major crop-growing countries. Although Bt toxin has defended crops in the field for over a decade, the emergence of resistance in previously susceptible pest species has spurred research into alternative insect-toxin transgenes.

A large number of insecticidal peptides have been isolated from venom of arthropod predators such as spiders, scorpions and centipedes. Many of these peptides have desirable properties for development as bioinsecticides, including high potency, rapid speed of kill, lack of vertebrate toxicity, low production costs, and activity against a wide range of insect pests. Some of these peptides have novel modes of action compared with extant chemical insecticides, and consequently they might be particularly useful for control of insect pests that have developed resistance to chemical insecticides. Research over the last 20 years has demonstrated that it is possible to develop insect-resistant plants by engineering them to express insecticidal venom peptides. Significant developments have also occurred in approaches to enhance the oral activity of venom peptides. In this thesis I aim to bring these two technologies together by developing transgenic plants that express an insecticidal spider-venom peptide fused to a carrier protein that enhances its oral activity.

More specifically, the overriding aim of this project was to explore whether Arabidopsis plants engineered to express the spider-venom peptide Hv1a, either alone or fused to a carrier protein that improves its oral activity, have enhanced resistance to the highly insecticide-resistant lepidopteran pest Helicoverpa armigera. The specific objectives of my

34

project were as follows:

Aim 1: Produce recombinant Hv1a, GNA, Hv1a-GNA and GNA-Hv1a in a yeast expression system and assess their insecticidal activity.

Aim 2: Construct transformation vectors for Agrobacterium-mediated transformation of Arabidopsis with transgenes encoding Hv1a or fusions of Hv1a with either GNA or PEMV CP. I aimed to create two sets of vectors, one with transgene expression under CaMV35S control and another under AtSUC2 promoter control.

Aim 3: Generate transgenic Arabidopsis plants expressing Hv1a and fusions of this toxin to either GNA or CP, with transgene expression under CaMV35S or AtSUC2 promoter control.

Aim 4: Determine whether in planta expression of Hv1a, GNA, Hv1a-GNA or GNA-Hv1a in Arabidopsis confers resistance against cotton bollworms (larvae of Helicoverpa armigera). The plan was to first characterise plants with transgene expression under CaMV35S control, and only examine plants with transgene expression under AtSUC2 promoter control if time permitted.

35 Chapter 2: Insecticidal activity of recombinant Hv1a fusion proteins

2.1 Introduction

The ongoing search for new and safe insecticidal compounds has led to the discovery of promising insect-specific neurotoxins derived from the venoms of predatory animals that include spiders, scorpions, centipedes, and wasps. The venoms from predators that kill or paralyse insect prey are an excellent natural source of insect-specific neurotoxins (Windley et al., 2012; King and Hardy, 2013; Smith et al., 2013). These insecticidal neurotoxins typically target sodium, potassium, calcium, or chloride channels (Windley et al., 2012; King and Hardy, 2013; Smith et al., 2013). However most insect specific neurotoxins are not orally active and require an appropriate delivery system to access the nerves they target (Fitches et al., 2004; Fitches et al., 2012; Bonning and Chougule, 2014).

Spiders have complex venoms containing hundreds of peptides that are mostly insecticidal (Pelhate and Zlotkin, 1982; Windley et al., 2012; King and Hardy, 2013; Smith et al., 2013). Based on the extant number of spider species (~45,000; see http://research.amnh.org/iz/spiders/catalog/counts.html), it has been estimated that spider venoms contain at least 10 million bioactive peptides (King et al., 2008). There are 800 spider-venom peptides currently included in the ArachnoServer 2.0 Database (www.arachnoserver.org). Of these, 136 have been shown to be insecticidal: 38 are insect-selective, 34 are non-selective, and 64 have unknown phyletic selectivity (Windley et al., 2012).

Although Hv1a and other insecticidal peptides present in spider venoms are toxic to invertebrate insect pests when injected (Mukherjee et al., 2006), the same peptides are typically ineffective when delivered orally or applied topically due to their inability to access the insect hemocoel (Fitches et al., 2012; King and Hardy, 2013). The target specificity of these naturally occurring arthropod-derived insecticidal peptides makes them particularly appealing for insect pest management technologies if the need for injection can be circumvented.

Orally ingested insecticidal peptide neurotoxins must permeate several barriers in the insect digestive system, including the peritrophic membrane and mid-gut epithelium, in order to reach their neuronal targets (Fiandra et al., 2009). The efficacy of the orally ingested toxin cannot be easily predicted because of the limited information available on how the insect gut absorbs these peptides (Fiandra et al., 2009).

36 Fusion protein approaches can enable oral delivery of peptides to their targets by exploiting “carrier” proteins that bind to receptors on the invertebrate gut epithelium before being translocated into the hemocoel. The fusion of an insect toxin to a carrier protein creates an environmentally benign bioinsecticide whose properties can be selected and modified. The construction of a fusion protein involves the linking of two proteins or protein domains by a linker sequence that helps the proteins fold independently and therefore behave as might be expected. The selection of the linker sequence is important, with several studies concluding that the flexibility and hydrophilicity of the linker is key to domain functionality (Argos, 1990; Alfthan et al., 1995; Robinson and Sauer, 1998; Arai et al., 2001). The functionality of the protein on either the N- or C-terminus, the relative orientation of the individual proteins, the linker length and sequence, the need for specific oligomerisation, and the introduction of mutations or truncations that might enhance or eliminate certain features are all important considerations in fusion protein design (Schmidt, 2013).

Fusing a venom peptide such as Hv1a with a protein that facilitates transport across the insect gut is a promising option to increase the oral activity of insecticidal venom peptides. In this respect, the best-studied carrier protein is Galanthus nivalis agglutinin (GNA), a mannose-specific lectin from the snowdrop plant (Fitches et al., 2012). In order to determine whether fusion of Hv1a with GNA increases its oral activity, we produced recombinant Hv1a-GNA and GNA-Hv1a fusion proteins using the yeast Pichia pastoris and compared the oral activity of the insecticidal fusion proteins with Hv1a.

P. pastoris expression system

P. pastoris belongs to one of the four methylotrophic yeast genera (the others are Candida, Hansenula, and Torulopsis) that can metabolize methanol as their sole carbon source (Higgins, 2001). P. pastoris was developed in the early 1970s as a biological tool to convert methanol into high quality protein that can be used as a component in livestock feed (Higgins, 2001). However, this application was never employed widely due to the oil crisis that began in the mid-1970s and the development of soybeans as a cheap and high-quality livestock feed component (Higgins, 2001). In the early 1980s, P. pastoris was developed as a eukaryotic host for the production of heterologous proteins (Higgins, 2001). Philips Petroleum Company and Research Corporation Technologies agreed to release the P. pastoris expression system for academic research laboratories in 1993 (Cregg et al., 2000). As a result, P. pastoris is now widely being used in research and industrial settings.

37 P. pastoris is as easy to manipulate as Escherichia coli (Cregg et al., 1985). However, in contrast to E. coli, P. pastoris has the advantage of being a eukaryotic expression system, which can be critical for allowing proper processing, folding, and post-translational modification of eukaryotic proteins which are often not completely processed in E. coli, resulting in non-functional proteins (Cregg et al., 1985). When compared to other eukaryotic expression systems, such as insect or mammalian cell cultures, P. pastoris expression is faster, easier, cheaper, and allows high levels of expression of the target protein (Daly and Hearn, 2005). Furthermore, spurious glycosylation is limited in P. pastoris compared to the model yeast Saccharomyces cerevisiae, where proteins can be excessively glycosylated (Higgins, 2001).

During methanol metabolism, P. pastoris produces the enzyme alcohol oxidase (AOX), which catalyses the first step in the methanol utilisation pathway (Higgins, 2001). Oxidation of methanol leads to the formation of formaldehyde and hydrogen peroxide, which is toxic to yeast cells (Higgins, 2001). To avoid the toxic effects of hydrogen peroxide, methanol oxidation takes place in specialised organelles called peroxisomes (Cregg et al., 2000).

There are two genes encoding for AOX in P. pastoris, AOX1 and AOX2 (Cregg et al., 1989). The two proteins encoded by these genes have nearly identical amino acid sequences (Koutz et al., 1989), but there is more sequence variation in the promoter region of the genes (Ellis et al., 1985). The AOX1 gene is responsible for the vast majority of alcohol oxidase activity in yeast cells (Cregg et al., 1989). P. pastoris can produce AOX1 as ≥ 30% of the total soluble protein when grown on media with methanol as the main carbon source (Higgins, 2001).

Regulation of the AOX1 gene involves a two-step mechanism: repression/depression and induction, which is very similar to regulation of the GAL1 gene in S. cerevisiae (Cregg et al., 2000). However, in S. cerevisiae, the repression/depression mechanism itself is enough to activate GAL1 (Cregg et al., 2000; Higgins, 2001). In P. pastoris, the AOX1 promoter is repressed by glucose while methanol, which acts as an inducer (the second mechanism), is always required to initiate production of AOX1 (Cregg et al., 2000). The AOX1 promoter might not be suitable for heterologous protein production in some circumstances. In that case, other promoters can be considered such as GAP (derived from P. pastoris glyceraldehyde-3-phosphate dehydrogenase) (Waterham et al., 1997), FLD1 (derived from the P. pastoris FLD1 gene) (Shen et al., 1998), PEX8 (a gene encoding a peroxisomal matrix protein which is essential for peroxisome biogenesis) (Klaas Nico et al., 2001), and YPT7 (a gene encoding a GTPase involved in secretion)

38 (Sears et al., 1998). The GAP promoter is more effective for producing isotope-labelled proteins for NMR studies (Li et al., 2007). The FLD1 promoter offers the flexibility of high-level protein expression induced with methylamine, a cheap nitrogen source (Li et al., 2007). The PEX8 and YPT1 promoters are often used when moderate levels of expression are required (Li et al., 2007).

There are three types of P. pastoris host strains available: Mut+ (methanol utilisation plus), Muts (methanol utilisation slow), and Mut– (methanol utilisation minus). Mut+ is the wild- type strain, which is commonly used for protein production (Cregg et al., 2000). One of the most commonly used strains is X33, which depends on methanol as the main carbon source (Cregg et al., 2000); it is the strain used in the current study. Muts and Mut– are two modified strains where one or two AOX genes have been deleted from the genome (Cregg et al., 2000). These two strains require less methanol and they are sometimes better for protein production than Mut+, especially when a large amount of methanol in the fermentor is considered a fire hazard (Higgins, 2001). KM71 is an example of a host strain that relies on the weaker AOX2 promoter due to replacement of chromosomal AOX1 with the S. cerevisiae ARG4 gene (Cregg et al., 2000).

In some cases, the foreign protein expressed in P. pastoris is rapidly degraded by proteases in the culture medium (Li et al., 2007). Major vacuolar proteases seem to be a significant cause of this degradation, particularly in fermentor cultures where a high cell density environment combined with lysis of a small percentage of cells can activate the proteases (Li et al., 2007). To avoid this problem, the use of host strains that are defective in proteases can be helpful. SMD1163 (his4 pep4 prb1), SMD1165 (his4 prb1), and SMD1168 (his4 pep4) are protease-deficient strains that may provide a more suitable environment for unstable heterologous proteins (Li et al., 2007).

A wide range of heterologous proteins have been successfully produced in P. pastoris via the intracellular or secreted routes (Higgins, 2001). Secreted protein production in P. pastoris requires a signal sequence on the expressed protein, such as the native secretion signal of the protein, the secretion signal sequence from S. cerevisiae α factor prepropeptide (MF-α1), or the P. pastoris acid phosphatase signal (PHO1) (Li et al., 2007). The signal sequence directs the target protein to the secretory pathway (Cregg et al., 2009). As P. pastoris naturally secretes a very low amount of native proteins, the culture media is enriched with the target protein upon induction. This is another contributing factor to why the P. pastoris expression system has become major alternative to other eukaryotic expression systems for the production of heterologous proteins.

39 The P. pastoris expression system was used to produce fusion proteins of Hv1a with GNA. The insecticidal activity of the recombinant fusion proteins was then compared with unfused Hv1a in both injection and oral feeding assays.

2.2 Materials and methods

2.2.1 Designing synthetic gene constructs for Hv1a, GNA, and GNA-Hv1a fusion proteins

Synthetic genes, with codons optimised for P. pastoris expression, were commercially synthesised by GeneArt based on the sequences shown in Fig. 2.1. In this figure, the sequences of Hv1a (red), GNA (green), His6 tag (brown), and TEV protease cleavage site

(ENLYFQS; blue) are indicated in different colours. A His6 tag was added at the N-terminus of each sequence to facilitate purification of the recombinant protein using nickel affinity chromatography. Hv1a was fused to either the N- or C-terminus of GNA via a triple-alanine linker (light blue letters). The synthetic genes were cloned into the pPICZαA expression vector from GeneArt.

Figure 2.1: Sequences of Hv1a, GNA, and fusions thereof that were used to design synthetic genes for expression in P. pastoris. In the sequences of His6-TEV-Hv1a (A) and His6-TEV-Hv1a-GNA (C), the final amino acid residue of the TEV recognition site (Ser) is the first residue of Hv1a. In the case of the fusions of Hv1a with GNA, a triple-alanine linker (i.e., Ala-Ala-Ala; shown in light blue) was added between the Hv1a and GNA.

40 Fig. 2.2 shows the major features of pPICZαA (3.6 kb), the expression vector in this study. pPICZαA uses the AOX1 gene promoter to drive expression of the gene of interest and the α-factor signal sequence for secretion of the target protein into the culture media. A zeocin resistance gene is also integrated into the plasmid for use as a selection marker in both E. coli and P. pastoris (Invitrogen, 2010).

Figure 2.2: pPICZα Expression vector for P. pastoris expression system (Invitrogen, 2010). The α-factor signal sequence is used to target the expressed protein to the culture media. A zeocin resistance gene is included as a selection marker. Genes encoding heterologous proteins are cloned into the multiple cloning site (shown in expansion at top) that follows immediately after the α-factor signal sequence. (Figure taken from user manual for EasySelect Pichia Expression Kit; Invitrogen, 2010).

The E. coli strain TOP10 was used in this study to propagate DNA. Bacterial transformation was carried out using the heat shock method. Briefly, 100 ng of plasmid containing the gene of interest was incubated with 0.1 mL of competent TOP10 cells on ice for 15–30 min. Cells were then heat shocked at 42oC for exactly 60 s, then tubes were kept on ice for 3–5 min. About 900 µL of low-salt Luria-Bertani media (low-salt LB, composed of tryptone 10 g/L, sodium chloride 5 g/L and yeast extract 5 g/L, pH 7.5) was added to the heat-shocked bacterial cells, which were then incubated at 37°C for 60–90 min. After incubation, cells were centrifuged at 5000 rcf then the cell pellet was re- suspended in 100 µL of low-salt LB and plated on low-salt LB agar plates (composed of tryptone 10 g/L, sodium chloride 5 g/L and yeast extract 5 g/L, pH 7.5, agar 15 g/L) containing the antibiotic zeocin (25 µg/mL). The low-salt LB agar plates were then

41 incubated at 37°C overnight. As a negative control, low-salt LB agar plates were inoculated with untransformed E. coli without adding plasmid (no cells are expected to grow on the negative control plates). Five colonies were cultured in 5 mL low-salt LB media (containing 25 µg/mL of Zeocin) at 37°C and glycerol stocks were made from the overnight cultures.

2.2.2 Transformation of P. pastoris expression host strain X-33

About 25–50 mL of low-salt LB containing zeocin (25 µg/mL) was inoculated from a glycerol stock from the bacterial transformation and cultured at 37°C overnight. Miniprep and midiprep procedures (Invitrogen) were used to extract plasmid from E. coli cells. After plasmid extraction, the concentration was measured using NanoDrop 2000 spectrophotometer (Thermo Scientific). About 5 µg of plasmid was then linearised by overnight incubation with SacI-HF restriction enzyme (New England Biolabs) at 37°C.

The linearised plasmid was then transformed into competent P. pastoris (X-33, Mut+) cells using the Pichia EasySelect kit (Invitrogen, 2010). The transformed X-33 cells were then plated on Yeast Peptone Dextrose (YPD) agar plates (1% yeast extract, 2% peptone, 2% dextrose and 20 g/L of agar) containing zeocin (100 µg/ml), and plates were incubated at 30°C for 3–4 days until colonies appeared on the plate. About 10 conspicuous colonies were picked randomly from the YPD agar plate and cultured in YPD media (containing 1% yeast extract, 2% peptone, 2% dextrose and 100 µg/mL zeocin) for 24–36 h. Glycerol stocks were made from these cultures and stored at –80°C until later use for protein production.

2.2.3 Small-scale protein production and detection of clones

The X-33 glycerol stocks (10 colonies) were used for test protein expressions. Briefly, each colony was inoculated in 10 mL YPD medium and cultured at 30°C, with shaking at

220–250 rpm, for 1–2 days (OD600 = 6). Cells were then harvested by centrifugation at 5,000 g for 10 min at room temperature. The supernatant was discarded and the cell pellet was re-suspended in 10 mL of Yeast Nitrogen Base (YNB) media (yeast nitrogen base 3.4 g/L, ammonium sulfate 10 g/L, potassium phosphate buffer 100 mM with pH 8.0, 0.02% biotin). Methanol was then added to a final concentration of either 0.5% or 1% every 24 h to maintain the induction.

For the time-course experiment, a single colony was used to inoculate 50 ml of YPD in a 250 mL flask. The culture conditions for both the growth and induction phase were maintained as described above. During the induction period, 1 mL of culture media was

42 collected every 12 h. The samples were then centrifuged at 8000 rcf for 5 min, and the supernatant was collected (Note: the cell pellet was kept to ensure that the target protein was secreted). The protein expression levels were examined by SDS-Polyacrylamide Gel Electrophoresis (SDS-PAGE) and western blot analysis.

2.2.4 Large-scale protein production

The best clone from the small-scale expression was selected for large-scale expression (1–2 L). About 50 µL of glycerol stock was added to 50 mL of YPD medium and cultured overnight at 30°C with shaking at 220–250 rpm in a rotating incubator. The overnight 50 mL culture was then transferred into 1 L of YPD media and cultured for a further 48 h under the same conditions. The cells were then collected by centrifugation at 5,000 g for 10 min at room temperature. The supernatant was discarded, and the cell pellet was re-suspended in YNB media. Methanol was added to a final concentration of 1% at 24-h intervals for 2 days. The culture supernatant was collected by centrifugation at 8000 rcf for 20 min and used for protein purification. The cell pellet obtained after the induction phase was used for a second round of protein production using fresh YNB medium.

2.2.5 Ni-NTA affinity purification

Protein constructs were designed with an N-terminal His6 tag so they could be purified using Ni-NTA (Nickel- nitrilotriacetic acid) affinity chromatography. For recharging the Ni-NTA column, the NTA resin was first washed with 50 mM EDTA containing 0.1% SDS (pH 8) to get remove residual bound proteins. It was then washed with distilled water (5-10 column volumes) to remove the EDTA. The column was then treated with 5 resin volumes of 100 mM nickel sulfate solution to recharge the beads (after all the nickel sulfate solution is discarded from the column, the beads should be green). Excess nickel sulfate was then removed by washing with water followed by equilibration buffer (20 mM Tris, 150 mM sodium chloride, pH 8). The column was then ready for purification of His-tagged proteins.

The yeast culture supernatant obtained after induction was dialysed overnight against distilled water and then equilibration buffer (pH 8.0) (about 6 h) at 4°C. The dialysed supernatant was applied to the Ni-NTA column and allowed to pass through by gravity flow. Then column was washed with 50 ml of washing buffer (pH 8.0) to remove residual bound proteins.

After the brief wash, the remaining bound proteins were eluted twice with 15 mL of elution buffer (20 mM Tris, 150 mM sodium chloride and 300 mM imidazole, pH 8.0). The 30 mL

43 eluate was dialysed overnight against equilibration buffer at 4°C using a membrane with a molecular weight cut-off of 1 kDa (Spectra/Por® 6 Dialysis Tubing, Spectrum Laboratories). The dialysed protein solution was frozen in liquid nitrogen or on dry ice and then the sample was lyophilised (alternatively, the protein was concentrated or buffer exchanged using a Millipore Amicon Ultra-15 concentrator). The final product from lyophilisation or concentration was then further purified using reverse phase high performance liquid chromatography (RP-HPLC). About 2 mL of sample was collected during all the purification steps for SDS-PAGE or western blot detection.

2.2.6 RP-HPLC purification and mass analysis

Two different reverse phase (RP) columns were used for HPLC; a C18 analytical column (250 mm x 4.6 mm, particle size 5 µm, Vydac) was used for purification of Hv1a and a C4 analytical column (150 mm x 4.6 mm, particle size 5 µm, Supelco Analytical) was used for the purification of GNA, Hv1a-GNA, and GNA-Hv1a. The solvents used for RP-HPLC were solvent A (0.09% trifluoroacetic acid in double distilled water) and solvent B (0.09% trifluoroacetic acid in 90% acetonitrile).

Lyophilised protein samples were dissolved in either distilled water or appropriate buffer and then solvent B was added to a final concentration of 5% (as the RP-HPLC column was equilibrated with 5% solvent B). The acidified samples were centrifuged at 12,500 rpm using bench-top centrifuge and then filtered through 0.2 µM filters. Proteins were purified using a 20%–60% gradient of solvent B over 40 min. Samples were manually collected as single fractions and lyophilised prior to further analysis.

Protein molecular masses were determined via electrospray ionisation mass spectrometry (ESI-MS) using an Applied Biosystem API 2000 system or via matrix-assisted laser desorption ionisation (MALDI) mass spectrometry using an AB SCIEX MALDI TOF/TOF 4700 mass spectrometer. SDS-PAGE and western blot analyses were also conducted for identification of GNA and GNA-Hv1a fusion proteins.

2.2.7 SDS-PAGE and western blotting of the purified fusion proteins

SDS-PAGE is routinely used to separate protein samples according to their electrophoretic mobility and thereby identify their molecular mass. SDS-PAGE gels were prepared in a hand-casted apparatus (1.5 mm thick; BIO-RAD). Table 2.1 shows the recipe for preparation of 16.5% polyacrylamide gels.

44 Table 2.1: Recipe for separating and stacking gels for 16.5% polyacrylamide gel.

Materials Required for One Gel 8 mL of 16.5% Separating Gel 5 mL of 6% Stacking Gel 2.5 mL Water 2.9 mL Water 3.3 mL 40% acrylamide 0.75 mL 40% acrylamide 2 mL 1.5 M Tris pH 8.8 1.25 mL 1.5 M Tris pH 8.8 80 µL 10% SDS 50 µL 10% SDS 80 µL 10% ammonium persulfate 50 µL 10% ammonium persulfate 8 µL tetramethylethylenediamine 5 µL tetramethylethylenediamine

Samples collected during various steps of protein purification were analysed using SDS-PAGE. About 50 µL of each sample was mixed with 50 µL of 3× loading dye (2% SDS, 0.1% bromophenol blue, 10% glycine) and samples were boiled at 95°C for 10 min. A portion (20 µL) of each sample was then electrophoresed on a 16.5% SDS-PAGE gel.

Gels were electophoresed in BIO-RAD Mini-PROTEAN® Tetra cell gel electrophoresis apparatus (vertical mini gel electrophoresis) with tris-glycine-SDS electrophoresis buffer (25 mM Tris, 192 mM glycine, 0.1% SDS, pH 8.3). Precision Plus Protein Dual Color standards were used as molecular mass markers. Gels were electrophoresed at 90 V for 20–30 min, and then the voltage was increased to 120 V for another 45 min. The gel was then either stained with Coomassie brilliant blue staining solution (40% methanol, 10% acetic acid, 0.1% coomassie brilliant blue) for 2–3 h at a room temperature or used immediately for western blot analysis. Stained gels were then immersed in de-staining solution (20% methanol, 10% acetic acid, 70% double distilled water) for 2-3 h or until proteins bands were clearly visible. Gel images were recorded using a BIO-RAD Gel-Doc® system.

Proteins were separated under reducing conditions on an SDS-PAGE gel then transferred to Hybond ECL nitrocellulose membrane (Amersham) by electroblotting at 400 mA for 1 h. The membrane was washed with phosphate buffer saline (PBS, 3.2 mM disodium hydrogen phosphate, 0.5 mM potassium dihydrogen phosphate, 1.3 mM potassium chloride and 135 mM sodium chloride, pH 7.4) and blocked with blocking buffer (Odyssey® Western Blocking Buffer) for 1 h at a room temperature. The membranes were incubated with anti-His antibody (His-Probe H-15 rabbit polyclonal IgG, Santa Cruz Biotechnology, CA, USA) overnight at 4°C on shaker. A rabbit polyclonal anti-GNA antibody (from Dr Elaine Fitches, Food & Environment Research Agency, Sand Hutton, UK) was used to detect GNA and GNA fusion proteins. In all the cases, the primary antibody was diluted to 1:2000. After 4-5 washes with PBST (3.2 mM disodium hydrogen phosphate, 0.5 mM potassium dihydrogen phosphate, 1.3 mM potassium chloride,

45 135 mM sodium chloride and 0.05% tween 20, pH 7.4), the membrane was incubated with secondary antibody (IR Dye 800CW goat polyclonal anti-rabbit IgG, LICOR Biosciences, NE, USA) for 1 h at room temperature (1:10,000 dilution). The membrane was then washed 4-5 times with PBST followed by a final wash with PBS to remove the detergent. The blot was finally analysed by scanning the membrane in Odyssey® Infrared Imaging System at 800 nm for fluorescence from the bound secondary antibody.

2.2.8 Injection bioassay to determine the insecticidal properties of recombinant fusion proteins

The insecticidal activity of the purified recombinant proteins, Hv1a, GNA-Hv1a and Hv1a- GNA, was determined by injecting them into houseflies (Musca domestica) with the help of Dr Volker Herzig (Institute for Molecular Bioscience, the University of Queensland, Australia). The recombinant proteins were injected into the ventro-lateral thoracic region of the houseflies using a 1.0 mL Terumo Insulin syringe (B-D Ultra-Fine, Terumo Medical Corporation, Maryland, USA) with a fixed 29 G needle fitted to an Arnold hand micro-applicator (Burkard Manufacturing Co. Ltd., England). After injection, the flies were individually housed in 2-mL tubes and observed over the first 30 min, and then at 1 h and 24 h after injection.

2.2.9 Feeding assay for oral activity of recombinant fusion proteins

The oral activity of the recombinant fusion proteins was determined by feeding second/third instar diamondback moth (Plutella xyllostella) larva on Arabidopsis leaves coated with purified recombinant Hv1a 0.633 µg/cm2 leaf, recombinant GNA 1.90 µg/cm2 leaf, recombinant Hv1a-GNA 2.532 µg/cm2 leaf and recombinant GNA-Hv1a 2.532 µg/cm2 leaf. Test leaves were prepared by adding droplets of protein (re-suspended in distilled water to make total 5 µL) onto the upper and lower surfaces of leaves. Control leaves were prepared with distilled water. Larvae were reared from hatch to second/third instars on non-treated Arabidopsis and then a single larvae was released into a ventilated plastic dish containing coated leaf and 2% agar gel at the bottom to prevent desiccation. The insect feeding behaviour and the severity of leaf damage was observed after 2 days of insect release. Magnified images were taken to determine the damage caused by the insect feeding on the recombinant toxin-coated and control leaves under microscope and quantitatively analysed using ImageJ programme (Rasband, 1997—2014) and one-way ANOVA using GraphPad Prism version 6.00 (GraphPad Software, La Jolla California USA, www.graphpad.com) to examine the differences between the mean values of undamaged leaf area. A P value less than 0.05 was considered statistically significant.

46 2.3 Results

2.3.1 Small-scale protein production and detection of clones

About 8–10 colonies were randomly selected from each YPD agar plate containing transformed P. pastoris cells carrying genes encoding Hv1a, GNA, and GNA-toxin fusion proteins. These colonies were cultured in small-scale culture media (10 mL) for small-scale expressions as described in Materials and Methods. Based on the results from both SDS-PAGE and western blots we confirmed expression of the recombinant proteins in P. pastoris.

SDS-PAGE analysis of Hv1a expression (not shown) revealed a band of the expected size (~6 kDa for His6-TEV-Hv1a) for three colonies. One of these colonies, number 3 was selected for large-scale production of the Hv1a.

In case of GNA, two protein bands, one with a mass of ~13.5 kDa and another with a mass of ~27 kDa were detected in the western blot (Fig. 2.3A). The band at ~13.5 kDa is much more intense and corresponds to the expected size of a GNA monomer. The weaker band at ~27 kDa is consistent with a GNA dimer. Based on the results of the small-scale expressions, colony 4 was selected for large-scale production of GNA.

As shown in Fig. 2.3B, two different sized bands (~20 kDa and ~15 kDa) were detected in the western blots used to analyse Hv1a-GNA expression. We presume that these correspond to the intact Hv1a-GNA fusion protein (~18 kDa for the His6-TEV-Hv1a-GNA) and a GNA fragment (~15 kDa) resulting from proteolytic cleavage of the fusion protein. Based on the results of the small-scale expressions, Hv1a-GNA colony 5 was selected for large-scale production of this fusion protein.

In case of GNA-Hv1a, colonies 4, 6 and 7 (0.5% methanol induction) and colony 3 (1% methanol induction) produced a band (~20 kDa) corresponding to the expected mass of the GNA-Hv1a fusion protein (~18 kDa for the His6-TEV-GNA-Hv1a) (Fig. 2.4). Colony 4 and 6 appeared to produce the highest level of protein expression at 0.5% methanol induction and hence they were chosen for large-scale protein production.

47

Figure 2.3: Western blot (probed with rabbit anti-GNA polyclonal antibody) showing expression of GNA and Hv1a-GNA in P. pastoris. (A) Western blot analysis of

His6-TEV-GNA expression revealed an intense band at ~13.5 kDa corresponding to the expected mass of a GNA monomer and a weak band at ~27 kDa, consistent with a GNA dimer. (B) Western blot analysis of His6-TEV-Hv1a-GNA revealed two major protein bands at ~20 kDa and ~15 kDa which were presumed to correspond to the intact fusion protein and a degradation product, respectively.

Figure 2.4: Western blot (probed with rabbit anti-GNA polyclonal antibody) showing expression of GNA-HV1a. Lanes 1–5 represent the expressed protein from colonies 3 to 7 respectively, each induced with 0.5% methanol, and lanes 7 to 10 represent the expressed protein from colonies 3–6 respectively, each induced with 1% methanol. The arrow indicates the expected running position (~18 kDa) of the His6-TEV-GNA-Hv1a fusion protein. The level of protein production was higher with 0.5% methanol and colonies 4 and 6 produced the highest levels of GNA-Hv1a.

48 2.3.2 Large-scale production of recombinant proteins

Recombinant proteins were produced from large-scale culture and purified using Ni-NTA affinity chromatography. The recombinant proteins were then concentrated, lyophilised and resuspended prior to further purification using RP-HPLC. RP-HPLC fractions were collected manually and analysed using ESI and MALDI mass spectrometry (MS). Fig. 2.5A shows an RP-HPLC chromatogram with three peaks marked with arrows resulting from purification of Hv1a using a C18 RP-HPLC analytical column.

Figure 2.5: Purification of recombinant Hv1a expressed in P. pastoris. (A) Chromatogram from RP-HPLC purification of Hv1a. The mass of the arrowed fractions matches that expected for fully oxidised His6-TEV-Hv1a. (B) ESI-MS spectrum from analysis of the first arrowed peak in panel A. (Similar results were obtained for the other two arrowed peaks.) The measured mass (5940.43 Da) matches that expected for

His6-TEV-Hv1a.

These three peaks seen in Fig. 2.5A could be peptide isoforms resulting from different disulfide-bonds connectivities. MS analysis (Fig. 2.5B) revealed that the three peaks

49 marked with arrows had masses corresponding to that of fully oxidised Hv1a (i.e., all six Cys residues involved in disulfide bonds); the predicted mass of the fully oxidised

His6-TEV-Hv1a is 5940.4 Da and the monoisotopic mass obtained from MS analysis of the arrowed peaks in Fig. 2.5A was 5940.3 Da. This indicates that P. pastoris is able to produce Hv1a in soluble, disulfide-bonded form.

Recombinant GNA was purified using a C4 RP-HPLC analytical column (Fig. 2.6A). The peaks marked with arrows in Fig. 2.6A had masses similar to that predicted for

His6-TEV-GNA (~13 kDa) when analysed by western blot with anti-GNA antibody (Fig. 2.6B). At this stage, it is unclear why separate RP-HPLC fractions were obtained with masses corresponding to GNA.

Figure 2.6: Purification and detection of recombinant GNA expressed in P. pastoris. (A) Chromatogram from C4 RP-HPLC purification of recombinant GNA. The peak fractions were selected after matching with the expected size for His6-TEV-GNA (~13 kDa) according to western blot analysis (panel B). (B) Western blot analysis of RP-HPLC fractions resulting from purification of GNA (panel A). Several RP-HPLC fractions yielded protein bands with masses matching the expected size of His6-TEV-GNA (~13 kDa).

50 The recombinant Hv1a-GNA fusion protein was purified using a C4 RP-HPLC analytical column (Fig. 2.7A). The peak marked with an arrow in Fig. 2.7A had a mass similar to that expected for His6-TEV-Hv1a-GNA (~18 kDa) when analysed by western blot with anti-GNA antibody (Fig. 2.7B). In contrast with the small-scale expressions, where two protein bands with different sizes were evident, Western blot analysis of the RP-HPLC fractions showed only a single protein band of mass of ~18 kDa, close to that expected for

His6-TEV-Hv1a-GNA.

Figure 2.7: Purification and detection of recombinant Hv1a-GNA expressed in P. pastoris. (A) Chromatogram from RP-HPLC purification of Hv1a-GNA. The peak fraction that has a mass matching that expected for His6-TEV-Hv1a-GNA (~18 kDa) according to western blot analyses (panel B) is highlighted with an arrow. (B) Western blot analysis of RP-HPLC fractions resulting from purification of Hv1a-GNA (panel A).

51 The recombinant GNA-Hv1a fusion protein was purified using a C4 RP-HPLC analytical column (Fig. 2.8A). The GNA-Hv1a fusion protein eluted as a peak a with retention time of ~22 min (arrowed peak in Fig. 2.8A). Western blot analysis of the RP-HPLC fractions using anti-GNA antibody (Fig. 2.8B) revealed that the peak marked with an arrow in

Fig. 2.8B has a mass corresponding to that predicted for His6-TEV-GNA-Hv1a (~18 kDa) and this was confirmed using MS (data not shown).

Figure 2.8: Purification and detection of recombinant GNA-Hv1a expressed in P. pastoris. (A) Chromatogram from RP-HPLC purification of GNA-Hv1a. An arrow marks the peak with mass matching that expected for His6-TEV-GNA-Hv1a. (B) Western blot analysis of HPLC fractions resulting from purification of GNA-Hv1a secreted by colonies 4 (lanes 6–10) and 6 (Lane 2–5). These two colonies were selected for large-scale production. However, in subsequent experiments (as shown here), colony 4 consistently produced higher levels of protein. The GNA-Hv1a band is evident at ~20 kDa.

2.3.3 Insecticidal activity of the recombinant fusion proteins

Protein concentrations for stock solutions of each of the recombinant protein were calculated using the theoretical extinction coefficients of the proteins in combination with sample absorbance at 280 nm measured using a NanoDrop 2000 spectrophotometer (Thermo Scientific). The final stock concentration of the GNA-Hv1a, Hv1a-GNA, and Hv1a was 3.9 µg/µL, 0.13 µg/µL, and 2.6 µg/µL, respectively. These solutions were used to inject M. domestica (average body weight 12.2 mg) with Hv1a or fusion proteins (Table 2.2).

52 Table 2.2. Insecticidal activity of recombinant Hv1a, GNA-Hv1a, and Hv1a GNA after injection into houseflies (Musca domestica)

Dose 0.5 hour 1 hour 24 hours Recombinant (µg/mg Paralysed Dead Paralysed Dead Paralysed Dead peptides insect) (%) (%) (%) (%) (%) (%) Water — 0 0 0 0 0 0 Hv1a 0.730 100 0 100 0 100 100 Hv1a-GNA 0.615 20 0 20 20 80 40 GNA-Hv1a 0.574 0 0 0 0 100 40

Fletcher et al. (1997) observed that, the insecticidal activity of the recombinant Hv1a was observed immediately after injection as the injected insects became paralysed within

30 min, which is the characteristic effect produced by block of insect Cav channels by Hv1a (Fletcher et al., 1997). In contrast, the insects injected with recombinant fusions of Hv1a with GNA (i.e., Hv1a-GNA or Hv1a-GNA) also showed paralysis but the effect was not immediate. However the insecticidal efficacy of the injected toxins after 24 h (Table 2.2) indicates that fusion of Hv1a to GNA does not alter its intrinsic insecticidal activity. It should be noted that the proteins were injected at similar doses by mass, and therefore the molar dose of the fusion proteins was about one-quarter that of Hv1a. Higher doses of the fusion protein are likely to induce more immediate paralysis.

In the leaf disc-feeding assay, P. Xylostella larvae were fed with Arabidopsis leaves coated with different recombinant toxins for 2 days and then the severity of feeding damage was observed (Fig. 2.9A and Fig. 2.9B). The analysis of the magnified images of the undamaged leaf tissues using ImageJ programme shows significant difference between the undamaged tissues in untreated leaves and leaves treated with recombinant Hv1a/GNA fusion proteins (Fig. 2.9B, P ≤ 0.0001). Significant difference was also found between the undamaged tissues treated with Hv1a alone and recombinant Hv1a/GNA fusion proteins (Fig. 2.9B, P≤ 0.0001). Significant difference was also found between the undamaged tissues treated with GNA alone and recombinant Hv1a/GNA fusion proteins (Fig. 2.9 B, P≤ 0.05). Untreated leaves Treatment with recombinant Hv1a or GNA and showed similar effect but GNA alone was found more effective than Hv1a alone. These results indicate that fusion of Hv1a to GNA greatly magnifies its oral activity. Fitches et al. found that fusion of Hv1a to the N-terminus of GNA (i.e., a Hv1a-GNA fusion) significantly enhanced its oral activity against larvae of the cabbage moth Mamestra brassicae, but these authors did not examine the effects of a GNA-Hv1a fusion (Fitches et al., 2012) .

53

Figure 2.9: Feeding damage by P. xylostella larvae on toxin treated Arabidopsis leaves. (A) Arabidopsis leaves were coated with water (control, top panel), recombinant Hv1a (0.633 µg/cm2 leaf), recombinant GNA (1.90 µg/cm2 leaf), recombinant Hv1a-GNA (2.532 µg/cm2 leaf) or recombinant GNA-Hv1a (2.532 µg/cm2 leaf) as indicated by the labels to the left of each panel. (B) Quantitative analysis of undamaged leaf tissues after feeding.

2.4 Discussion

Hv1a isolated from the venom of the Australian funnel-web spider Hadronyche versuta (Windley et al., 2012) has potential as an effective and environmentally safe insecticide.

This 37-residue peptide has potent insecticidal activity due to its block of insect Cav channels (Tedford et al., 2004; Chong et al., 2007). Spiders inject Hv1a (and other venom

54 components) directly into the insect haemolymph, thereby enabling it to reach its molecular target (i.e., neuronal Cav channels). In contrast, for agricultural applications; Hv1a needs to be orally active. Unfortunately, Hv1a has a poor oral activity due to its inability to pass through the insect gut layers into the haemolymph (Fitches et al., 2012).

GNA, a mannose-binding lectin, has a natural ability to cross the insect gut into the haemolymph via recognition by specific receptors (Fitches et al., 2012). Consequently, GNA has been exploited as a transporter for several different insecticidal toxins (Fitches et al., 2001; Fitches et al., 2004; Down et al., 2006; Trung et al., 2006; Wakefield et al., 2006; Fitches et al., 2012). GNA itself is weakly insecticidal in some insect species, but not as potent as insecticidal spider toxins. By fusing GNA to an insecticidal spider-venom peptide, one can couple the high insecticidal potency of the spider toxin with the gut permeability of GNA. Thus, in the work described in this chapter, we examined whether fusion of Hv1a to GNA enhanced it oral activity against diamondback moth larvae.

Glycosylation and possible proteolytic degradation

The GNA-Hv1a fusion protein produced by P. pastoris appeared as ~20 kDa band in both SDS-PAGE and western blot analyses (Figs. 2.4 and 2.8B). The same size band was also observed for Hv1a-GNA (Fig. 2.3B) in small-scale expressions. The expected theoretical size of these fusion proteins is ~18 kDa, a combination of Hv1a (~4 kDa), GNA (~12 kDa), the His6-tag, and the TEV protease cleavage site. The additional ~2 kDa on the recombinant fusion proteins may be caused by glycosylation, which is a common problem for recombinant proteins produced in yeast expression system (Palomares et al., 2004; Li et al., 2007).

P. pastoris often produces recombinant proteins that contain N- and/or O-linked glycosylation (Li et al., 2007). P. pastoris even produces glycosylated protein products when the protein is not normally glycosylated by the native host (Palomares et al., 2004). The glycosylated gene products from P. pastoris generally have shorter sugar chains compared to the products from S. cerevisiae, another yeast species commonly used for protein expression (Cregg et al., 2000). This makes P. pastoris more suitable for the production of many recombinant eukaryotic proteins.

As in the current study, Fitches et al. (2012) also found that Hv1a-GNA was glycosylated when produced in P. pastoris (Fitches et al., 2012). Despite the fact that the Hv1a-GNA and GNA-Hv1a fusion proteins in our study may have been glycosylated, the insecticidal activity of the fusion proteins seems not to have been affected by this post-translational

55 modification. Therefore, actions to avoid or to remove the glycosylation on the recombinant fusion proteins were not considered necessary.

In addition to the ~20 kDa band, a band of mass ~15 kDa was evident in the western blot analysis of small-scale expressions of Hv1a-GNA (Fig. 2.3A). This led to the hypothesis that proteolytic degradation might have occurred in the culture media. In theory, proteolytic degradation could be avoided by producing the recombinant protein in protease-deficient host strains such as SMD1163, SMD1165, and SMD1168 (Li et al., 2007). However, the same two bands (~20 kDa and ~15 kDa) were evident in western blot analyses of the recombinant Hv1a-GNA produced in P. pastoris by Fitches et al. (Fitches et al., 2012) where the SMD1168H host strain was used. However, these two proteins showed potent insecticidal activity when tested in insect bioassays. Thus, regardless of the cause of this problem, it does not seem to adversely affect the insecticidal activity of the recombinant fusion proteins.

2.5 Summary and conclusion

In summary, recombinant Hv1a, GNA-Hv1a, Hv1a-GNA and GNA were successfully produced in soluble form using P. pastoris. All four recombinant proteins were purified to >95% purity using a combination of Ni-NTA affinity chromatography and RP-HPLC. The production and purification methods established in the current study will serve as a basis for further improvements in recombinant production of these proteins.

Preliminary insect bioassays revealed that Hv1a retains its intrinsic insecticidal activity when fused to the N- or C-terminus of GNA. However, a feeding bioassay using Arabidopsis leaves revealed that the Hv1a-GNA and GNA-Hv1a fusion proteins had significantly higher oral activity than Hv1a as judged by their ability to reduce leaf damage caused by diamondback moths. Thus, expression of Hv1a-GNA or GNA-Hv1a in transgenic plants might confer resistance against lepidopteran pests, and possibly other insect pest species.

56 Chapter 3: Genetic engineering of Arabidopsis to express Hv1a proteins

3.1 Introduction

Crop loss to insect pests is a global problem. With more than 1,000,000 extant species and high evolutionary variability, insects destroy ~14% of the world's total agricultural production each year, despite extensive control measures (Oerke et al., 1994; Oerke and Dehne, 2004; Oerke, 2006). Conventional chemical approaches that were initially highly successful have waned in their ability to control insect pests due to the widespread development of resistance in insect populations, a consequence of repeated and indiscriminate use of insecticides (Windley et al., 2012; Smith et al., 2013).

Genetic engineering techniques have made it possible to transform crop plants with desirable traits from any source. The most successful and widespread insecticidal traits transferred into plants are sequences coding for Bacillus thuringiensis (Bt) Cry toxins, which target the insect digestive system. Since the introduction of Bt crops, some lepidopteran species have evolved various levels of resistance (Tabashnik et al., 2013). Thus, different Bt genes have been isolated to cope with resistance to specific Bt toxins.

The search for new genes with insecticidal traits is ongoing. Insecticidal peptide toxins from spider venom are one of the most promising candidates for new insecticide development (King and Hardy, 2013). Attempts to engineer insect-resistant plants expressing insecticidal spider-toxin transgenes began almost 30 years ago (Hong et al., 1996).

Insecticidal spider-venom peptides are highly toxic to insect pests when injected (Fitches et al., 2012; Smith et al., 2013). However, these peptides are largely ineffective when delivered orally or when applied topically as their molecular targets are located in the central or peripheral nervous system, or other sites only accessible from the circulatory system (Fitches et al., 2012; King and Hardy, 2013; Smith et al., 2013). The low level of oral activity presumably results from a slow rate of absorption of insecticidal venom peptides in the insect gut, as observed previously for insect neuropeptides (Audsley et al., 2008) and disulfide-rich peptides from scorpion and snake venoms (Casartelli et al., 2005). Enhancement of the oral activity of insecticidal venom peptides by any suitable approach could improve the commercial importance of those peptides in crop protection (King and Hardy, 2013). One promising approach is to fuse the peptides to carrier proteins that facilitate their transport across the insect gut. As demonstrated in the previous chapter, fusion of venom peptides with GNA (Fitches et al., 2012) was an effective way to increase oral activity. An alternative approach that was recently reported was fusion of

57 insecticidal venom peptides to the coat protein (CP) of pea enation mosaic virus (PEMV); this approach provided selective insecticidal activity against aphids, which are vectors for PEMV and other luteovoiruses and have gut receptors that bind CP (Miller and Bonning, 2003; Bonning et al., 2014). This approach dramatically improved the oral activity of Hv1a against a wide range of aphids (Bonning et al., 2004).

Thus, we decided to use Arabidopsis as a model system to examine whether GNA and CP fusions to Hv1a could provide resistance to lepidopterans and aphids, respectively. Arabidopsis has been used as a model plant for more than 60 years, initially because of its short life cycle, high yield of seeds and its ability to self-pollinate (Somerville and Koornneef, 2002). Arabidopsis was studied more vigorously in the 1990s as it was realised that, in addition to these desirable genetic traits (Redei, 1992), Arabidopsis had a small genome (125 MB) and adult plants could be transformed without tissue culture by Agrobacterium tumefaciens (Bechtold et al., 1993). The Arabidopsis genome was sequenced in 2000 (The Arabidopsis Initiative, 2000) and it is now in use for plant research in over 16,000 labs worldwide.

Although several methods for plant transformation are available, those that use the soil bacterium Agrobacterium are the most commonly used (Hooykaas, 1989). Agrobacterium is a widespread, naturally occurring soil bacterium that causes crown gall and it has the ability to introduce new genetic material into plant cells (Gelvin, 2003). Initially it was believed that Agrobacterium only infects dicotyledonous plants, but it was later established that it could also be used to transform monocotyledonous plants such as rice (Ignacimuthu and Raveendar, 2011). The genetic material that is introduced is called T-DNA (transferred DNA), which is located on a Ti plasmid. This natural ability to alter the plant genome was harnessed in the 1980s. Since it was first used to generate transgenic plants (Fraley et al., 1983), Agrobacterium has been widely used for introducing transgenes into plants for the purposes of both basic research and the generation of commercially used transgenic crops (Hinchee et al., 1988; Broothaerts et al., 2005; Bhalla and Singh, 2008; Oltmanns et al., 2010; Pitzschke and Hirt, 2010). The overall advantages of using Agrobacterium-mediated transformation over other transformation methods are simpler transgene insertion into the plant genome with less rearrangements (Jones et al., 2005).

The Arabidopsis “floral dip” method (Bechtold et al., 1993; Clough and Bent, 1998) technique involves immersion of flowers into a suspension of Agrobacterium carrying a gene of interest. The seeds collected from these transformed “T0” plants are germinated under selection to identify transgenic “T1” individuals. This method has been used in

58 hundreds of laboratories throughout the world, and using this approach ~1% of T1 seedlings are transgenic (Bent, 2006).

In this chapter, we designed vectors encoding fusions of Hv1a to GNA or CP that were used for Agrobacterium-mediated genetic transformation of Arabidopsis.

3.2 Materials and methods

3.2.1 Fusion protein expression constructs

We designed sequences with Hv1a fused at its N- or C-terminus to GNA (GNA-Hv1a and Hv1a-GNA) or CP (CP-Hv1a and Hv1a-CP) (Fig. 3.1). An ER signal sequence (23 amino acids, MAKASLLILAAIFLGVITPSCLS) preceded these sequences. We also designed sequences encoding unfused GNA, CP, or Hv1a (Fig. 3.1). A triplet of alanine residues (AAA) was used to link Hv1a to the fusion proteins. For GNA, residue Cys109 was mutated to Ser mutation to avoid non-native disulfide bond formation.

Synthetic genes encoding each of these sequences were made by GeneArt® (Fig. 3.2) and cloned into their in-house plasmid pMA-T with restriction sites that would enable cloning into the binary vector pAOV (Fig. 3.3) (Mylne and Botella, 1998) with the Cauliflower Mosaic Virus 35S promoter (CaMV35S or 35S) (Odell et al., 1985) or Arabidopsis thaliana SUCROSE TRANSPORTER 2 promoter (SUC2, At1g22710) (Sauer and Stolz, 1994; Truernit and Sauer, 1995). At the N-terminus of each ORF we added BamHI and NcoI sites, and at the C-termini we added a SacI site; these restriction sites allowed cloning of the ORFs as BamHI/SacI fragments into pAOV. It also allowed cloning of each synthetic gene as an NcoI/SacI fragment into pAOV modified to contain the SUC2 promoter.

59

Figure. 3.1: Amino acid sequences encoding Hv1a (red), GNA (green), CP (purple), and fusions of Hv1a to either the N- or C-terminus of GNA (GNA-Hv1a and Hv1a- GNA) or CP (CP-Hv1a and Hv1a-CP). All constructs include an ER signal sequence (brown) and fusion proteins contain a tri-alanine linker (blue).

60

Figure 3.2: Synthetic genes encoding the designed peptide sequences. (A) Hv1a; (B) GNA; (C) CP; (D) Hv1a-GNA; (E) GNA-Hv1a; (F) Hv1a-CP; and (G) CP-Hv1a. The size of each construct (in bp) is indicated, and engineered restriction sites are indicated.

Figure 3.3: Schematic representation of pAOV (Mylne and Botella, 1998). Restriction enzyme sites are shown for the cloning of inserts in multiple cloning sites (MCS). LB and RB represent T-DNA left and right border sequences, respectively. The nopaline synthase promoter and terminators are represented as nos and nos3′, respectively. 35S and ocs3′ indicates CaMV35S promoter and octopine synthase 3′ UTR respectively. The bialophos resistance gene (bar) encodes phosphinothricin acetyltransferase (Thompson et al., 1987). 1Restriction site not tested. ★Not unique.

61 To construct pAOV transformation vectors (35S-MCS-nos), 25–80 ng of each ORF (25–30 ng for ~200 bp ORF, 50 ng for ~400 bp ORF and 80 ng for ~700 bp ORF) was obtained from the GeneArt® vector using BamHI and SacI digestion followed by agarose gel electrophoresis and gel purification (QIAGEN). Fragments were then ligated directly into similarly cut pAOV (200 ng) using Quick-Stick Ligase (Bioline) (Sambrook et al., 1989).

For construction of transformation vectors with a SUC2 promoter, 25–80 ng of each ORF (25–30 ng for ~200 bp ORF, 50 ng for ~400 bp ORF and 80 ng for ~700 bp ORF) destined for 35S-less pAOV was obtained from the GeneArt® vector by digestion with NcoI and SacI. Fragments were triple ligated with SUC2 (120 ng) liberated from XhoI/NcoI digested pGEMT-easy (Promega) vector carrying SUC2 (obtained from Dr Joshua S. Mylne, Institute for Molecular Bioscience, The University of Queensland, Australia) and XhoI/SacI-digested pAOV (200 ng), which liberates the 35S promoter. There was no selectivity in the ligation for SUC2 over 35S apart from a higher concentration of SUC2 fragment, but the size differential between SUC2 and 35S was used to ensure SUC2 incorporated.

3.2.1.1 Diagnostic restriction digestion of the transformation vectors plasmids

The final constructs were ~25 kb in size and were digested with different sets of restriction enzymes and their sizes judged by agarose gel electrophoresis was used to confirm constructs instead of conventional sequencing, which is difficult with such large plasmids.

The final constructs were digested with three sets of restriction endonucleases (Fig. 3.4). Those containing the 35S promoter were digested with XhoI/EcoRI, XhoI/SacI or BamHI/EcoRI (Fig. 3.4A). Those containing the SUC2 promoter were digested with XhoI/EcoRI, XhoI/SacI or NcoI/EcoRI (Fig. 3.4B). The expected size of the digested fragments is given in Table 3.1. The plasmids for each construct that produced the three expected sized fragments were selected for transformation of A. tumefaciens.

62

Figure 3.4: Diagnostic restriction digestion strategies for (A) vectors containing the 35S promoter, and (B) vectors containing the SUC2 promoter. The horizontal lines parallel to the plasmid sequence represent the fragment size after double digestion with three combinations of restriction endonuclease.

Table 3.1: Expected fragment size after double digestion of the constructs

ORF Expected fragment size after double digestion Sl. Constructs size (Approximate bp) No. (bp) XhoI/EcoRI XhoI/SacI BamHI/EcoRI NcoI/EcoRI 1 pAOV-35S-ER-Hv1a 204 1300 1050 450 - 2 pAOV-35S-ER-GNA 408 1510 1260 660 - 3 pAOV-35S-ER-CP 750 1850 1600 1000 - 4 pAOV-35S-ER-Hv1a-GNA 528 1630 1380 780 - 5 pAOV-35S-ER-GNA-Hv1a 528 1630 1380 780 - 6 pAOV-35S-ER-Hv1a-CP 870 1970 1720 1120 - 7 pAOV-35S-ER-CP-Hv1a 870 1970 1720 1120 - 8 pAOV-SUC2-ER-Hv1a 204 2550 2300 - 450 9 pAOV-SUC2-ER-GNA 408 2760 2510 - 660 10 pAOV-SUC2-ER-CP 750 3100 2850 - 1000 11 pAOV-SUC2-ER-Hv1a-GNA 528 2880 2630 - 780 12 pAOV-SUC2-ER-GNA-Hv1a 528 2880 2630 - 780 13 pAOV-SUC2-ER-Hv1a-CP 870 3220 2970 - 1120 14 pAOV-SUC2-ER-CP-Hv1a 870 3220 2970 - 1120

63 3.2.2 Transformation of A. tumefaciens by tri-parental mating

Cloning plasmids can be introduced into A. tumefaciens by electroporation or tri-parental mating. We introduced the binary vectors into Agrobacterium by tri-parental mating (Hoekema et al., 1983) using Agrobacterium strain LBA4404 as the recipient, E. coli (TOP10, Invitrogen) carrying the construct as the donor, and a helper pilus positive E. coli DH5α (HB101 pRK2013). The three strains were mixed on LB-agar plates for 20 h at 28°C and streaked onto Rif50Tet2 LB-agar plates and re-streaked to Rif50Tet2Strep25 LB-agar plates. Agrobacterium colonies raised form this second selection were cultured and glycerol stocks made.

3.2.3 Plant materials

Arabidopsis thaliana (L.) Heynh. (Brassicaceae) Columbia-0 wild type (WT) seeds obtained from Dr Joshua S. Mylne (Institute for Molecular Bioscience, The University of Queensland, Australia) were used for expression of transgenes.

Seeds from a quadruple mutant (cyp79B2 cyp79B3 myb28 myb29) glucosinolate null (hereafter referred to as gluc-null) Arabidopsis were obtained from Professor Jonathan Gershenzon (Max Plank Institute for Chemical Ecology, Germany) via his collaboration with Professor Myron P. Zalucki (School of Biological Sciences, The University of Queensland, Australia). This gluc-null was also used for transformation because it is devoid of indole and aliphatic glucosinolates (Muller et al., 2010).

Glucosinolates are a diverse group of defensive secondary metabolites (Muller et al., 2010) which is used in endogenous chemical defence against attack by insect herbivores (Stauber et al., 2012). These plant defence compounds are amino-acid derived thioglucosides that are present in essentially all genera of the Brassicales (Fahey et al., 2001; Halkier and Gershenzon, 2006; Mithen et al., 2010). The glucosinolates in Arabidopsis can be converted into a large number of defensive metabolites during herbivory (Halkier and Gershenzon, 2006; Hopkins et al., 2009; Ahuja et al., 2010; Wittstock and Burow, 2010; Bohinc et al., 2012; Kos et al., 2012; Mithofer and Boland, 2012; Rohr et al., 2012; Stauber et al., 2012).

WT and gluc-null Arabidopsis plants were grown on plastic trays/pots containing Arabidopsis soil mix (mixture of peat moss, vermiculite and washed sand). Seed germination was synchronized by treatment at 4°C for 3–5 days. Trays were placed under long day-light (16 h light) or short day light (12 h light) at 23°C.

64 3.2.4 Floral-dip transformation of Arabidopsis

The best plants for transformation have many immature flower clusters and few fertilized siliques, although a range of plant stages can be successfully transformed (Clough and Bent, 1998). Pre-existing siliques were cut off using scissors, and the prepared plants were transformed by floral dip (Fig. 3.5) (Bechtold et al., 1993; Clough and Bent, 1998).

Figure 3.5: Standard floral dip method to transform Arabidopsis. (A) Arabidopsis plants ready for transformation (B) dipping/immersion.

Transformed plants were left overnight on their sides in a plastic box sealed with plastic film and uncovered the following day and raised to maturity. Seeds were harvested, sieved and stored in paper envelopes with proper labelling in an airtight plastic container containing silica gel.

3.2.5 Selection of transgenic plants using selectable markers

The transformation vector we constructed to use in plant transformation confers resistance to Basta herbicide (glufosinate ammonium, Bayer Crop Science). Basta selection was performed by sowing approximately 0.5 mL of T0/T1 seeds in soil trays. Seedlings were sprayed first upon emergence and twice afterwards at 3-day intervals with Basta solution (0.2 g glufosinate ammonium in 1 L water). Only Basta-resistant plants survived and produced true leaves. The Basta-resistant plants were T1 transgenic plants and these were transplanted to pots containing Arabidopsis soil mix and grown to obtain seeds.

65 3.2.6 PCR confirmation of transgene integration

To confirm the transgenic lines bearing the desired Hv1a gene, genomic DNA was extracted from Arabidopsis T2 lines (Edwards et al., 1991).

For 35S containing lines a primer for the 35S (5′-TTC GCA AGA CCC TTC CTC TA-3′) and nopaline synthase (nos) terminator (5′-AAG ACC GGC AAC AGG ATT C-3′) were used for PCR to amplify each ORF plus 128 bp of its flanking sequence from the binary vector. For SUC2-containing lines, a primer for SUC2 (5′-CAC GTG TCA CGA AGA TAC CC-3′) was used with the aforementioned primer for the nos terminator and this produced PCR product 268 bp larger than the ORF.

As a positive control for the extracted gDNA, ROC7 forward (5′-TGA AGT GCG CCT AAT TTG TG-3′) and reverse (5′-AGG CAA AGA GCC GAT GTA AA-3′) primers were used to amplify a 800 bp fragment of the endogenous Arabidopsis cyclophilin gene ROC7 (At5g58710).

For transgenic lines in the gluc-null background where Hv1a expression is under control of the 35S promoter, a primer for the GNA signal sequence (5′-TTC TCG CTG CTA TCT TTC TCG-3′) and nopaline synthase (nos) terminator (5′- AAG ACC GGC AAC AGG ATT C-3′) were used for PCR to amplify each ORF plus 46 bp of flanking sequence from the binary vector.

3.2.7 Segregation analysis of transgenic lines

To obtain single locus, homozygous transgenic lines we monitored the herbicide selectable marker. Approximately 200 seeds from T2 transgenic lines were sprinkled on moist filter paper (Whatman No.1 filter paper, 90 mm diameter) in a tissue culture dish (92 × 17 mm) with a layer of ground vermiculite (10–12 g/dish) pre-wet with 15 mL Basta herbicide-solution (13.33 mg/L glufosinate ammonium). Dishes were sealed with parafilm or porous tape and, after 3–4 days of stratification at 4°C, grown in default condition as stated before. The plants were grown only to the cotyledon stage at which point it was possible to distinguish herbicide-resistant from non-resistant seedlings. Non-transgenic seedlings germinated, but became pale yellow or white soon after and the cotyledons did not fully expand. Resistant seedlings were green in colour and the cotyledons fully expanded.

The numbers of herbicide-resistant and susceptible seedlings were recorded to calculate the segregation ratio and select the single locus transgenic lines (75% resistant plants,

66 25% susceptible). Single-locus lines were then grown to obtain T3 seeds so that a proportion of these would be homozygous lines and 100% resistant to Basta. These genetically stable, single-locus, homozygous transgenic lines were used in further studies.

3.2.8 Detection of in planta expressed proteins

To detect in planta expression of Hv1a and Hv1a fusion proteins, protein was extracted from transgenic Arabidopsis leaf tissue (Martinez-Garcia et al., 1999). Approximately 0.1 g of leaf tissue from each line was collected in a 1.5 ml tube then frozen immediately in either dry ice or liquid nitrogen and stored at –80°C. Approximately 0.1 g of floral tissue was collected from the transgenic lines expressing GNA, as the antibody used to detect GNA was found to cross-react with leaf protein.

Each 0.1 g of plant tissue was ground in 1.5-ml tubes using plastic pestle with a pinch of glass beads (Sigma-Aldrich, Cat. G4649) and 0.5 mL of plant protein extraction buffer (25 mM Tris, 0.15 M sodium chloride, pH 7.6). Samples were vortexed for 30 s and centrifuged at 10,000 g for 15 min at 4°C. After centrifugation, 0.3 mL of clarified supernatant was transferred to a new tube and stored at –80°C.

Protein extracts were separated by SDS-PAGE and transferred to Hybond ECL nitrocellulose membrane (Amersham) by electroblotting to be analysed by western blotting. Membranes were blocked with Odyssey western blocking buffer (LICOR Biosciences) and incubated with primary antibody (anti-Hv1a or anti-GNA) overnight at 4°C. A rabbit anti-Hv1a polyclonal antibody (Vestaron Corporation, MI, USA) was used to detect Hv1a. A rabbit polyclonal anti-GNA antibody (from Dr Elaine Fitches, Food & Environment Research Agency, Sand Hutton, UK) was used to detect GNA. No antibody was available for CP. The primary antibody solutions were diluted 1:2000 in Odyssey western blocking buffer.

A 1:10,000 dilution of IRDye® 800CW goat polyclonal anti-rabbit IgG (LICOR Biosciences) in Odyssey western blocking buffer was used as secondary antibody. Blots were scanned with an Odyssey infrared imaging system (LICOR Biosciences) using the 800 nm channel to detect fluorescence from the bound secondary antibody.

The mature Hv1a peptide (predicted 4055.44 Da) was detectable as a band on blots probed with rabbit polyclonal anti-Hv1a antibodies. Mature GNA (11641.95 Da) was detectable in blots probed with rabbit polyclonal anti-GNA antibodies. Hv1a fusions with GNA (predicted 15,892.61 Da) were also detected using rabbit polyclonal anti-Hv1a. The

67 Hv1a fusions with CP (predicted 28,170.64 Da) were detected using rabbit polyclonal anti-Hv1a antibodies.

3.3 Results

3.3.1 Detection of the transgenes in Arabidopsis

A Total of 64 transgenic lines were confirmed by PCR (Figs 3.6–3.9). PCR products were obtained for 58 and all 58 matched the expected size. The 6 lines that did not produce bands might be due to the failure of that particular PCR reaction and were not used for further studies. These results confirmed successful integration of transgenes into the plant genome and a low frequency of escapes.

3.3.2 Detecting in planta peptide expression

The in planta expression of Hv1a and Hv1a fusion proteins under control of the 35S promoter was confirmed by western blotting of the plant protein extract probed with specific antibodies (Figs. 3.10–3.15). In all cases, the protein band matched the expected sizes. Several faint bands were also found in all samples (Figs. 3.10–3.15), which are either an artifact of the gel or represents cross reactivity of the antibody with endogenous Arabidopsis proteins. Some bands smaller than the expected size were also observed (Figs. 3.13–3.15), which might represent degradation products of the fusion proteins. Cross reactivity was observed with anti-GNA antibody with endogenous Arabidopsis proteins (Figs. 3.11 and 3.15b).

In case of western blotting with protein extracts from transgenic plants expressing the proteins in phloem tissue under SUC2 promoter control, a very faint band was observed for expected protein size (data not shown). Phloem tissue specific protein expression under SUC2 promoter control was expected to be much lower than the 35S lines.

68

Figure 3.6: PCR analysis of transgenic Arabidopsis lines generated on a WT background. PCR amplifications are for the transgene (T) using primers for the nos sequence and either the 35S or SUC2 sequence. The endogenous ROC7 gene (R) provided a positive control for the genomic DNA. Lanes are labelled above the gel with their particular transgene.

69

Figure 3.7: PCR analysis of transgenic Arabidopsis lines in WT background. PCR amplifications are for the transgene (T) using primers for the nos sequence and SUC2 sequence. Lanes are labelled above with their particular transgene.

Figure 3.8: PCR analysis of transgenic Arabidopsis lines generated in the gluc-null background. PCR amplifications are of the transgene using primers for the ER signal and nos sequence. Lanes are labelled above the gel with their particular transgene.

70

Figure 3.9: PCR analysis of transgenic Arabidopsis lines generated in WT background. PCR amplifications are of either the transgene using primers for the 35S and nos sequence or SUC2 and nos sequence. Lanes are labelled above the gel with their particular transgene.

71

Figure 3.10: Western blot analysis of 35S-Hv1a homozygous transgenic Arabidopsis lines, probed with rabbit polyclonal anti-Hv1a antibody (1:2000 dilution). Arrow indicates the expected size of Hv1a.

Figure 3.11: Western blot analysis of 35S-GNA homozygous transgenic Arabidopsis lines, probed with rabbit polyclonal anti-GNA antibody (1:2000 dilution). Arrow indicates the expected size of GNA.

72

Figure 3.12: Western blot analysis of 35S-Hv1a-GNA homozygous transgenic Arabidopsis lines probed with rabbit polyclonal anti-Hv1a antibody (1: 2000 dilution). Arrow indicates the expected size of Hv1a-GNA.

Figure 3.13: Western blot analysis of 35S-GNA-Hv1a homozygous transgenic Arabidopsis lines, probed with rabbit polyclonal anti-Hv1a antibody (1: 2000 dilution). Arrow indicates the expected size of GNA-Hv1a.

Figure 3.14: Western blot analysis of (A) 35S-Hv1a-CP and (B) 35S-CP-Hv1a homozygous transgenic Arabidopsis lines, probed with rabbit polyclonal anti-Hv1a antibody (1: 2000 dilution). Arrows indicate the expected size of Hv1a-CP and CP-Hv1a.

73

Figure 3.15: Western blot analysis of proteins in 35S homozygous transgenic Arabidopsis lines in the gluc-null background, probed with either (A) rabbit polyclonal anti-Hv1a antibody (1:2000 dilution), or (B) rabbit polyclonal anti-GNA antibody (1:2000 dilution). Arrows indicate the expected sizes of proteins.

74 3.4 Discussion

In the present study, transgenic Arabidopsis plants were generated to express Hv1a and its fusions with either GNA or CP. The transgenes encoding these peptides were inserted into the binary vector pAOV under control of either the constitutive 35S or phloem-specific SUC2 promoter.

Transformation vectors were introduced into Agrobacterium and used to generate transgenic Arabidopsis plants via Agrobacterium-mediated transformation using the floral dip technique. Transgenic Arabidopsis lines were selected for resistance to

Basta herbicide. PCR amplification of genomic DNA from T2 generation transgenic Arabidopsis lines proved the transgenes successfully integrated into the Arabidopsis genome. Western blots of T3 homozygous transgenic Arabidopsis lines confirmed that we had expressed Hv1a and its fusions with either GNA or CP in planta in Arabidopsis. These transgenic Arabidopsis lines were generated to use for subsequent analysis to determine the host plant resistance conferred by the in planta expressed Hv1a alone or fused with GNA.

The recombinant fusion proteins of GNA and Hv1a showed better insecticidal activity by helping the delivery of the orally fed Hv1a from insect gut to haemocoel. Both the C and N terminal fusions of GNA and Hv1a also showed insecticidal activity in injection bioassay and better resistance against insects in feeding bioassay (Chapter 2). The transgenic plants expressing the C and N terminal fusions of GNA and Hv1 will provide the opportunity to assess the insecticidal performance of the Hv1a with improved delivery system (mentioned in Chapter 4). The native glucosinolates in the test plants used in generating transgenic plants might interfere in the feeding behaviours and developments of some insect pest species. So to reduce the interfering effects of native glucosinolates, a set of transgenic plants expressing the C and N terminal fusions of GNA and Hv1 also generated in Gluc-null mutant background.

The in planta expressed fusion protein of Hv1a with viral coat protein showed better insecticidal activity by helping the delivery of the orally fed Hv1a-CP fusion protein in aphid vector. The transgenic plants expressing the C and N terminal fusions of Hv1a and CP will provide the opportunity to assess any enhancement in the delivery of in planta expressed toxins by insecticidal performance of the fusions of Hv1a with CP in different combinations. Insecticidal assays using the generated transgenic lines expressing the fusions of Hv1a and CP are still ongoing and the result is not

75 mentioned in this thesis.

As mentioned in Chapter 1, the tissue specific expression of the toxins in planta might be an important mechanism to improve specificity to the target pests and thus limit the non-target effects of the expressed toxins. The transgenic plants expressing the fusions of Hv1a and GNA or the fusions of Hv1a and CP under phloem tissue specific SUC2 promoter will provide the opportunity to assess insecticidal performance and the efficacy of the tissue specific in planta expressed toxins in controlling phloem feeding insect pests. Insecticidal assays using the generated transgenic lines expressing the fusions of Hv1a and GNA or the fusions of Hv1a and CP under phloem tissue specific SUC2 promoter are still ongoing and the result is not mentioned in this thesis.

3.5 Summary and conclusion

Hv1a and fusions with either GNA or CP were successfully expressed in planta in Arabidopsis. These transgenic plants provide an opportunity to compare the level of insect resistance conferred by each construct.

76 Chapter 4: Hv1a transgenes protect plants from lepidopteran pests

4.1 Introduction

Phytophagous lepidopterans are considered the most destructive crop pests; about 40% of global chemical insecticide use is to control heliothine moths such as the cotton bollworm Helicoverpa armigera and the tobacco budworm Heliothis virescens (McCaffery, 1998). Helicoverpa species (Fig. 4.1) are polyphagous pests of around 200 plant species including cotton, corn, soybeans, tobacco, tomato, chickpea, legumes and other vegetable crops (Zalucki et al., 1986; Fitt, 1989). They are one of the most serious agricultural pests in cotton-producing countries including Australia, China, India and the USA, causing huge economic losses each year (Stevens et al., 2012). These pests are problematic because of their feeding preference for nitrogen-rich plant structures, especially reproductive structures and growing points such as flowers, fruits and young leaves (Stevens et al., 2012). Damage to these reproductive structures in plants of economic importance has a direct influence on crop yield.

Figure 4.1: The life cycle of the cotton bollworm (H. armigera) (Stevens et al., 2012).

Cotton bollworms are leaf feeders at the early instar stage and move to developing fruits at later stages. They are a major problem throughout the world, including Australia, as they have become resistant to most commercially available chemical insecticides (McCaffery, 1998; DAFF, 2011; Yang et al., 2013). Compared to other lepidopteran pest species, H. armigera is less migratory from their infestation area due to different ecology and behaviour; thus, field populations subject to insecticide control regimes are exposed to consistent selection pressure, thereby accelerating the evolution of resistance to the applied insecticide (Fitt, 1994; DAFF, 2011).

77 Long-term use of chemical insecticides has also led to concerns regarding food safety, human health, and the environment (Stevens et al., 2012). The importance of chemical insecticides to prevent crop losses from insect pests cannot be ignored, but there is a huge demand to develop alternative or additional approaches that would permit more selective use of chemical insecticides (Sharma et al., 2000). Enhancing the insect resistance of host plants through incorporation of transgenes encoding insecticidal proteins is one of the most recently introduced insect control technologies. As the products of most transgenes are ingested by the insect pest and therefore act through the gut, most research has concentrated on transgene-encoded proteins that target the insect mid-gut and/or the peritrophic membrane to interrupt digestion or nutrition (Murdock et al., 1990; Eisemann et al., 1994; Harper et al., 1998; Hopkins and Harper, 2001). Generally, the detrimental effects of these proteins on larval growth results from limited assimilation of nutrients due to disrupted digestion (Williams, 1999). Furthermore, the induced delay in growth and development prolongs the period in which the larvae are exposed to natural enemies such as mice, spiders and insect predators (Stevens et al., 2012).

As discussed in Chapter 2, in order for insecticidal peptides that target the insect nervous system to be orally active, they must be fused to a carrier protein that enables them to traverse the insect gut epithelium in order to reach their molecular target. We demonstrated that the insecticidal activity of Hv1a could be markedly improved by fusing it to the plant lectin GNA. In this chapter, I examine whether a transgene encoding Hv1a-GNA or GNA-Hv1a in Arabidopsis confers resistance against the recalcitrant lepidopteran pest H. armigera.

4.2 Materials and methods

4.2.1 Test plants

For these experiments, we selected single locus homozygous transgenic Arabidopsis lines expressing Hv1a, GNA, Hv1a-GNA or GNA-Hv1a under 35S promoter control. Wild type (WT) Arabidopsis were used as a control. Arabidopsis were grown in 12 h days of cool white fluorescent light for six weeks at 23°C with a relative humidity of ~70% until a ~20 rosette leaf was reached. In order to study the effect on H. armigera of endogenous glucosinolates produced by WT Arabidopsis, we also included a glucosinolate null (gluc-null) line (cyp79B2 cyp79B3 myb28 myb29 quadruple mutant, deficient in production of glucosinolates) (Sun et al., 2009). Single locus homozygous transgenic lines expressing the transgenes under 35S promoter control in gluc-null background were also included in the experiment.

78 4.2.2 Test insect species

Laboratory cultures of H. armigera neonates were obtained from Professor Myron Zalucki (School of Biological Sciences, The University of Queensland, Australia) and AgBiTech Pty Ltd. (Queensland, Australia).

4.2.3 Insect bioassay with H. armigera feeding on transgenic Arabidopsis

To investigate whether host-plant resistance was conferred by the incorporated transgenes, we performed feeding bioassays with H. armigera on transgenic Arabidopsis lines expressing the transgenes. The bioassays were designed to determine insect feeding preferences as well as post-ingestion effects.

4.2.3.1 Whole-plant bioassays

In whole-plant bioassays, neonates of H. armigera were introduced onto 6-week-old transgenic or WT Arabidopsis, then the plants were caged and larvae allowed consume leaves freely (Fig. 4.2). Mortality was recorded on the 3rd, 5th, and 7th day after insect release and larval weight was recorded on the 9th day after insect release.

Figure 4.2: Set up of whole-plant insect bioassay.

4.2.3.2 Detached-leaf bioassays

In detached-leaf assays, H. armigera neonate was placed in a humid Petri dish and allowed to feed continuously on leaves from 6-week-old Arabidopsis for 7–15 days (Fig. 4.3). Agar gel (2%) was used in the bottom of the Petri dish to keep leaves turgid and thus more appetizing for larvae. Petri dishes were maintained at 23°C and 70% relative humidity. All larvae were fed with a continuous supply of leaf and the quantity increased with larval growth. Mortality of the neonatal larvae was recorded on the 3rd, 5th, and 7th day

79 after insect release and the larval weight was recorded on 9th day after insect release. In an extended-duration feeding assay, mortality was recorded 3, 5, 7, 9, 11, 13, and 15 days after the release of larvae.

Figure 4.3: Set up of detached-leaf insect bioassay with H. armigera.

To assess the extent of insect resistance conferred by the various transgenes, H. armigera neonates were allowed to feed continuously for 5 days on leaves (~0.1 g leaf tissue) from transgenic Arabidopsis lines. Leaf weight was measured before insect release and after 5 days of feeding to determine the amount of leaf tissue consumed by the larvae.

4.2.4 Western blot analysis of insect hemolymph

To verify whether the insect mortality observed in bioassays was due to ingestion of in planta expressed insecticidal toxin, western blot analysis was performed with insect hemolymph. Insect hemolymph was collected after crushing and centrifuging dead larvae from the feeding bioassays. Western blotting was performed with rabbit anti-Hv1a polyclonal antibody to detect the presence of Hv1a in insect hemolymph.

4.2.5 Statistical analysis

Kaplan-Meier survival curves were constructed using R software (R Development Core Team., 2013) using the recorded survival data for larvae feeding on different Arabidopsis lines. Kaplan-Meier survival curves were compared using a log-rank test (Dalgaard, 2008). Data were analysed using a Student’s t-test or one-way ANOVA using GraphPad™ Prism 6.00 (GraphPad Software, La Jolla California USA, www.graphpad.com) software to examine the differences between the mean values of parameters such as larval mortality, larval weight, and leaf damage. A P value less than 0.05 was considered statistically significant.

80 4.3 Results

4.3.1 Resistance of transgenic Arabidopsis to H. armigera

4.3.1.1 Mortality and development of H. armigera larvae fed on Hv1a transgenic plants constructed on a WT background

We initially examined the mortality of H. armigera larvae fed on transgenic plants constructed in a WT background (i.e., plants containing endogenous glucosinolates). No significant difference was found between the survival of H. armigera larvae feeding on WT and Hv1a plants (P = 0.656) or WT and GNA plants (P = 0.804) (Fig. 4.4A, log-rank test on Kaplan–Meier survival curves)

Surprisingly, no significant difference was found between the survival of H. armigera larvae feeding on WT and Hv1a-GNA plants (Fig. 4.4A, black line versus dark blue dashed/dotted line; log-rank test on Kaplan–Meier curves yielded P = 0.353). However, the difference in survival between H. armigera larvae feeding on WT and GNA-Hv1a plants (Fig. 4.4A, black line versus cyan dashed line) was close to being statistically significant; a log rank test on the Kaplan–Meier survival curves yielded P = 0.0554.

The average weight of H. armigera larvae fed on WT or different transgenic Arabidopsis expressing Hv1a, GNA, Hv1a-GNA and GNA-Hv1a was found to be 3.134 mg, 0.925 mg, 2.05 mg, 1.86 mg and 0.72 mg, respectively (Fig. 4.4B). A one-way ANOVA followed by multiple comparisons test showed significant difference between the average weight of larvae after feeding on WT and transgenic Arabidopsis expressing Hv1a, GNA, Hv1a-GNA or GNA-Hv1a (F = 4.92, d.f. = 4, P = 0.0031).

The highest mortality and the largest decrease in larval weight relative to WT controls were found for GNA-Hv1a plants. This suggests that this fusion protein is conferring resistance against H. armigera. However, a major problem with the interpretation of these experiments was the high level of mortality observed with WT control plants. This made it difficult to determine whether the increased mortality seen in larvae feeding on transgenic plants expressing Hv1a transgenes is significant.

81

Figure 4.4: Insect bioassay with H. armigera neonates feeding on leaves of WT or transgenic Arabidopsis expressing Hv1a, GNA, GNA-Hv1a, or Hv1a-GNA in a WT background. (A) Kaplan-Meier survival curves for H. armigera neonates feeding on leaves of WT or transgenic Arabidopsis expressing Hv1a, GNA, GNA-Hv1a, or Hv1a-GNA in a WT background. (B) Weight of larvae (mean ± SEM) after 9 days feeding on WT or transgenic Arabidopsis lines. Asterisks indicate statistically significant differences (p < 0.05).

We wondered whether the high mortality seen with control plants might be due to the endogenous glucosinolates produced by Arabidopsis. Glucosinolates are plant secondary metabolites, found in members of the Brassicaceae family, which have insecticidal effects on insect pests such as H. armigera (Hopkins et al., 2009; Muller et al., 2010; Bohinc et al., 2012; Kos et al., 2012; Mithofer and Boland, 2012). Herbivory of Arabidopsis by H. armigera has been shown in induce production of indolic glucosinolates (Badenes- Perez et al., 2013) and H. armigera were shown to avoid feeding on the mid-vein and

82 periphery of Arabidopsis rosette leaves where the concentration of the major glucosinolates is highest (Shroff et al., 2008).

Figure 4.5: Insect bioassay with H. armigera neonates fed on WT and gluc-null Arabidopsis for 9 days. (A) Mortality and (B) weight-gain observed for H. armigera neonates fed on WT and gluc-null Arabidopsis for 9 days. Data are mean ± SEM. (C) Photograph of larvae after feeding on WT and gluc-null Arabidopsis for 9 days.

In order to test this hypothesis, we compared the mortality and weight-gain of H. armigera neonates fed on either a gluc-null or WT Arabidopsis for 9 days (Fig. 4.5). Significantly higher larval mortality (~40%) was found in larvae feeding on WT plants compared to gluc-null plants (~7%) (Student’s t-test, P < 0.05; Fig. 4.5A). Larvae feeding on gluc-null plants for 9 days also attained a significantly larger size (40.3 mg) than larvae feeding on WT Arabidopsis for a similar period of time (3.4 mg) (student’s t-test, P < 0.05; Figs. 4.5B and C). Clearly, the endogenous glucosinolates in WT plants significantly increase mortality and reduce development of H. armigera neonates feeding on these plants.

83 We found that the endogenous glucosinolates content was the reason behind the high mortality of H. armigera larvae fed on WT Arabidopsis. We therefore decided to engineer the GNA/Hv1a transgenes into the gluc-null quadruple mutant Arabidopsis.

4.3.1.2 H. armigera fed leaves from Hv1a transgenics in a gluc-null background We examined survival of H. armigera neonates feeding on detached leaves from gluc-null control and gluc-null transgenic lines expressing GNA/Hv1a transgenes.

A significant difference was found between the survival of H. armigera larvae feeding on gluc-null control plants or gluc-null Arabidopsis expressing Hv1a transgenes (Fig. 4.6A, log-rank tests on Kaplan–Meier survival curves yielded P = 0.00255). This suggests that expression of insecticidal peptides in planta provided resistance against H. armigera larvae. Larval mortality increased with feeding time and the lowest survival was observed in larvae fed on leaves from plants expressing GNA-Hv1a.

No significant difference was found between the survival of H. armigera larvae fed on leaves of transgenic plants expressing Hv1a and GNA (P = 0.288), Hv1a and Hv1a-GNA (P = 0.0678) or Hv1a and GNA-Hv1a (P = 0.298) (Fig. 4.6A, log-rank tests on Kaplan-Meier survival curves). However, after 3 days of insect release, significantly lower survival was observed on GNA-Hv1a or Hv1a-GNA leaves compared to Hv1a leaves (Fig. 4.6A). This suggests that the GNA-Hv1a and Hv1a-GNA transgenes provide the best protection from H. armigera larvae.

There was no significant difference between H. armigera larval survival on GNA and Hv1a-GNA leaves (Fig. 4.6A, log-rank tests on Kaplan–Meier survival curves yielded P = 0.153). However, a significant difference was found between the survival of larvae feeding on GNA and GNA-Hv1a leaves (Fig. 4.6A, log-rank tests on Kaplan–Meier survival curves yielded P = 0.0381). Thus, in planta expressed GNA-Hv1a confers better resistance against H. armigera than Hv1a-GNA. However, no significant difference was found between survival of H. armigera larvae feeding on Hv1a-GNA and GNA-Hv1a leaves (Fig. 4.6A, log-rank tests on Kaplan–Meier survival curves yielded P = 0.57), though higher mortality was observed in case of larvae feeding on GNA-Hv1a leaves for 7 days (Fig. 4.6A).

84

Figure 4.6: Insect bioassay with H. armigera neonates feeding on leaves of gluc-null plants or transgenic plants expressing Hv1a, GNA, Hv1a-GNA or GNA-Hv1a generated in a gluc-null Arabidopsis background. (A) Kaplan-Meier survival curves for H. armigera neonates feeding on leaves of gluc-null plants or transgenic plants expressing Hv1a, GNA, Hv1a-GNA or GNA-Hv1a generated in a gluc-null Arabidopsis background. (B) Weight of larvae (mean ± SEM) after 9 days of feeding on leaves of gluc-null or transgenic Arabidopsis.

The average weight of H. armigera larvae fed on leaves of gluc-null control plants or gluc-null plants expressing GNA, Hv1a, GNA-Hv1a or Hv1a-GNA was found to be 19.28 mg, 12.65 mg, 9.87 mg, 8.93 mg and 5.81 mg, respectively (Fig. 4.6B). One-way ANOVA followed by a multiple comparisons test revealed significant differences between the average weight of H. armigera larvae fed on leaves of gluc-null plants and the Hv1a, GNA, GNA-Hv1a or Hv1a-GNA plants (Fig. 4.6B, F = 20.63, d.f. = 4, P < 0.0001). While all of the transgenes impeded larval growth, larvae gained least weight when fed on leaves of GNA-Hv1a plants (Fig. 4.6B).

85

Figure 4.7: Consumption of leaf tissue by H. armigera larvae during five days of feeding on detached leaves of gluc-null control or different transgenic plants in gluc-null background.

4.3.1.3 H. armigera leaf damage on Hv1a transgenics in gluc-null background

To determine the severity of leaf tissue damage by H. armigera larvae, neonates were fed detached leaves from gluc-null or Hv1a/GNA lines in the gluc-null background. The amount of the leaf tissue consumed after five days of feeding was higher in gluc-null control plants than gluc-null plants expressing Hv1a, GNA, Hv1a-GNA or GNA-HV1a (Fig. 4.7). One-way ANOVA revealed a significant difference between gluc-null plants and gluc-null plants expressing containing Hv1a, GNA Hv1a-GNA or GNA-HV1a transgenes (Fig. 4.7, F = 4.132, d.f. = 4, P = 0.0071). However, one-way ANOVA followed by a multiple comparisons test indicated that only the plants expressing GNA-Hv1a had significantly less leaf damage than the gluc-null control plants (Fig. 4.7).

4.3.1.4 Extended feeding assays

Feeding assays were extended to 15 days to compare the resistance conferred by the Hv1a and GNA-Hv1a transgenes over a longer period of time. We found no significant difference in the survival of H. armigera larvae fed on gluc-null plants or gluc-null plants

86 containing an Hv1a transgene (Fig. 4.8, black line versus red dashed line, log-rank test on Kaplan–Meier survival curves yielded P = 0.0635) even though there was clearly decreased survival on the gluc-null/Hv1a plants. However, a highly significant decrease in survival was found for larvae feeding on gluc-null/GNA-Hv1a leaves compared to those fed on gluc-null leaves (Fig. 4.8, black line versus green dotted line, log-rank test on Kaplan–Meier curves yielded P = 0.00000712) or gluc-null/Hv1a leaves (Fig. 4.8, red dashed line versus green dotted line, log-rank test on Kaplan–Meier curves yielded P = 0.0318). These results suggest that in planta expressed GNA-Hv1a, but not Hv1a alone, confers considerable resistance against H. armigera larvae.

Fig. 4.9 shows the severity of damage caused by H. armigera larvae fed on detached leaves of gluc-null, gluc-null/Hv1a, and gluc-null/GNA-Hv1a Arabidopsis. Larvae feeding on gluc-null leaves consumed almost all the plant tissue supplied during the assay and they attained the largest size. The larvae consumed much less tissue when fed on gluc-null/GNA-Hv1a leaves.

Figure 4.8: Kaplan-Meier survival curves for H. armigera neonates fed for 15 days with leaves from gluc-null, gluc-null/Hv1a or gluc-null/GNA-Hv1a plants.

87

Figure 4.9: Severity of leaf tissue damage caused by H. armigera larvae supplied continuously for 11 days with leaves from gluc-null plants or gluc-null plants expressing Hv1a or GNA-Hv1a.

4.3.2 Detection of insecticidal peptides in insect hemolymph

We examined the hemolymph from insects fed on each of the transgenic plants constructed in a gluc-null background to determine whether we could detect the insecticidal peptides produced by the plant transgenes. Using an anti-Hv1a antibody, we were able to detect Hv1a-GNA and GNA-Hv1a (~16 kDa) in the hemolymph of H. armigera fed for 3–7 days on plants engineered to express these fusion proteins (Fig. 4.10). A band of mass <16 kDa was also observed in western blots (Fig. 4.10, marked with arrow), which might represent a proteolytic degradation product.

Figure 4.10: Western blot analysis of hemolymph from dead larvae fed on different transgenic leaves. Proteins were detected using an anti-Hv1a polyclonal antibody (1:2000 dilution).

88 4.4 Discussion

4.4.1 Effect of endogenous glucosinolates on H. armigera survival

Initially, we generated homozygous transgenic Arabidopsis lines expressing Hv1a, GNA, Hv1a-GNA and GNA-Hv1a under 35S promoter control in a WT background to study the insecticidal effect of in planta expressed Hv1a and Hv1a/GNA fusions. Although H. armigera neonates fed on plants expressing Hv1a or Hv1a/GNA fusions appeared to have mortality than neonates fed on wild-type Arabidopsis (Fig. 4.4A), the differences were not statistically significant. However, larval growth was impaired by a statistically significant amount in neonates fed on plants expressing Hv1a or the GNA-Hv1a fusion protein (Fig. 4.4B). Thus, while these experiments provided some evidence that in planta expression of Hv1a and Hv1a/GNA fusions can provide resistance against H. armigera larvae, it was difficult to make definitive conclusions because of the high level of mortality in control, wild-type Arabidopsis.

We surmised that the high levels of larval mortality on wild-type plants might be due to expression of endogenous glucosinolates. We therefore compared the mortality of H. armigera larvae on wild-type plants and gluc-null plants deficient in production of glucosinolates. Larval mortality was much lower and weight gain was much higher for larvae fed on gluc-null plants (Fig. 4.5). Thus, we conclude that the glucosinolates present in WT Arabidopsis are responsible for high level of mortality of H. armigera larvae. Similar effects have been reported for Mamestra brassicae larvae feeding on Arabidopsis. Beekwilder et al. (2008) found that weight gain was 2.6-fold higher for larvae fed on double-mutant Arabidopsis (myb28 myb29) completely lacking aliphatic glucosinolates and 1.8-fold higher for larvae fed on single-mutant Arabidopsis with intermediate levels of aliphatic glucosinolates compared to WT plants (Beekwilder et al., 2008). Our results are consistent with the report by Shroff et al. (2008) that H. armigera larvae avoid the glucosinolate-rich mid-vein and leaf periphery when feeding on Arabidopsis leaves. Endogenous levels of glucosinolates inhibited larval growth and appeared to mask most effects of the Hv1a transgenes, suggesting that the two “defence” pathways are not synergistic.

4.4.2 in planta expression of Hv1a/GNA fusions offers resistance to H. armigera

When Hv1a and HV1a/GNA fusions were expressed in a gluc-null background, it became evident that these transgenes confer a significant level of resistance against H. armigera. This was most evident in 15-day feeding trials in which larval survival was much lower in

89 plants expressing Hv1a or GNA-Hv1a compared to gluc-null control plants (Fig. 4.8). As predicted, larval mortality was significantly higher in GNA-Hv1a plants than those expressing Hv1a alone. After 15 days, larval mortality in the GNA-Hv1a plants was >90% compared to ~50% for Hv1a plants and ~20% for gluc-null control plants (Fig. 4.8). Consistent with the effects on larval mortality, leaf damage was massively reduced in GNA-Hv1a plants compared to gluc-null control plants (Fig. 4.9). We were able to detect GNA-Hv1a in the hemolymph of larvae fed on plants expressing GNA-Hv1a (Fig. 4.10), thereby confirming that GNA is capable of ferrying Hv1a across the insect gut when in planta expressed GNA-Hv1a fusion protein is consumed by cotton bollworms.

4.4.3 Comparison with other plants engineered to express Hv1a

Hv1a has been expressed in tobacco plants (Nicotiana tabacum). When fed to H. armigera and Spodoptera littoralis larvae, these plants caused 100% mortality by 48 h (Khan et al., 2006; Shah et al., 2011). The high level of mortality reported is rather surprising as we found a much lower level of mortality for H. armigera fed on Arabidopsis expressing Hv1a alone. The same authors also reported that a recombinant thioredoxin-Hv1a fusion protein is lethal to H. armigera and S. littoralis larvae when applied topically (Khan et al., 2006). Again, this result is rather surprising; however, in these assays the fusion protein was applied topically in a solution containing high levels of imidazole, a compound known to have contact insecticidal activity (Pence, 1965), which casts doubt on the validity of their conclusions. An ER signal in their transgene constructs also seems to be missing.

Later, it has even been mentioned that Hvt-cotton expressing Hv1a is as effective as Monsanto’s pyramided Bollgard II® cotton in controlling major cotton pests (Omar and Chatha, 2012).

Recently, Bonning et al. (2014) developed transgenic Arabidopsis expressing a fusion gene consisting of the viral coat protein and a small portion of the read through domain of Pea enation mosaic virus (PEMV) to the highly insect-specific spider venom peptide Hv1a. Green peach aphid Myzus persicae feeding on transgenic Arabidopsis showed paralysis, a reduction in the population and a reduced infestation (Bonning et al., 2014).

This thesis presents the first work that tests for resistance conferred by in planta expressed GNA/Hv1a fusions against H. armigera or other lepidopteran pests.

90 4.5 Summary and conclusions

Using transgenic lines generated in a gluc-null background, we demonstrated that in planta expression of a GNA-Hv1a fusion protein provides a high level of resistance to H. armigera. Larval mortality was very high and plant damage very low in this transgenic plant. These exciting results flag GNA-Hv1a as a promising candidate bioinsecticide for insect pest control, either as an alternative to harmful chemical insecticides or for trait staking or pyramiding with Bt in transgenic crops to widen the targeted pest range and to reduce the risk of resistance development. It was recently shown that an Hv1a-GNA fusion protein has no adverse effects on honeybees (Nakasu et al., 2014), which further highlights the potential of GNA/Hv1a fusion proteins as eco-friendly bioinsecticides.

91 Chapter 5: Summary, general discussion and future directions

5.1 Summary of findings

5.1.1 Recombinant production of Hv1a/GNA fusion proteins

Recombinant Hv1a, GNA, Hv1a-GNA and GNA-Hv1a were successfully produced in soluble form using P. pastoris. All four recombinant proteins were purified to >95% using a combination of Ni-NTA affinity chromatography and RP-HPLC. The production and purification methods established in the current study will serve as a basis for further improvements in recombinant production of these proteins.

5.1.2 Insecticidal activity of recombinant Hv1a/GNA fusion proteins

Injection bioassays with houseflies (M. domestica) revealed that recombinant Hv1a-GNA and GNA-Hv1a fusion proteins retain the intrinsic insecticidal activity of Hv1a. Based on the extent of leaf damage, the Hv1a-GNA and GNA-Hv1a fusion proteins showed a higher level of insecticidal activity than Hv1a, consistent with the expectation that coupling Hv1a to GNA would increase its oral insecticidal activity.

5.1.3 Transformation vector for Agrobacterium mediated plant transformation

Vectors for Arabidopsis transformation were constructed that encoded codon-optimised genes for Hv1a, GNA, or Hv1a fused to the N- or C-terminus of either GNA or the coat protein of pea enation mosaic virus (CP). Two sets of vectors were constructed: in one set the genes were under control of the strong 35S promoter and in the other set they were under control of the phloem-specific SUC2 promoter. The transformation vectors were then introduced into Agrobacterium so that transgenes could be transformed into the plant genome.

5.1.4 Generation and characterization of transgenic Arabidopsis plants

Transgenic Arabidopsis plants were generated using the floral dip method. Integration of transgenes into the plant genome was confirmed using PCR. Genetically stable, single-locus homozygous lines were chosen by following the herbicide selectable marker. Transgene expression in transgenic plants was confirmed by western blotting with specific antibodies.

92 5.1.5 Insecticidal activity of in planta expressed toxins against H. armigera

Initial bioassays performed with H. armigera larvae (cotton bollworms) feeding on transgenic Arabidopsis plants revealed that the endogenous glucosinolates present in wild-type plants caused considerable mortality, which masked the insecticidal effect of in planta expressed insecticidal peptides. A second set of transgenic plants was therefore constructed in which the insect-toxin transgenes were expressed in a gluc-null Arabidopsis plant that is completely deficient in glucosinolates production. Bioassays with these transgenic plants revealed that Hv1a alone conferred some resistance against H. armigera larvae, but the level of resistance was considerably improved when Hv1a was fused to GNA in order to improve its oral activity. The larvae feeding on transgenic GNA-Hv1a and Hv1a-GNA plants had reduced survival; retarded growth and they also caused significantly less leaf tissue damage. The plants that expressed GNA-Hv1a showed the highest level of resistance against H. armigera.

5.2 General discussion

Chemical insecticides were the dominant method of insect pest control for the second half of the 20th century. However, the repeated use of some of these insecticides over many decades inevitably provided sufficient selection pressure that many pest insects developed resistance to these chemicals. Insecticide resistance, in concert with increased concern about the adverse effects of some chemical insecticides on the environment and human health, has led to a spate of voluntary withdrawals as well as forced de-registrations and use cancellations for many insecticides. This has greatly diminished the number of available chemical insecticides and created to an urgent need to develop new insect control methods (Windley et al., 2012; King and Hardy, 2013; Smith et al., 2013).

Recombinant DNA technology makes it possible to insert foreign DNA into plant genomes with a view to generating plants expressing desired traits such as resistance against insect pests. Insect-resistant transgenic crops, which were introduced in the late 1990s, have been the most successful application of plant genetic engineering technology to agriculture (Gatehouse, 2008). The first generation of genetically modified insect-resistant crops contained a transgene encoding an insecticidal toxin from the soil bacterium Bacillus thuringiensis (so-called Bt crops). However, from the 1990s, an enormous amount of work has been carried out to develop transgenic plants that express putative insecticidal genes from natural sources such as bacteria, plants, and animal venoms (Tian et al., 1991; Yao et al., 1996; Anderson et al., 1997; Rao et al., 1998)

93 Although plant-derived insecticidal compounds such as lectins might have a potential advantage over toxins derived from venoms or microbes in terms of public acceptance, they are much less potent than neurotoxic venom peptides (Fitches et al., 1997) and thus have failed to enter the commercial insect pest control arena. Venom-derived peptides such as those from scorpions (Rodriguez de la Vega and Possani, 2005; Gurevitz et al., 2007) and spiders (Tedford et al., 2004; Windley et al., 2012; King and Hardy, 2013; Smith et al., 2013) are highly potent insecticidal toxins that are effective at very low doses, and thus they have greater potential to be utilized in commercial insect pest control strategies. However, the phyletic specificity of these toxins is critical; in order for them to be viable for insect pest control, they must be devoid of vertebrate activity (Wang et al., 2000).

Many of the insecticidal toxins derived from arthropod venoms are active against neuronal ion channels that are not targeted by current chemical insecticides (Tedford et al., 2004; Smith et al., 2013). For example, Hv1a, the subject of this thesis, targets voltage-gated calcium channels. However, these toxins need to access the nervous system of insects to reach their sites of action in order to be effective (Fitches et al., 2012). In nature, venomous animals inject these neurotoxins into the insect hemolymph from where they can access the nervous system. However, if used in crop protection, venom peptides would need to pass through the insect cuticle if applied topically or cross the insect gut epithelium if administered orally (Fitches et al., 2012; King and Hardy, 2013). One of the most promising options is to fuse these toxins to proteins that can translocate across the insect gut, epithelium such as plant lectins (Fitches et al., 2012) or virus coat proteins (Bonning et al., 2014).

GNA has been used to improved the oral activity of a variety of insecticidal neurotoxins (Fitches et al., 1997; Fitches et al., 2001; Fitches et al., 2002; Fitches et al., 2004; Down et al., 2006; Trung et al., 2006; Wakefield et al., 2006; Fitches et al., 2010; Wakefield et al., 2010; Fitches et al., 2012). GNA and CP both bind to receptors in the insect gut and subsequently cross the epithelium by transcytosis (Fitches et al., 1997; Fitches et al., 2001; Fitches et al., 2012; Bonning and Chougule, 2014; Bonning et al., 2014) .

Stability within the insect gut environment is a major factor when considering fusion protein toxicity. However, it is also important to consider the stability of the fusion proteins within the insect hemolymph. To be successfully toxic the fusion protein needs to be able to withstand gut proteolysis but after transport into the hemolymph it might be important for the insecticidal toxin to be released from the carrier protein. If a fusion protein is too

94 stable when transported into the hemolymph, its toxin component might not be able to interact with its molecular target in the nervous system.

In this study, the Arabidopsis plant expression system was used to express GNA/Hv1a fusion proteins in planta. Although expression in P. pastoris yielded relatively high levels of recombinant GNA/Hv1a fusion proteins, which could potentially be used as exogenously applied insecticides, expression of the fusion proteins in planta offers the possibility of producing endogenous resistance to insect pests in crops. Expression in planta allows for a more direct approach to season-long control of lepidopteran pest species, particularly where expression of insecticidal proteins can offer direct protection to parts of crop plants, such as the cotton boll, which are difficult to protect with exogenously applied insecticides.

In order to be a commercially successful insect control agent, the in planta expressed GNA/Hv1a fusion proteins would have to show a comparable level of resistance against insect pests to the currently approved Bt crops. However, it should be noted that despite their success in protecting important crops like cotton and corn from lepidopteran pests, Bt toxins are only effective against a small range of insects (lepidopterans, coleopterans, and dipterans) (Sharma et al., 2004). This deficiency has been addressed by stacking Bt plants with insecticidal transgenes having different modes of action. For example, in planta expression of Bt toxins plus GNA yielded increased toxicity and also increased the range of susceptible insect species, with the expanded range including hemipterans (Maqbool et al., 2001; Ramesh et al., 2004).

In addition to increasing toxicity and expanding the range of susceptible insects, pyramiding insecticidal transgenes can avoid or delay the development of resistance. An example of a such as transgenic plant is Bollgard II cotton (Monsanto), which contains two transgenes encoding the Bt toxins Cry1Ac and Cry2Ab, where Cry1Ac is specific to lepidopterans and Cry2Ab is more broadly active against dipterans and lepidopterans. An extreme example of trait stacking is the insect-resistant, herbicide-tolerant Genuity® strains of transgenic corn (Monsanto/Dow collaboration), which contain as many as eight transgenes, including multiple insect resistance genes and two transgenes conferring tolerance to glyphosate and glufosinate-ammonium herbicides (Monsanto, 2014).

Most of the insecticidal peptide toxins explored to date have not been shown to target specific insect orders. But there are still a large number of insect-specific toxins to be explored that have the potential to also be order-specific (Escoubas et al., 2006). An alternative way of limiting the host range of toxins expressed in planta is to limit their

95 tissue expression. For example, one could place the transgene under the control of a phloem-specific promoter in order to limit the effects of the toxins to sap sucking insects (Sauer and Stolz, 1994; Stadler and Sauer, 1996; Juergensen et al., 2003; Dinant et al., 2004; Dutt et al., 2012). Alternatively, one could fuse the toxin to a carrier protein that is only recognised by specific insects; for example, fusion to luteovirus coat proteins can be used to specifically target sap-sucking aphids which act as vectors of these plant viruses (Bonning et al., 2014).

5.3 Future work

In this project, we generated transgenic Arabidopsis plants that express Hv1a alone, or fused to GNA or CP, under control of either the constitutive 35S promoter or the phloem-specific SUC2 promoter. We demonstrated that transgenes encoding the Hv1a/GNA fusion proteins conferred a high level of resistance against H. armigera when expressed using the strong 35S promoter, which is not only constitutive but also active in all plant tissues (Odell et al., 1985; Benfey and Chua, 1990; de Mesa et al., 2004). The fusions of Hv1a with CP were designed to target sap-sucking aphids but there was insufficient time to test these plants. Collaborations are ongoing to test whether expression of the Hv1a/CP fusions in Arabidopsis confers resistance against the green peach aphid, Myzus persicae.

In addition, the time available did not permit testing of any of the transgenes expressed under SUC2 control, which limits transgene expression to the phloem and therefore has the potential to limit affects on non-target insects, particularly beneficial insects such as pollinators and natural insect predators. Plans are underway to test these plants against a range of sap sucking and lepidopteran pests, with the anticipation that only sap-sucking insects will be affected. In future studies it will also be important to study the phenotype and agronomic parameters of each of the transgenic plants in order to determine whether there is any fitness cost associated with expression of the insecticidal toxin transgenes.

In future studies, it will be critical to examine how well the Hv1a transgenes perform in crop plants. While Arabidopsis is a valuable model system, the real-life performance of the GNA/Hv1a and CP/Hv1a transgenes expressed in crop plants such as corn and cotton will be critical if they are to be commercially viable. Moreover, it will be extremely interesting to see how well the Hv1a transgenes perform when stacked with Bt transgenes. Since Cry toxins damage the integrity of the insect gut, they might enhance the activity of Hv1a transgenes by providing the spider toxin with greater access to the hemolymph. Hence,

96 stacking of Hv1a and Bt transgenes might lead to a synergistic increase in insecticidal activity.

Finally, there are questions that remained to be answered with regard to safety and public acceptance. Hv1a has no vertebrate activity in any species that have been examined thus far (rats, mice, and rabbits) but more widespread testing is required against ecologically important vertebrate species such as birds and , as well as non-target beneficial insects. It also remains to be seen how well the public will react to GM crops expressing a spider toxin, although this concern is somewhat ameliorated by the acceptance of Bt, which was initially accepted in non-food crops like cotton, but now GM food crops like Bt-corns (Agrisure® Duracade™; Herculex™ RW; YieldGard™ etc.) and Bt-soybean (Intacta™ Roundup Ready™ 2 Pro) are being cultivated in Argentina, Brazil, Canada, China, Chile, Colombia, European Union, Mexico, Paraguay, Philippines, South Africa, Taiwan, Uruguay and USA with country specific approval (ISAAA, 2014).

97 References

Abe, T.; Sugita, M.; Fujikura, T.; Hiyoshi, J. & Akasu, M. 2000. Giant hornet (Vespa mandarinia) venomous phospholipases. The purification, characterization and inhibitory properties by biscoclaurine alkaloids. Toxicon, 38, 1803–1816.

Ahuja, I.; Rohloff, J. & Bones, A. 2010. Defence mechanisms of Brassicaceae: implications for plant-insect interactions and potential for integrated pest management. Agron. Sustain. Dev., 30, 311–348.

Alfthan, K.; Takkinen, K.; Sizmann, D.; Soderlund, H. & Teeri, T. T. 1995. Properties of a single-chain antibody containing different linker peptides. Protein Eng., 8, 725– 731.

Ali, M. I. & Luttrell, R. G. 2007. Susceptibility of bollworm and tobacco budworm (Lepidoptera: Noctuidae) to Cry2Ab2 insecticidal protein. J. Econ. Entomol., 100, 921–931.

Ali, M. I.; Luttrell, R. G. & Young, S. Y., 3rd 2006. Susceptibilities of Helicoverpa zea and Heliothis virescens (Lepidoptera: Noctuidae) populations to Cry1Ac insecticidal protein. J. Econ. Entomol., 99, 164–175.

Ali, S. A.; Stoeva, S.; Grossmann, J. G.; Abbasi, A. & Voelter, W. 2001. Purification, characterization, and primary structure of four depressant insect-selective neurotoxin analogs from scorpion (Buthus sindicus) venom. Arch. Biochem. Biophys., 391, 197–206.

An, Y. Q.; McDowell, J. M.; Huang, S. R.; McKinney, E. C.; Chambliss, S. & Meagher, R. B. 1996. Strong, constitutive expression of the Arabidopsis ACT2/ACT8 actin subclass in vegetative tissues. Plant J., 10, 107–121.

Anderson, M. A.; Van Heeswijck, R.; West, J.; Bateman, K.; Lee, M.; Christeller, J. T.; McDonald, G. & Heath, R. L. 1997. Proteinase inhibitors from Nicotiana alata enhance plant resistance to insect pests. J. Insect Physiol., 43, 833–842.

Arai, R.; Ueda, H.; Kitayama, A.; Kamiya, N. & Nagamune, T. 2001. Design of the linkers which effectively separate domains of a bifunctional fusion protein. Protein Eng., 14, 529–532.

Argos, P. 1990. An investigation of oligopeptides linking domains in protein tertiary structures and possible candidates for general gene fusion. J. Mol. Biol., 211, 943– 958.

Arnon, T.; Potikha, T.; Sher, D.; Elazar, M.; Mao, W.; Tal, T.; Bosmans, F.; Tytgat, J.; Ben- Arie, N. & Zlotkin, E. 2005. BjalphaIT: a novel scorpion alpha-toxin selective for insects-unique pharmacological tool. Insect Biochem. Mol. Biol., 35, 187–195.

Audsley, N.; Matthews, J.; Nachman, R. J. & Weaver, R. J. 2008. Transepithelial flux of an allatostatin and analogs across the anterior midgut of Manduca sexta larvae in vitro. Peptides, 29, 286–294.

Badenes-Perez, F. R.; Reichelt, M.; Gershenzon, J. & Heckel, D. G. 2013. Interaction of glucosinolate content of Arabidopsis thaliana mutant lines and feeding and oviposition by generalist and specialist lepidopterans. Phytochemistry, 86, 36–43.

98 Baptista-Saidemberg, N. B.; Saidemberg, D. M. & Palma, M. S. 2011. Profiling the peptidome of the venom from the social wasp Agelaia pallipes pallipes. J. Proteomics, 74, 2123–2137.

Barton, K. A. & Miller, M. J. 1993. Insecticidal toxins in plants. United States Patent, U. S. Patent No. 5177308 A.

Bebber, D. P.; Ramotowski, M. A. T. & Gurr, S. J. 2013. Crop pests and pathogens move polewards in a warming world. Nature Clim. Change, 3, 985–988.

Bechtold, N.; Ellis, J. & Pelletier, G. 1993. In planta Agrobacterium-mediated gene transfer by infiltration of adult Arabidopsis thaliana plants. Comp. Rend. L’Acad. des Sci. , Serie III 1194–1199.

Bechtold, N.; Jaudeau, B.; Jolivet, S.; Maba, B.; Vezon, D.; Voisin, R. & Pelletier, G. 2000. The maternal chromosome set is the target of the T-DNA in the in planta transformation of Arabidopsis thaliana. Genetics, 155, 1875–1887.

Bechtold, N. & Pelletier, G. 1998. In planta Agrobacterium-mediated transformation of adult Arabidopsis thaliana plants by vacuum infiltration. Methods Mol. Biol., 82, 259–266.

Beekwilder, J.; van Leeuwen, W.; van Dam, N. M.; Bertossi, M.; Grandi, V.; Mizzi, L.; Soloviev, M.; Szabados, L.; Molthoff, J. W.; Schipper, B.; Verbocht, H.; de Vos, R. C.; Morandini, P.; Aarts, M. G. & Bovy, A. 2008. The impact of the absence of aliphatic glucosinolates on insect herbivory in Arabidopsis. PLoS One, 3, e2068.

Bende, N. S.; Kang, E.; Herzig, V.; Bosmans, F.; Nicholson, G. M.; Mobli, M. & King, G. F. 2013. The insecticidal neurotoxin Aps III is an atypical knottin peptide that potently blocks insect voltage-gated sodium channels. Biochem. Pharmacol., 85, 1542– 1554.

Benfey, P. N. & Chua, N. H. 1990. The Cauliflower Mosaic Virus 35S promoter: combinatorial regulation of transcription in plants. Science, 250, 959–966.

Bent, A. 2006. Arabidopsis thaliana floral dip transformation method. In: Wang, K. (ed.) Agrobacterium Protocols. Humana Press.

Bevan, M. W.; Flavell, R. B. & Chilton, M. D. 1983. A chimeric antibiotic-resistance gene as a selectable marker for plant-cell transformation. Nature, 304, 184–187.

Bhalla, P. L. & Singh, M. B. 2008. Agrobacterium-mediated transformation of Brassica napus and Brassica oleracea. Nat. Protoc., 3, 181–189.

Bohinc, T.; Ban, S. G.; Ban, D. & Trdan, S. 2012. Glucosinolates in plant protection strategies: a review. Arch. Biol. Sci., 64, 821–828.

Bonning, B. C. & Chougule, N. P. 2014. Delivery of intrahemocoelic peptides for insect pest management. Trends Biotechnol., 32, 91–98.

Bonning, B. C.; Pal, N.; Liu, S.; Wang, Z.; Sivakumar, S.; Dixon, P. M.; King, G. F. & Miller, W. A. 2014. Toxin delivery by the coat protein of an aphid-vectored plant virus provides plant resistance to aphids. Nat. Biotechnol., 32, 102–105.

99 Borchani, L.; Mansuelle, P.; Stankiewicz, M.; Grolleau, F.; Cestele, S.; Karoui, H.; Lapied, B.; Rochat, H.; Pelhate, M. & el Ayeb, M. 1996. A new scorpion venom toxin paralytic to insects that affects Na+ channel activation. Purification, structure, antigenicity and mode of action. Eur. J. Biochem., 241, 525–532.

Borchani, L.; Stankiewicz, M.; Kopeyan, C.; Mansuelle, P.; Kharrat, R.; Cestele, S.; Karoui, H.; Rochat, H.; Pelhate, M. & el Ayeb, M. 1997. Purification, structure and activity of three insect toxins from Buthus occitanus tunetanus venom. Toxicon, 35, 365–382.

Boulter, D.; Gatehouse, J. A.; Gatehouse, A. M. R. & Hilder, V. A. 1990. Genetic- engineering of plants for insect resistance. Endeavour, 14, 185–190.

Bravo, A.; Gill, S. S. & Soberon, M. 2007. Mode of action of Bacillus thuringiensis Cry and Cyt toxins and their potential for insect control. Toxicon, 49, 423–435.

Bravo, A.; Likitvivatanavong, S.; Gill, S. S. & Soberon, M. 2011. Bacillus thuringiensis: A story of a successful bioinsecticide. Insect Biochem. Mol. Biol., 41, 423–431.

Brigatte, P.; Cury, Y.; de Souza, B. M.; Baptista-Saidemberg, N. B.; Saidemberg, D. M.; Gutierrez, V. P. & Palma, M. S. 2011. Hyperalgesic and edematogenic effects of peptides isolated from the venoms of honeybee (Apis mellifera) and neotropical social wasps (Polybia paulista and Protonectarina sylveirae). Amino Acids, 40, 101–111.

Brogdon, W. G. & McAllister, J. C. 1998. Insecticide resistance and vector control. Emerg. Infect. Dis., 4, 605–613.

Broothaerts, W.; Mitchell, H. J.; Weir, B.; Kaines, S.; Smith, L. M.; Yang, W.; Mayer, J. E.; Roa-Rodriguez, C. & Jefferson, R. A. 2005. Gene transfer to plants by diverse species of bacteria. Nature, 433, 629–633.

Cai, M.; Wei, J.; Li, X.; Xu, C. & Wang, S. 2007. A rice promoter containing both novel positive and negative cis-elements for regulation of green tissue-specific gene expression in transgenic plants. Plant Biotechnol. J., 5, 664–674.

Cao, C. W.; Liu, G. F.; Wang, Z. Y.; Yan, S. C.; Ma, L. & Yang, C. P. 2010. Response of the gypsy moth, Lymantria dispar to transgenic poplar, Populus simonii x P. nigra, expressing fusion protein gene of the spider insecticidal peptide and Bt-toxin C- peptide. J. Insect. Sci., 10, 200.

Casartelli, M.; Corti, P.; Giovanna Leonardi, M.; Fiandra, L.; Burlini, N.; Pennacchio, F. & Giordana, B. 2005. Absorption of albumin by the midgut of a lepidopteran larva. J. Insect. Physiol., 51, 933–940.

Cascone, O.; Amaral, V.; Ferrara, P.; Vita, N.; Guillemot, J. C. & Diaz, L. E. 1995. Purification and characterization of two forms of antigen 5 from polybia scutellaris venom. Toxicon, 33, 659–665.

Cestele, S.; Borchani, L.; El Ayeb, M. & Rochat, H. 1997. Bot IT2: a new scorpion toxin to study receptor site on insect sodium channels. FEBS Lett., 405, 77–80.

Cestele, S.; Stankiewicz, M.; Mansuelle, P.; De Waard, M.; Dargent, B.; Gilles, N.; Pelhate, M.; Rochat, H.; Martin-Eauclaire, M. F. & Gordon, D. 1999. Scorpion alpha-like toxins, toxic to both mammals and insects, differentially interact with

100 receptor site 3 on voltage-gated sodium channels in mammals and insects. Eur. J. Neurosci., 11, 975–985.

Chang, S. S.; Park, S. K.; Kim, B. C.; Kang, B. J.; Kim, D. U. & Nam, H. G. 1994. Stable genetic transformation of Arabidopsis thaliana by Agrobacterium inoculation in planta. Plant J., 5, 551–558.

Chilton, M. D.; Drummond, M. H.; Merio, D. J.; Sciaky, D.; Montoya, A. L.; Gordon, M. P. & Nester, E. W. 1977. Stable incorporation of plasmid DNA into higher plant cells: the molecular basis of crown gall tumorigenesis. Cell, 11, 263–271.

Chitkowski, R. L.; Turnipseed, S. G.; Sullivan, M. J. & Bridges, W. C., Jr. 2003. Field and laboratory evaluations of transgenic cottons expressing one or two Bacillus thuringiensis var. kurstaki Berliner proteins for management of noctuid (Lepidoptera) pests. J. Econ. Entomol., 96, 755–762.

Chong, Y.; Hayes, J. L.; Sollod, B.; Wen, S. P.; Wilson, D. T.; Hains, P. G.; Hodgson, W. C.; Broady, K. W.; King, G. F. & Nicholson, G. M. 2007. The ω-atracotoxins: Selective blockers of insect M-LVA and HVA calcium channels. Biochem. Pharmacol., 74, 623–638.

Chowell, G.; Diaz-Duenas, P.; Bustos-Saldana, R.; Mireles, A. A. & Fet, V. 2006. Epidemiological and clinical characteristics of scorpionism in Colima, Mexico (2000-2001). Toxicon, 47, 753–758.

Clough, S. J. & Bent, A. F. 1998. Floral dip: a simplified method for Agrobacterium- mediated transformation of Arabidopsis thaliana. Plant J., 16, 735–743.

Coddington, J. A. & Levi, H. W. 1991. Systemics and evolution of spiders (Araneae). Annu. Rev. Ecol. Syst., 22, 565–592.

Corzo, G.; Bernard, C.; Clement, H.; Villegas, E.; Bosmans, F.; Tytgat, J.; Possani, L. D.; Darbon, H. & Alagon, A. 2009. Insecticidal peptides from the theraposid spider Brachypelma albiceps: an NMR-based model of Ba2. Biochim. Biophys. Acta., 1794, 1190–1196.

Corzo, G.; Escoubas, P.; Stankiewicz, M.; Pelhate, M.; Kristensen, C. P. & Nakajima, T. 2000. Isolation, synthesis and pharmacological characterization of delta-palutoxins IT, novel insecticidal toxins from the spider Paracoelotes luctuosus (Amaurobiidae). Eur. J. Biochem., 267, 5783–5795.

Corzo, G.; Escoubas, P.; Villegas, E.; Karbat, I.; Gordon, D.; Gurevitz, M.; Nakajima, T. & Gilles, N. 2005. A spider toxin that induces a typical effect of scorpion α-toxins but competes with β-toxins on binding to insect sodium channels. Biochem., 44, 1542– 1549.

Corzo, G.; Gilles, N.; Satake, H.; Villegas, E.; Dai, L.; Nakajima, T. & Haupt, J. 2003. Distinct primary structures of the major peptide toxins from the venom of the spider Macrothele gigas that bind to sites 3 and 4 in the sodium channel. FEBS Lett., 547, 43–50.

Costa, H. & Palma, M. S. 2000. Agelotoxin: a phospholipase A(2) from the venom of the neotropical social wasp cassununga (Agelaia pallipes pallipes) (Hymenoptera- Vespidae). Toxicon, 38, 1367–1379.

101 Cregg, J.; Madden, K.; Barringer, K.; Thill, G. & Stillman, C. 1989. Functional characterization of the two alcohol oxidase genes from the yeast Pichia pastoris. Mol. Cell. Biol., 9, 1316–1323.

Cregg, J. M.; Barringer, K.; Hessler, A. & Madden, K. 1985. Pichia pastoris as a host system for transformations. Mol. Cell. Biol., 5, 3376–3385.

Cregg, J. M.; Cereghino, J. L.; Shi, J. Y. & Higgins, D. R. 2000. Recombinant protein expression in Pichia pastoris. Mol. Biotechnol., 16, 23–52.

Cregg, J. M.; Tolstorukov, I.; Kusari, A.; Sunga, J.; Madden, K. & Chappell, T. 2009. Expression in the yeast Pichia pastoris. Methods enzymol., 463, 169.

Curtis, I. S. 2005. Production of transgenic crops by the floral-dip method. Methods Mol. Biol., 286, 103–110.

Curtis, I. S. & Nam, H. G. 2001. Transgenic radish (Raphanus sativus L. longipinnatus Bailey) by floral-dip method–plant development and surfactant are important in optimizing transformation efficiency. Transgenic Res., 10, 363–371.

DAFF. 2011. Helicoverpa and insecticide resistance [Online]. Queensland, Australia: Department of Agriculture, Fisheries and Forestry, Queensland Government, Australia. Available: http://www.daff.qld.gov.au/plants/field-crops-and- pastures/broadacre-field-crops/integrated-pest-management/a-z-insect-pest- list/helicoverpa/insecticide-resistance.

Dalgaard, P. 2008. Introductory statistics with R, New York, USA, Springer.

Daly, R. & Hearn, M. T. W. 2005. Expression of heterologous proteins in Pichia pastoris: a useful experimental tool in protein engineering and production. J. Mol. Recognit., 18, 119–138. de Figueiredo, S. G.; de Lima, M. E.; Nascimento Cordeiro, M.; Diniz, C. R.; Patten, D.; Halliwell, R. F.; Gilroy, J. & Richardson, M. 2001. Purification and amino acid sequence of a highly insecticidal toxin from the venom of the brazilian spider Phoneutria nigriventer which inhibits NMDA-evoked currents in rat hippocampal neurones. Toxicon, 39, 309—317.

De Lima, M. E.; Martin, M. F.; Diniz, C. R. & Rochat, H. 1986. Tityus serrulatus toxin VII bears pharmacological properties of both β-toxin and insect toxin from scorpion venoms. Biochem. Biophys. Res. Commun., 139, 296–302. de Mesa, M. C.; Santiago-Doménech, N.; Pliego-Alfaro, F.; Quesada, M. A. & Mercado, J. A. 2004. The CaMV 35S promoter is highly active on floral organs and pollen of transgenic strawberry plants. Plant Cell Rep., 23, 32–38.

Dehesa-Davila, M.; Martin, B. M.; Nobile, M.; Prestipino, G. & Possani, L. D. 1994. Isolation of a toxin from Centruroides infamatus infamatus Koch scorpion venom that modifies Na+ permeability on chick dorsal root ganglion cells. Toxicon, 32, 1487–1493.

Deng, M.; Luo, X.; Meng, E.; Xiao, Y. & Liang, S. 2008. Inhibition of insect calcium channels by -V, a neurotoxin from Chinese Ornithoctonus huwena venom. Eur. J. Pharmacol., 582, 12–16.

102 Devos, Y.; Meihls, L. N.; Kiss, J. & Hibbard, B. E. 2013. Resistance evolution to the first generation of genetically modified Diabrotica-active Bt-maize events by western corn rootworm: management and monitoring considerations. Transgenic Res., 22, 269–299.

Dhurua, S. & Gujar, G. T. 2011. Field-evolved resistance to Bt toxin Cry1Ac in the pink bollworm, Pectinophora gossypiella (Saunders) (Lepidoptera: Gelechiidae), from India. Pest Manag. Sci., 67, 898–903.

Dibden, J.; Gibbs, D. & Cocklin, C. 2013. Framing GM crops as a food security solution. J. Rural Stud., 29, 59–70.

Dinant, S.; Ripoll, C.; Pieper, M. & David, C. 2004. Phloem specific expression driven by wheat dwarf geminivirus V-sense promoter in transgenic dicotyledonous species. Physiol. Plant, 121, 108–116.

DiRita, V. J. & Gelvin, S. B. 1987. Deletion analysis of the mannopine synthase gene promoter in sunflower crown gall tumors and Agrobacterium tumefaciens. Mol. Gen. Genet., 207, 233–241.

Down, R. E.; Fitches, E. C.; Wiles, D. P.; Corti, P.; Bell, H. A.; Gatehouse, J. A. & Edwards, J. P. 2006. Insecticidal spider venom toxin fused to snowdrop lectin is toxic to the peach-potato aphid, Myzus persicae (Hemiptera: Aphididae) and the rice brown planthopper, Nilaparvata lugens (Hemiptera: Delphacidae). Pest Manag. Sci., 62, 77–85.

Dutt, M.; Ananthakrishnan, G.; Jaromin, M. K.; Brlansky, R. H. & Grosser, J. W. 2012. Evaluation of four phloem-specific promoters in vegetative tissues of transgenic citrus plants. Tree Physiol., 32, 83–93.

Edwards, K.; Johnstone, C. & Thompson, C. 1991. A simple and rapid method for the preparation of plant genomic DNA for PCR analysis. Nucleic. Acids. Res., 19, 1349.

Eisemann, C. H.; Donaldson, R. A.; Pearson, R. D.; Cadogan, L. C.; Vuocolo, T. & Tellam, R. L. 1994. Larvicidal activity of lectins on Lucilia cuprina: mechanism of action. Entomol. Exp. Appl., 72, 1–10.

Ellis, S. B.; Brust, P. F.; Koutz, P. J.; Waters, A.; Harpold, M. M. & Gingeras, T. R. 1985. Isolation of alcohol oxidase and two other methanol regulatable genes from the yeast Pichia pastoris. Mol. Cell. Biol., 5, 1111–1121.

Escoubas, P.; Diochot, S. & Corzo, G. 2000a. Structure and pharmacology of spider venom neurotoxins. Biochimie., 82, 893–907.

Escoubas, P.; Sollod, B. & King, G. F. 2006. Venom landscapes: mining the complexity of spider venoms via a combined cDNA and mass spectrometric approach. Toxicon, 47, 650–663.

Escoubas, P.; Stankiewicz, M.; Takaoka, T.; Pelhate, M.; Romi-Lebrun, R.; Wu, F. Q. & Nakajima, T. 2000b. Sequence and electrophysiological characterization of two insect-selective excitatory toxins from the venom of the Chinese scorpion Buthus martensi. FEBS Lett., 483, 175–180.

103 Estrada, G.; Villegas, E. & Corzo, G. 2007. Spider venoms: a rich source of acylpolyamines and peptides as new leads for CNS drugs. Nat. Prod. Rep., 24, 145–161.

Fahey, J. W.; Zalcmann, A. T. & Talalay, P. 2001. The chemical diversity and distribution of glucosinolates and isothiocyanates among plants. Phytochemistry, 56, 5–51.

FAO 2011. Save and Grow: a policymaker’s guide to the sustainable intensification of smallholder crop production. Rome: Food and Agricultural Organization of the United Nations. .

Feldmann, K. A. & David Marks, M. 1987. Agrobacterium-mediated transformation of germinating seeds of Arabidopsis thaliana : A non-tissue culture approach. Mol. Gen. Genet., 208, 1–9.

Fernandes-Pedrosa, M. F.; Félix-Silva, J. & Menezes, Y. A. S. 2013. Toxins from venomous animals: gene cloning, protein expression and biotechnological applications. In: Baptista, G. R. (ed.) An Integrated View of the Molecular Recognition and Toxinology - From Analytical Procedures to Biomedical Applications. InTech.

Feyereisen, R. 1995. Molecular biology of insecticide resistance. Toxicol. Lett., 82–83, 83–90.

Fiandra, L.; Casartelli, M.; Cermenati, G.; Burlini, N. & Giordana, B. 2009. The intestinal barrier in lepidopteran larvae: permeability of the peritrophic membrane and of the midgut epithelium to two biologically active peptides. J. Insect Physiol., 55, 10–18.

Figueiredo, S. G.; Garcia, M. E.; Valentim, A. C.; Cordeiro, M. N.; Diniz, C. R. & Richardson, M. 1995. Purification and amino acid sequence of the insecticidal neurotoxin Tx4(6-1) from the venom of the 'armed' spider Phoneutria nigriventer (Keys). Toxicon, 33, 83–93.

Fitches, E.; Audsley, N.; Gatehouse, J. A. & Edwards, J. P. 2002. Fusion proteins containing neuropeptides as novel insect contol agents: snowdrop lectin delivers fused allatostatin to insect haemolymph following oral ingestion. Insect Biochem. Mol. Biol., 32, 1653–1661.

Fitches, E.; Edwards, M. G.; Mee, C.; Grishin, E.; Gatehouse, A. M. R.; Edwards, J. P. & Gatehouse, J. A. 2004. Fusion proteins containing insect-specific toxins as pest control agents: snowdrop lectin delivers fused insecticidal spider venom toxin to insect haemolymph following oral ingestion. J. Insect Physiol., 50, 61–71.

Fitches, E.; Gatehouse, A. M. R. & Gatehouse, J. A. 1997. Effects of snowdrop lectin (GNA) delivered via artificial diet and transgenic plants on the development of tomato moth (Lacanobia oleracea) larvae in laboratory and glasshouse trials. J. Insect Physiol., 43, 727–739.

Fitches, E.; Woodhouse, S. D.; Edwards, J. P. & Gatehouse, J. A. 2001. In vitro and in vivo binding of snowdrop (Galanthus nivalis agglutinin; GNA) and jackbean (Canavalia ensiformis; Con A) lectins within tomato moth (Lacanobia oleracea) larvae; mechanisms of insecticidal action. J. Insect Physiol., 47, 777–787.

Fitches, E. C.; Bell, H. A.; Powell, M. E.; Back, E.; Sargiotti, C.; Weaver, R. J. & Gatehouse, J. A. 2010. Insecticidal activity of scorpion toxin (ButaIT) and

104 snowdrop lectin (GNA) containing fusion proteins towards pest species of different orders. Pest Manag. Sci., 66, 74–83.

Fitches, E. C.; Pyati, P.; King, G. F. & Gatehouse, J. A. 2012. Fusion to snowdrop lectin magnifies the oral activity of insecticidal ω-hexatoxin-Hv1a peptide by enabling its delivery to the central nervous system. PLoS One, 7, e39389.

Fitt, G. P. 1989. The ecology of Heliothis species in relation to agroecosystems. Annu. Rev. Entomol., 34, 17–53.

Fitt, G. P. 1994. Cotton pest management: Part 3. An Australian perspective. Annu. Rev. Entomol., 39, 543–562.

Fletcher, J. I.; Smith, R.; Odonoghue, S. I.; Nilges, M.; Connor, M.; Howden, M. E. H.; Christie, M. J. & King, G. F. 1997. The structure of a novel insecticidal neurotoxin, ω-atracotoxin-HV1, from the venom of an Australian funnel web spider. Nat. Struc. Biol., 4, 559–566.

Fraley, R. T.; Rogers, S. G.; Horsch, R. B.; Sanders, P. R.; Flick, J. S.; Adams, S. P.; Bittner, M. L.; Brand, L. A.; Fink, C. L.; Fry, J. S.; Galluppi, G. R.; Goldberg, S. B.; Hoffmann, N. L. & Woo, S. C. 1983. Expression of bacterial genes in plant cells. Proc. Natl. Acad. Sci. USA, 80, 4803–4807.

Froy, O. & Gurevitz, M. 2003. New insight on scorpion divergence inferred from comparative analysis of toxin structure, pharmacology and distribution. Toxicon, 42, 549–555.

Froy, O.; Zilberberg, N.; Gordon, D.; Turkov, M.; Gilles, N.; Stankiewicz, M.; Pelhate, M.; Loret, E.; Oren, D. A.; Shaanan, B. & Gurevitz, M. 1999. The putative bioactive surface of insect-selective scorpion excitatory neurotoxins. J. Biol. Chem., 274, 5769–5776.

Gassmann, A. J.; Petzold-Maxwell, J. L.; Keweshan, R. S. & Dunbar, M. W. 2011. Field- evolved resistance to Bt maize by western corn rootworm. PLoS One, 6, e22629.

Gatehouse, A. M. R. 1998. Biotechnological applications of plant genes in the production of insect-resistant crops. Global plant genetic resources for insect-resistant crops. CRC Press.

Gatehouse, A. M. R.; Ferry, N.; Edwards, M. G. & Bell, H. A. 2011. Insect-resistant biotech crops and their impacts on beneficial arthropods. Philos. T. Roy. Soc. B, 366, 1438–1452.

Gatehouse, J. A. 2008. Biotechnological prospects for engineering insect-resistant plants. Plant Physiol., 146, 881–887.

Gelvin, S. B. 2003. Agrobacterium-mediated plant transformation: the biology behind the "Gene-Jockeying" tool. Microbiol. Mol. Biol. Rev., 67, 16–37.

Gordon, D.; Martin-Eauclaire, M. F.; Cestele, S.; Kopeyan, C.; Carlier, E.; Khalifa, R. B.; Pelhate, M. & Rochat, H. 1996. Scorpion toxins affecting sodium current inactivation bind to distinct homologous receptor sites on rat brain and insect sodium channels. J. Biol. Chem., 271, 8034–8045.

105 Goudet, C.; Chi, C. W. & Tytgat, J. 2002. An overview of toxins and genes from the venom of the Asian scorpion Buthus martensi Karsch. Toxicon, 40, 1239–1258.

Gould, F. 2003. Bt-resistance management–theory meets data. Nat. Biotechnol., 21, 1450–1451.

Gray, S. & Gildow, F. E. 2003. Luteovirus-aphid interactions. Ann. Rev. Phytopath., 41, 539–566.

Greenop, K. R.; Peters, S.; Bailey, H. D.; Fritschi, L.; Attia, J.; Scott, R. J.; Glass, D. C.; de Klerk, N. H.; Alvaro, F.; Armstrong, B. K. & Milne, E. 2013. Exposure to pesticides and the risk of childhood brain tumors. Cancer Cause Control, 24, 1269–1278.

Grube, A.; Donaldson, D.; Kiely, T. & Wu, L. 2011. Pesticides industry sales and usage: 2006 and 2007 market estimates. U.S. Environmental Protection Agency, Washington, DC 20460.

Gunning, S. J.; Maggio, F.; Windley, M. J.; Valenzuela, S. M.; King, G. F. & Nicholson, G. M. 2008. The Janus-faced atracotoxins are specific blockers of invertebrate KCa channels. FEBS J., 275, 4045–4059.

Gurevitz, M.; Karbat, I.; Cohen, L.; Ilan, N.; Kahn, R.; Turkov, M.; Stankiewicz, M.; Stuhmer, W.; Dong, K. & Gordon, D. 2007. The insecticidal potential of scorpion β- toxins. Toxicon, 49, 473–489.

Halkier, B. A. & Gershenzon, J. 2006. Biology and biochemistry of glucosinolates. Annu. Rev. Plant Biol., 57, 303-333.

Hamon, A.; Gilles, N.; Sautiere, P.; Martinage, A.; Kopeyan, C.; Ulens, C.; Tytgat, J.; Lancelin, J. M. & Gordon, D. 2002. Characterization of scorpion α-like toxin group using two new toxins from the scorpion Leiurus quinquestriatus hebraeus. Eur. J. Biochem., 269, 3920–3933.

Harper, M. S.; Hopkins, T. L. & Czapla, T. H. 1998. Effect of wheat germ agglutinin on formation and structure of the peritrophic membrane in European corn borer (Ostrinia nubilalis) larvae. Tissue Cell, 30, 166–176.

Hawker, J. S. & Hatch, M. D. 1965. Mechanism of sugar storage by mature stem tissue of sugarcane. Physiol. Plantarum, 18, 444–453.

Hemingway, J. & Ranson, H. 2000. Insecticide resistance in insect vectors of human disease. Annu. Rev. Entomol., 45, 371–391.

Hernandez-Campuzano, B.; Suarez, R.; Lina, L.; Hernandez, V.; Villegas, E.; Corzo, G. & Iturriaga, G. 2009. Expression of a spider venom peptide in transgenic tobacco confers insect resistance. Toxicon, 53, 122–128.

Hester, G. & Wright, C. S. 1996. The mannose-specific bulb lectin from Galanthus nivalis (Snowdrop) binds mono-and dimannosides at distinct sites. Structure analysis of refined complexes at 2.3 Å and 3.0 Å resolution. J. Mol. Biol., 262, 516–531.

Higgins, D. R. 2001. Overview of protein expression in Pichia pastoris. Current protocols in protein science. John Wiley & Sons, Inc.

106 Hilder, V. A. & Boulter, D. 1999. Genetic engineering of crop plants for insect resistance - a critical review. Crop Prot., 18, 177–191.

Hinchee, M. A. W.; Connorward, D. V.; Newell, C. A.; McDonnell, R. E.; Sato, S. J.; Gasser, C. S.; Fischhoff, D. A.; Re, D. B.; Fraley, R. T. & Horsch, R. B. 1988. Production of transgenic soybean plants using Agrobacterium-mediated DNA transfer. Bio-Technol, 6, 915–921.

Hoekema, A.; Hirsch, P. R.; Hooykaas, P. J. J. & Schilperoort, R. A. 1983. A binary plant vector strategy based on separation of vir-region and T-region of the Agrobacterium tumefaciens Ti-plasmid. Nature, 303, 179–180.

Hoffman, D. R. 1993. Allergens in Hymenoptera venom. XXV: The amino acid sequences of antigen 5 molecules and the structural basis of antigenic cross-reactivity. J. Allergy Clin. Immunol., 92, 707–716.

Hong, J.; Yu-xian, Z. & Zhang-liang, C. 1996. Insect resistance of transformed tobacco plants with gene of the spider insecticidal peptide. Acta. Bot. Sin., 38, 95–99.

Hooykaas, P. J. 1989. Transformation of plant cells via Agrobacterium. Plant Mol. Biol., 13, 327–336.

Hopkins, R. J.; van Dam, N. M. & van Loon, J. J. A. 2009. Role of Glucosinolates in insect-plant relationships and multitrophic interactions. Annu. Rev. Entomol., 54, 57–83.

Hopkins, T. L. & Harper, M. S. 2001. Lepidopteran peritrophic membranes and effects of dietary wheat germ agglutinin on their formation and structure. Arch. Insect Biochem. Physiol., 47, 100–109.

Horsch, R. B.; Fry, J. E.; Hoffmann, N. L.; Eichholtz, D.; Rogers, S. G. & Fraley, R. T. 1985. A simple and general-method for transferring genes into plants. Science, 227, 1229–1231.

Huang, J. Q.; Wei, Z. M.; An, H. L. & Zhu, Y. X. 2001. Agrobacterium tumefaciens- mediated transformation of rice with the spider insecticidal gene conferring resistance to leaffolder and striped stem borer. Cell Res., 11, 149–155.

Ignacimuthu, S. & Raveendar, S. 2011. Agrobacterium mediated transformation of indica rice (Oryza sativa L.) for insect resistance. Euphytica, 179, 277–286.

Imlau, A.; Truernit, E. & Sauer, N. 1999. Cell-to-cell and long-distance trafficking of the green fluorescent protein in the phloem and symplastic unloading of the protein into sink tissues. Plant Cell, 11, 309–322.

Invitrogen 2010. EasySelectTM Pichia expression kit - Protein expression: a manual of methods for expression of recombinant proteins using pPICZ and pPICZα in Pichia pastoris (Catalog No. K1740-01).

ISAAA. 2014. ISAAA's GM approval database. [Online]. The International Service for the Acquisition of Agri-biotech Applications (ISAAA). Available: http://www.isaaa.org/gmapprovaldatabase/.

107 James, C. 2012. Global status of commercialized biotech/GM crops: 2012. ISAAA Brief 44. Ithaca, NY: The International Service for the Acquisition of Agri-biotech Applications (ISAAA).

Ji, Y. H.; Mansuelle, P.; Terakawa, S.; Kopeyan, C.; Yanaihara, N.; Hsu, K. & Rochat, H. 1996. Two neurotoxins (BmK I and BmK II) from the venom of the scorpion Buthus martensi Karsch: purification, amino acid sequences and assessment of specific activity. Toxicon, 34, 987–1001.

Johnson, J. H.; Bloomquist, J. R.; Krapcho, K. J.; Kral, R. M., Jr.; Trovato, R.; Eppler, K. G.; Morgan, T. K. & DelMar, E. G. 1998. Novel insecticidal peptides from Tegenaria agrestis spider venom may have a direct effect on the insect central nervous system. Arch. Insect Biochem. Physiol., 38, 19–31.

Johnson, J. H.; Kral, R. M. J. & Krapcho, K. 1997. Insecticidal peptides from spider venom. United States Patent, U. S. Patent No. 5688764 A.

Jones, H. D.; Doherty, A. & Wu, H. 2005. Review of methodologies and a protocol for the Agrobacterium-mediated transformation of wheat. Plant Methods, 1, 5.

Juergensen, K.; Scholz-Starke, J.; Sauer, N.; Hess, P.; van Bel, A. J. E. & Grundler, F. M. W. 2003. The companion cell-specific Arabidopsis disaccharide carrier AtSUC2 is expressed in nematode-induced syncytia. Plant Physiol., 131, 61–69.

Karp, A. 1995. Somaclonal variation as a tool for crop improvement. Euphytica, 85, 295– 302.

Khan, S. A.; Zafar, Y.; Briddon, R. W.; Malik, K. A. & Mukhtar, Z. 2006. Spider venom toxin protects plants from insect attack. Transgenic Res., 15, 349–357.

King, G. 2007a. Natural Insecticides from Spiders. Australasian Science. Hawksburn, Australia, Hawksburn: Control Publications Pty Ltd.

King, G. F. 2007b. Modulation of insect CaV channels by peptidic spider toxins. Toxicon, 49, 513–530.

King, G. F.; Escoubas, P. & Nicholson, G. M. 2008a. Peptide toxins that selectively target insect NaV and CaV channels. Channels, 2, 100–116.

King, G. F.; Gentz, M. C.; Escoubas, P. & Nicholson, G. M. 2008b. A rational nomenclature for naming peptide toxins from spiders and other venomous animals. Toxicon, 52, 264–276.

King, G. F. & Hardy, M. C. 2013. Spider-venom peptides: structure, pharmacology, and potential for control of insect pests. Annu. Rev. Entomol., 58, 475–496.

Klaas Nico, F.; Anita, M. K.; Michael, E. & Marten, V. 2001. A novel method to determine the topology of peroxisomal membrane proteins in vivo using the Tobacco Etch Virus protease. J. Biol. Chem., 276, 36501–36507.

Klein, T. M.; Wolf, E. D.; Wu, R. & Sanford, J. C. 1987. High-velocity microprojectiles for delivering nucleic acids into living cells. Nature, 327, 70–73.

108 Kolarich, D.; Leonard, R.; Hemmer, W. & Altmann, F. 2005. The N-glycans of yellow jacket venom hyaluronidases and the protein sequence of its major isoform in Vespula vulgaris. FEBS J., 272, 5182–5190.

Kopeyan, C.; Mansuelle, P.; Martin-Eauclaire, M. F.; Rochat, H. & Miranda, F. 1993. Characterization of toxin III of the scorpion Leiurus quinquestriatus quinquestriatus: a new type of α-toxin highly toxic both to mammals and insects. Nat. Toxins, 1, 308–312.

Kopeyan, C.; Mansuelle, P.; Sampieri, F.; Brando, T.; Bahraoui, E. M.; Rochat, H. & Granier, C. 1990. Primary structure of scorpion anti-insect toxins isolated from the venom of Leiurus quinquestriatus quinquestriatus. FEBS Lett., 261, 423–426.

Kos, M.; Houshyani, B.; Wietsma, R.; Kabouw, P.; Vet, L. E. M.; van Loon, J. J. A. & Dicke, M. 2012. Effects of glucosinolates on a generalist and specialist leaf- chewing herbivore and an associated parasitoid. Phytochemistry, 77, 162–170.

Koureas, M.; Tsakalof, A.; Tsatsakis, A. & Hadjichristodoulou, C. 2012. Systematic review of biomonitoring studies to determine the association between exposure to organophosphorus and pyrethroid insecticides and human health outcomes. Toxicol. Lett., 210, 155–168.

Koutz, P.; Davis, G. R.; Stillman, C.; Barringer, K.; Cregg, J. & Thill, G. 1989. Structural comparison of the Pichia pastoris alcohol oxidase genes. Yeast, 5, 167–177.

Krapcho, K. J.; Kral, R. M., Jr.; Vanwagenen, B. C.; Eppler, K. G. & Morgan, T. K. 1995. Characterization and cloning of insecticidal peptides from the primitive weaving spider Diguetia canities. Insect Biochem. Mol. Biol., 25, 991–1000.

Krimm, I.; Gilles, N.; Sautiere, P.; Stankiewicz, M.; Pelhate, M.; Gordon, D. & Lancelin, J. M. 1999. NMR structures and activity of a novel alpha-like toxin from the scorpion Leiurus quinquestriatus hebraeus. J. Mol. Biol., 285, 1749–1763.

Kuhn-Nentwig, L.; Stocklin, R. & Nentwig, W. 2011. Venom composition and strategies in spiders: is everything possible? In: Casas, J. (ed.) Advances in Insect Physiology. London: Academic Press

Labra, M.; Vannini, C.; Grassi, F.; Bracale, M.; Balsemin, M.; Basso, B. & Sala, F. 2004. Genomic stability in Arabidopsis thaliana transgenic plants obtained by floral dip. Theore. App. Genetic., 109, 1512–1518.

Larkin, P. J. & Scowcroft, W. R. 1981. Somaclonal variation — a novel source of variability from cell cultures for plant improvement. Theore. App. Genet., 60, 197–214.

Li, D.; Xiao, Y.; Hu, W.; Xie, J.; Bosmans, F.; Tytgat, J. & Liang, S. 2003. Function and solution structure of hainantoxin-I, a novel insect sodium channel inhibitor from the Chinese bird spider Selenocosmia hainana. FEBS Lett., 555, 616–622.

Li, P.; Anumanthan, A.; Gao, X. G.; Ilangovan, K.; Suzara, V.; Düzgüneş, N. & Renugopalakrishnan, V. 2007. Expression of recombinant proteins in Pichia pastoris. Appl. Biochem. Biotech., 142, 105–124.

Li, Y. J. & Ji, Y. H. 2000. Binding characteristics of BmK I, an alpha-like scorpion neurotoxic polypeptide, on cockroach nerve cord synaptosomes. J. Pept. Res., 56, 195–200.

109 Lima, M. E. D. & Martin-Eauclaire, M.-F. 1995. The toxins purified from Tityus Serrulatus (Lutz & Mello) venom. Toxin Rev., 14, 457–481.

LIN Tong, W. Z.-Y., LIU Kuan-Yu, JING Tian-Zhong, ZHANG Chuan-Xi 2006. Transformation of spider neurotoxin gene with prospective insecticidal properties into hybrid poplar Populus simonii x P. nigra. Acta Entomol. Sinica, 49, 593–598.

Liu, F.; Cao, M. Q.; Yao, L.; Li, Y.; Robaglia, C. & Tourneur, C. 1998. In planta transformation of Pakchoi (Brassica campestris L. ssp. Chinensis) by infiltration of adult plants with Agrobacterium. In: Rubatzky, V. E., Hang, C. & Peron, J. Y. (eds.) Third International Symposium on Diversification of Vegetable Crops.

Liu, Z. C.; Zhang, R.; Zhao, F.; Chen, Z. M.; Liu, H. W.; Wang, Y. J.; Jiang, P.; Zhang, Y.; Wu, Y.; Ding, J. P.; Lee, W. H. & Zhang, Y. 2012. Venomic and transcriptomic analysis of centipede Scolopendra subspinipes dehaani. J. Proteome Res., 11, 6197–6212.

Loret, E. P.; Mansuelle, P.; Rochat, H. & Granier, C. 1990. Neurotoxins active on insects: amino acid sequences, chemical modifications, and secondary structure estimation by circular dichroism of toxins from the scorpion Androctonus australis Hector. Biochem. , 29, 1492-1501.

Lu, C. & Kang, J. 2008. Generation of transgenic plants of a potential oilseed crop Camelina sativa by Agrobacterium-mediated transformation. Plant Cell Rep., 27, 273–278.

Maggio, F. & King, G. F. 2002a. Role of the structurally disordered N- and C-terminal residues in the Janus-faced atracotoxins. Toxicon, 40, 1355—1361.

Maggio, F. & King, G. F. 2002b. Scanning mutagenesis of a Janus-faced atracotoxin reveals a bipartite surface patch that is essential for neurotoxic function. J. Biol. Chem., 277, 22806 —22813.

Maggio, F.; Sollod, B. L.; Tedford, H. W. & King, G. F. 2005. Spider toxins and their potential for insect control. In: Gilbert, L. I. (ed.) Comprehensive Molecular Insect Science. Amsterdam: Elsevier.

Maiti, I. B.; Ghosh, S. K.; Gowda, S.; Kiernan, J. & Shepherd, R. J. 1997. Promoter/leader deletion analysis and plant expression vectors with the figwort mosaic virus (FMV) full length transcript (FLt) promoter containing single or double enhancer domains. Transgenic Res., 6, 143–156.

Malta, M. B.; Lira, M. S.; Soares, S. L.; Rocha, G. C.; Knysak, I.; Martins, R.; Guizze, S. P.; Santoro, M. L. & Barbaro, K. C. 2008. Toxic activities of Brazilian centipede venoms. Toxicon, 52, 255–263.

Maqbool, S.; Riazuddin, S.; Loc, N.; Gatehouse, A. R.; Gatehouse, J. & Christou, P. 2001. Expression of multiple insecticidal genes confers broad resistance against a range of different rice pests. Mol. Breeding, 7, 85–93.

Martinez-Garcia, J. F.; Monte, E. & Quail, P. H. 1999. A simple, rapid and quantitative method for preparing Arabidopsis protein extracts for immunoblot analysis. Plant J., 20, 251–257.

110 McCaffery, A. R. 1998. Resistance to insecticides in heliothine Lepidoptera: a global view. Philos. T. Roy. Soc. B, 353, 1735–1750.

Meinwald, J. & Eisner, T. 1995. The chemistry of phyletic dominance. Proc. Natl. Acad. Sci. USA, 92, 14–18.

Mendelsohn, M.; Kough, J.; Vaituzis, Z. & Matthews, K. 2003. Are Bt crops safe? Nat. Biotechnol., 21, 1003–1009.

Michiels, K.; Van Damme, E. J. M. & Smagghe, G. 2010. Plant-insect interactions: what can we learn from plant lectins? Arch. Insect Biochem. Physiol., 73, 193–212.

Miller, W. A. & Bonning, B. C. 2003. Plant resistance to insect pests mediated by viral proteins. United States Patent, US Patent No. 7312080.

Mithen, R.; Bennett, R. & Marquez, J. 2010. Glucosinolate biochemical diversity and innovation in the Brassicales. Phytochemistry, 71, 2074–2086.

Mithofer, A. & Boland, W. 2012. Plant defense against herbivores: chemical aspects. Annu. Rev. Plant Biol., 63, 431–450.

Monsanto. 2014. Genuity® VT Triple PRO® RIB Complete®: single-bag option for managing above- and below-ground pests [Online]. Available: http://www.monsanto.com/products/pages/genuity-vt-triple-pro-rib-complete- corn.aspx.

Mukherjee, A. K.; Sollod, B. L.; Wikel, S. K. & King, G. F. 2006. Orally active acaricidal peptide toxins from spider venom. Toxicon, 47, 182–187.

Muller, R.; de Vos, M.; Sun, J. Y.; Sonderby, I. E.; Halkier, B. A.; Wittstock, U. & Jander, G. 2010. Differential effects of indole and aliphatic glucosinolates on lepidopteran herbivores. J. Chem. Ecol., 36, 905–913.

Murdock, L. L.; Huesing, J. E.; Nielsen, S. S.; Pratt, R. C. & Shade, R. E. 1990. Biological effects of plant lectins on the cowpea weevil. Phytochemistry, 29, 85–89.

Mylne, J. & Botella, J. R. 1998. Binary vectors for sense and antisense expression of Arabidopsis ESTs. Plant Mol. Biol. Report, 16, 257–262.

Nakajima, T.; Uzu, S.; Wakamatsu, K.; Saito, K.; Miyazawa, T.; Yasuhara, T.; Tsukamoto, Y. & Fujino, M. 1986. Amphiphilic peptides in wasp venom. Biopolymers, 25 Suppl., S115–121.

Nakasu, E. Y.; Williamson, S. M.; Edwards, M. G.; Fitches, E. C.; Gatehouse, J. A.; Wright, G. A. & Gatehouse, A. M. 2014. Novel biopesticide based on a spider venom peptide shows no adverse effects on honeybees. Proc. R. Soc. B, 281, 20140619.

Narusaka, M.; Shiraishi, T.; Iwabuchi, M. & Narusaka, Y. 2010. The floral inoculating protocol: a simplified Arabidopsis thaliana transformation method modified from floral dipping. Plant Biotechnol., 27, 349–351.

Nicholson, G. M. 2007. Fighting the global pest problem: preface to the special Toxicon issue on insecticidal toxins and their potential for insect pest control. Toxicon, 49, 413–422.

111 Nolte, K. D. & Koch, K. E. 1993. Companion-cell specific localization of sucrose synthase in zones of phloem loading and unloading. Plant Physiol., 101, 899–905.

Novotny, V.; Basset, Y.; Miller, S. E.; Weiblen, G. D.; Bremer, B.; Cizek, L. & Drozd, P. 2002. Low host specificity of herbivorous insects in a tropical forest. Nature, 416, 841–844.

Odell, J. T.; Nagy, F. & Chua, N. H. 1985. Identification of DNA sequences required for activity of the Cauliflower Mosaic Virus 35S promoter. Nature, 313, 810–812.

Oerke, E. C. 2006. Crop losses to pests. J. Agric. Sci., 144, 31–43.

Oerke, E. C. & Dehne, H. W. 2004. Safeguarding production - losses in major crops and the role of crop protection. Crop Prot., 23, 275–285.

Oerke, E. C.; Dehne, H. W.; Schönbeck, F. & Weber, A. 1994. Crop Production and Crop Protection–Estimated Losses In Major Food and Cash Crops, Amsterdam, Elsevier Science Ltd.

Oliveira, L. C.; De Lima, M. E.; Pimenta, A. M.; Mansuelle, P.; Rochat, H.; Cordeiro, M. N.; Richardson, M. & Figueiredo, S. G. 2003. PnTx4-3, a new insect toxin from Phoneutria nigriventer venom elicits the glutamate uptake inhibition exhibited by PhTx4 toxic fraction. Toxicon, 42, 793–800.

Oltmanns, H.; Frame, B.; Lee, L.-Y.; Johnson, S.; Li, B.; Wang, K. & Gelvin, S. B. 2010. Generation of backbone-free, low transgene copy plants by launching T-DNA from the Agrobacterium chromosome. Plant Physiol., 152, 1158–1166.

Omar, A. & Chatha, K. A. 2012. National Institute for Biotechnology and Genetic Engineering (NIBGE): genetically modified spider cotton. Asian J. Manag. Case, 9, 33–58.

Oren, D. A.; Froy, O.; Amit, E.; Kleinberger-Doron, N.; Gurevitz, M. & Shaanan, B. 1998. An excitatory scorpion toxin with a distinctive feature: an additional alpha helix at the C terminus and its implications for interaction with insect sodium channels. Structure, 6, 1095–1103.

Orivel, J.; Redeker, V.; Le Caer, J. P.; Krier, F.; Revol-Junelles, A. M.; Longeon, A.; Chaffotte, A.; Dejean, A. & Rossier, J. 2001. Ponericins, new antibacterial and insecticidal peptides from the venom of the ant Pachycondyla goeldii. J. Biol. Chem., 276, 17823–17829.

Palma, M. S. 2006. Insect venom peptides. In: Kastin, A. J. (ed.) Handbook of Biologically Active Peptides. Burlington: Academic Press.

Palomares, L. A.; Estrada-Mondaca, S. & Ramírez, O. T. 2004. Production of recombinant proteins: challenges and solutions. Methods Mol. Biol., 267, 15–51.

Pelhate, M.; Stankiewicz, M. & Ben Khalifa, R. 1998. Anti-insect scorpion toxins: historical account, activities and prospects. C. R. Seances. Soc. Biol. Fil., 192, 463–484.

Pelhate, M. & Zlotkin, E. 1982. Actions of insect toxin and other toxins derived from the venom of the scorpion Androctonus australis on isolated giant axons of the cockroach (Periplaneta americana). J. Exp. Biol., 97, 67–77.

112 Pence, R. J. 1965. The antimetabolite: imidazole as a pesticide. California Agric., 19, 13– 15.

Pham, N. 2010. Tissue specific expression of insecticidal toxins in plants. M Sc Thesis, The University of Queensland.

Piek, T.; Duval, A.; Hue, B.; Karst, H.; Lapied, B.; Mantel, P.; Nakajima, T.; Pelhate, M. & Schmidt, J. O. 1991. Poneratoxin, a novel peptide neurotoxin from the venom of the ant, Paraponera clavata. Comp. Biochem. Physiol. C., 99, 487–495.

Pimenta, A. M.; Martin-Eauclaire, M.; Rochat, H.; Figueiredo, S. G.; Kalapothakis, E.; Afonso, L. C. & De Lima, M. E. 2001. Purification, amino-acid sequence and partial characterization of two toxins with anti-insect activity from the venom of the South American scorpion Tityus bahiensis (Buthidae). Toxicon, 39, 1009–1019.

Pimentel, D. & Levitan, L. 1986. Pesticides: amounts applied and amounts reaching pests. BioScience, 36, 86–91.

Pirpignani, M. L.; Rivera, E.; Hellman, U. & Biscoglio de Jimenez Bonino, M. 2002. Structural and immunological aspects of Polybia scutellaris antigen 5. Arch. Biochem. Biophys., 407, 224–230.

Pitzschke, A. & Hirt, H. 2010. New insights into an old story: Agrobacterium-induced tumour formation in plants by plant transformation. EMBO J., 29, 1021-1032.

Potenza, C.; Aleman, L. & Sengupta-Gopalan, C. 2004. Targeting transgene expression in research, agricultural, and environmental applications: promoters used in plant transformation. In Vitro Cell. Dev. Biol.–Plant, 40, 1–22.

Poulsen, M.; Kroghsbo, S.; Schroder, M.; Wilcks, A.; Jacobsen, H.; Miller, A.; Frenzel, T.; Danier, J.; Rychlik, M.; Shu, Q.; Emami, K.; Sudhakar, D.; Gatehouse, A.; Engel, K. H. & Knudsen, I. 2007. A 90-day safety study in Wistar rats fed genetically modified rice expressing snowdrop lectin Galanthus nivalis (GNA). Food Chem. Toxicol., 45, 350–363.

Powell, K. S.; Spence, J.; Bharathi, M.; Gatehouse, J. A. & Gatehouse, A. M. R. 1998. Immunohistochemical and developmental studies to elucidate the mechanism of action of the snowdrop lectin on the rice brown planthopper, Nilaparvata lugens (Stal). J. Insect Physiol., 44, 529–539.

Qaim, M. & Kouser, S. 2013. Genetically modified crops and food security. PLoS One, 8, e64879.

Qing, C.; Fan, L.; Lei, Y.; Bouchez, D.; Tourneur, C.; Yan, L. & Robaglia, C. 2000. Transformation of Pakchoi (Brassica rapa L. ssp. chinensis) by Agrobacterium infiltration. Mol. Breeding, 6, 67–72.

Quistad, G. B.; Reuter, C. C.; Skinner, W. S.; Dennis, P. A.; Suwanrumpha, S. & Fu, E. W. 1991. Paralytic and insecticidal toxins from the funnel web spider, Hololena curta. Toxicon, 29, 329–336.

R Development Core Team. 2013. R: a language and environment for statistical computing. R foundation for statistical computing.

113 Ramesh, S.; Nagadhara, D.; Pasalu, I. C.; Kumari, A. P.; Sarma, N. P.; Reddy, V. D. & Rao, K. V. 2004. Development of stem borer resistant transgenic parental lines involved in the production of hybrid rice. J. Biotechnol., 111, 131-41.

Rao, K. V.; Rathore, K. S.; Hodges, T. K.; Fu, X.; Stoger, E.; Sudhakar, D.; Williams, S.; Christou, P.; Bharathi, M.; Bown, D. P.; Powell, K. S.; Spence, J.; Gatehouse, A. M. & Gatehouse, J. A. 1998. Expression of snowdrop lectin (GNA) in transgenic rice plants confers resistance to rice brown planthopper. Plant J., 15, 469–77.

Rasband, W. S. 1997—2014. ImageJ. 1.48 ed. Bethesda, Maryland, USA: U. S. National Institutes of Health.

Rash, L. D. & Hodgson, W. C. 2002. Pharmacology and biochemistry of spider venoms. Toxicon, 40, 225–254.

Rates, B.; Bemquerer, M. P.; Richardson, M.; Borges, M. H.; Morales, R. A. V.; De Lima, M. E. & Pimenta, A. M. C. 2007. Venomic analyses of Scolopendra viridicornis nigra and Scolopendra angulata (Centipede, Scolopendromorpha): Shedding light on venoms from a neglected group. Toxicon, 49, 810–826.

Rauch, S. A.; Braun, J. M.; Barr, D. B.; Calafat, A. M.; Khoury, J.; Montesano, A. M.; Yolton, K. & Lanphear, B. P. 2012. Associations of prenatal exposure to organophosphate pesticide metabolites with gestational age and birth weight. Environ. Health Perspect, 120, 1055–1060.

Redei, G. P. 1992. A heuristic glance at the past of Arabidopsis genetics. In: Koncz, N. H. C. C. & Schell, J. (eds.) Methods in Arabidopsis research. Singapore: World Scientific.

Richardson, J. R.; Roy, A.; Shalat, S. L.; von Stein, R. T.; Hossain, M. M.; Buckley, B.; Gearing, M.; Levey, A. I. & German, D. C. 2014. Elevated serum pesticide levels and risk for alzheimer disease. JAMA Neurol., 71, 284–290.

Rivera, A. L.; Gomez-Lim, M.; Fernandez, F. & Loske, A. M. 2012. Physical methods for genetic plant transformation. Phys. Life Rev., 9, 308–345.

Robinson, C. R. & Sauer, R. T. 1998. Optimizing the stability of single-chain proteins by linker length and composition mutagenesis. Proc. Natl. Acad. Sci. USA, 95, 5929– 5934.

Rodriguez de la Vega, R. C. & Possani, L. D. 2005. Overview of scorpion toxins specific for Na+ channels and related peptides: biodiversity, structure-function relationships and evolution. Toxicon, 46, 831–44.

Rohr, F.; Ulrichs, C.; Schreiner, M.; Zrenner, R. & Mewis, I. 2012. Responses of Arabidopsis thaliana plant lines differing in hydroxylation of aliphatic glucosinolate side chains to feeding of a generalist and specialist caterpillar. Plant Physiol. Bioch., 55, 52–59.

Romeis, J.; Babendreier, D. & Wäckers, F. L. 2003. Consumption of snowdrop lectin (Galanthus nivalis agglutinin) causes direct effects on adult parasitic wasps. Oecologia, 134, 528–536.

Sadeghi, A.; Smagghe, G.; Broeders, S.; Hernalsteens, J.-P.; Greve, H.; Peumans, W. & Damme, E. M. 2008. Ectopically expressed leaf and bulb lectins from garlic (Allium

114 sativum L.) protect transgenic tobacco plants against cotton leafworm (Spodoptera littoralis). Transgenic Res., 17, 9–18.

Saez, N. J.; Senff, S.; Jensen, J. E.; Er, S. Y.; Herzig, V.; Rash, L. D. & King, G. F. 2010. Spider-venom peptides as therapeutics. Toxins, 2, 2851–2871.

Saha, P.; Chakraborti, D.; Sarkar, A.; Dutta, I.; Basu, D. & Das, S. 2007. Characterization of vascular-specific RSs1 and rolC promoters for their utilization in engineering plants to develop resistance against hemipteran insect pests. Planta, 226, 429– 442.

Sambrook, J.; Fritsch, E. F. & Maniatis, T. 1989. Molecular cloning: a laboratory manual Cold Spring Harbor, New York, Cold Spring Harbor Laboratory Press.

Sattelle, D. B.; Cordova, D. & Cheek, T. R. 2008. Insect ryanodine receptors: molecular targets for novel pest control chemicals. Invert. Neurosci., 8, 107–119.

Sauer, N. & Stolz, J. 1994. SUC1 and SUC2: 2 sucrose transporters from Arabidopsis thaliana; expression and characterization in bakers-yeast and identification of the histidine-tagged protein Plant J., 6, 67–77.

Schell, J. & Van Montagu, M. 1977. The Ti-plasmid of Agrobacterium tumefaciens, a natural vector for the introduction of NIF genes in plants? Basic Life Sci., 9, 159– 179.

Schmidt, S. R. 2013. Fusion proteins: applications and challenges. In: Schmidt, S. R. (ed.) Fusion protein technologies for biopharmaceuticals: applications and challenges. Hoboken, New Jersey: John Wiley & sons, Inc.

Schmulling, T.; Schell, J. & Spena, A. 1989. Promoters of the rolA, B, and C genes of Agrobacterium rhizogenesare differentially regulated in transgenic plants. Plant Cell, 1, 665–670.

Schwartz, E. F.; Mourao, C. B.; Moreira, K. G.; Camargos, T. S. & Mortari, M. R. 2012. Arthropod venoms: a vast arsenal of insecticidal neuropeptides. Biopolymers, 98, 385–405.

Sears, I. B.; O'Connor, J.; Rossanese, O. W. & Glick, B. S. 1998. A versatile set of vectors for constitutive and regulated gene expression in Pichia pastoris. Yeast, 14, 783– 790.

Shah, A.; Ahmed, M.; Mukhtar, Z.; Khan, S.; Habib, I.; Malik, Z.; Mansoor, S. & Saeed, N. 2011. Spider toxin (Hvt) gene cloned under phloem specific RSs1 and RolC promoters provides resistance against American bollworm (Heliothis armigera). Biotechnol. Lett., 33, 1457–1463.

Sharma, H. C.; Sharma, K. K. & Crouch, J. H. 2004. Genetic transformation of crops for insect resistance: potential and limitations. Crit. Rev. Plant Sci., 23, 47–72.

Sharma, H. C.; Sharma, K. K.; Seetharama, N. & Ortiz, R. 2000. Prospects for using transgenic resistance to insects in crop improvement. Electron. J. Biotech., 3, 21– 22.

115 Shen, S.; Sulter, G.; Jeffries, T. W. & Cregg, J. M. 1998. A strong nitrogen source- regulated promoter for controlled expression of foreign genes in the yeast Pichia pastoris. Gene, 216, 93–102.

Shi, Y.; Wang, M. B.; Powell, K. S.; Van Damme, E.; Hilder, V. A.; Gatehouse, A. M. R.; Boulter, D. & Gatehouse, J. A. 1994. Use of the rice sucrose synthase-1 promoter to direct phloem-specific expression of β-glucuronidase and snowdrop lectin genes in transgenic tobacco plants. J. Exp. Botany, 45, 623–631.

Shroff, R.; Vergara, F.; Muck, A.; Svatoš, A. & Gershenzon, J. 2008. Nonuniform distribution of glucosinolates in Arabidopsis thaliana leaves has important consequences for plant defense. Proc. Natl. Acad. Sci. USA, 105, 6196–6201.

Skinner, W. S.; Adams, M. E.; Quistad, G. B.; Kataoka, H.; Cesarin, B. J.; Enderlin, F. E. & Schooley, D. A. 1989. Purification and characterization of two classes of neurotoxins from the funnel web spider, Agelenopsis aperta. J. Biol. Chem., 264, 2150–2155.

Skinner, W. S.; Dennis, P. A.; Li, J. P. & Quistad, G. B. 1992. Identification of insecticidal peptides from venom of the trap-door spider, Aptostichus schlingeri (Ctenizidae). Toxicon, 30, 1043–1050.

Smith, J.; Herzig, V.; King, G. & Alewood, P. 2013. The insecticidal potential of venom peptides. Cell. Mol. Life Sci., 70, 3665–3693.

Soldatova, L.; Kochoumian, L. & King, T. P. 1993. Sequence similarity of a hornet (D. maculata) venom allergen phospholipase A1 with mammalian lipases. FEBS Lett., 320, 145–149.

Somerville, C. & Koornneef, M. 2002. A fortunate choice: the history of Arabidopsis as a model plant. Nat. Rev. Genet., 3, 883–889.

Stadler, R. & Sauer, N. 1996. The Arabidopsis thaliana AtSUC2 gene is specifically expressed in companion cells. Botanica Acta, 109, 299–306.

Stapleton, A.; Blankenship, D. T.; Ackermann, B. L.; Chen, T. M.; Gorder, G. W.; Manley, G. D.; Palfreyman, M. G.; Coutant, J. E. & Cardin, A. D. 1990. Curtatoxins. Neurotoxic insecticidal polypeptides isolated from the funnel-web spider Hololena curta. J. Biol. Chem., 265, 2054–2059.

Stauber, E. J.; Kuczka, P.; van Ohlen, M.; Vogt, B.; Janowitz, T.; Piotrowski, M.; Beuerle, T. & Wittstock, U. 2012. Turning the 'Mustard Oil Bomb' into a 'Cyanide Bomb': aromatic glucosinolate metabolism in a specialist insect herbivore. PLoS One, 7, e35545.

Stevens, J.; Dunse, K.; Fox, J.; Evans, S. & Anderson, M. 2012. Biotechnological approaches for the control of insect pests in crop plants. In: Soundararajan, R. P. (ed.) Pesticides–Advances in Chemical and Botanical Pesticides. InTech.

Storer, N. P.; Babcock, J. M.; Schlenz, M.; Meade, T.; Thompson, G. D.; Bing, J. W. & Huckaba, R. M. 2010. Discovery and characterization of field resistance to Bt maize: Spodoptera frugiperda (Lepidoptera: Noctuidae) in Puerto Rico. J. Econ. Entomol., 103, 1031–1038.

116 Storer, N. P.; Kubiszak, M. E.; Ed King, J.; Thompson, G. D. & Santos, A. C. 2012. Status of resistance to Bt maize in Spodoptera frugiperda: lessons from Puerto Rico. J. Invertebr. Pathol., 110, 294–300.

Sun, J. Y.; Sonderby, I. E.; Halkier, B. A.; Jander, G. & de Vos, M. 2009. Non-volatile intact indole glucosinolates are host recognition cues for ovipositing Plutella xylostella. J. Chem. Ecol., 35, 1427–1436.

Sunilkumar, G.; Mohr, L.; Lopata-Finch, E.; Emani, C. & Rathore, K. S. 2002. Developmental and tissue-specific expression of CaMV 35S promoter in cotton as revealed by GFP. Plant Mol. Biol., 50, 463–474.

Tabashnik, B. E.; Brevault, T. & Carriere, Y. 2013. Insect resistance to Bt crops: lessons from the first billion acres. Nat. Biotechnol., 31, 510–521.

Tabashnik, B. E.; Van Rensburg, J. B. & Carriere, Y. 2009. Field-evolved insect resistance to Bt crops: definition, theory, and data. J. Econ. Entomol., 102, 2011–2025.

Tamborindeguy, C.; Monsion, B.; Brault, V.; Hunnicutt, L.; Ju, H. J.; Nakabachi, A. & Van Fleet, E. 2010. A genomic analysis of transcytosis in the pea aphid, Acyrthosiphon pisum, a mechanism involved in virus transmission. Insect Mol. Biol., 19 Suppl. 2, 259–272.

Tedford, H. W.; Gilles, N.; Menez, A.; Doering, C. J.; Zamponi, G. W. & King, G. F. 2004a. Scanning mutagenesis of ω-atracotoxin-Hv1a reveals a spatially restricted epitope that confers selective activity against insect calcium channels. J. Biol. Chem., 279, 44133–44140.

Tedford, H. W., Jamie I Flecher, King G. F. 2001. Functional significance of the β-hairpin in the insecticidal neurotoxin ω-Atracotoxin-Hv1a. J. Biol. Chem., 276, 26568– 26576.

Tedford, H. W.; Sollod, B. L.; Maggio, F. & King, G. F. 2004b. Australian funnel-web spiders: master insecticide chemists. Toxicon, 43, 601–618.

Thacker, J. R. M. 2002. An introduction to arthropod pest control, Cambridge, United Kingdom, Cambridge University Press.

The Arabidopsis Initiative 2000. Analysis of the genome sequence of the flowering plant Arabidopsis thaliana. Nature, 408, 796–815.

Tian, Y. C.; Qin, X. F.; Xu, B. Y.; Li, T. Y.; Fang, R. X.; Mang, K. Q.; Li, W. G.; Fu, W. J.; Li, Y. P.; Zhang, S. F. & et al. 1991. Insect resistance of transgenic tobacco plants expressing δ-endotoxin gene of Bacillus thuringiensis. Chin. J. Biotechnol., 7, 1– 13.

Toenniessen, G. H.; O'Toole, J. C. & DeVries, J. 2003. Advances in plant biotechnology and its adoption in developing countries. Curr. Opin. Plant Biol., 6, 191–198.

Trieu, A. T.; Burleigh, S. H.; Kardailsky, I. V.; Maldonado-Mendoza, I. E.; Versaw, W. K.; Blaylock, L. A.; Shin, H.; Chiou, T. J.; Katagi, H.; Dewbre, G. R.; Weigel, D. & Harrison, M. J. 2000. Transformation of Medicago truncatula via infiltration of seedlings or flowering plants with Agrobacterium. Plant J., 22, 531–541.

117 Truernit, E. & Sauer, N. 1995. The promoter of the Arabidopsis thaliana SUC2 sucrose-H+ symporter gene directs expression of β-glucuronidase to the phloem: evidence for phloem loading and unloading by SUC2. Planta, 196, 564–570.

Trung, N. P.; Fitches, E. & Gatehouse, J. A. 2006. A fusion protein containing a lepidopteran-specific toxin from the south Indian red scorpion (Mesobuthus tamulus) and snowdrop lectin shows oral toxicity to target insects. BMC Biotechnol., 6, 18.

Tu, J.; Zhang, G.; Datta, K.; Xu, C.; He, Y.; Zhang, Q.; Khush, G. S. & Datta, S. K. 2000. Field performance of transgenic elite commercial hybrid rice expressing Bacillus thuringiensis δ-endotoxin. Nat. Biotechnol., 18, 1101–1104.

Undheim, E. A.; Sunagar, K.; Hamilton, B. R.; Jones, A.; Venter, D. J.; Fry, B. G. & King, G. F. 2014. Multifunctional warheads: diversification of the toxin arsenal of centipedes via novel multidomain transcripts. J. Proteomics, 102, 1–10.

Undheim, E. A. B. & King, G. F. 2011. On the venom system of centipedes (Chilopoda), a neglected group of venomous animals. Toxicon, 57, 512–524.

Van Maele-Fabry, G.; Hoet, P. & Lison, D. 2013. Parental occupational exposure to pesticides as risk factor for brain tumors in children and young adults: a systematic review and meta-analysis. Environ. Int., 56, 19–31. van Rensburg, J. B. J. 2007. First report of field resistance by the stem borer, Busseola fusca (Fuller) to Bt-transgenic maize. South African J. Plant Soil, 24, 147–151.

Vandenborre, G.; Smagghe, G. & Van Damme, E. J. M. 2011. Plant lectins as defense proteins against phytophagous insects. Phytochemistry, 72, 1538–1550.

Vargas, O.; Martin, M. F. & Rochat, H. 1987. Characterization of six toxins from the venom of the Moroccan scorpion Buthus occitanus mardochei. Eur. J. Biochem., 162, 589–599.

Vasconcelos, I. M. & Oliveira, J. T. 2004. Antinutritional properties of plant lectins. Toxicon, 44, 385–403.

Vassilevski, A. A.; Kozlov, S. A. & Grishin, E. V. 2009. Molecular diversity of spider venom. Biochemistry (Moscow), 74, 1505–1534.

Wakefield, M. E.; Bell, H. A.; Fitches, E. C.; Edwards, J. P. & Gatehouse, A. M. R. 2006. Effects of Galanthus nivalis agglutinin (GNA) expressed in tomato leaves on larvae of the tomato moth Lacanobia oleracea (Lepidoptera: Noctuidae) and the effect of GNA on the development of the endoparasitoid Meteorus gyrator (Hymenoptera: Braconidae). B. Entomol. Res., 96, 43–52.

Wakefield, M. E.; Fitches, E. C.; Bell, H. A. & Gatehouse, A. M. R. 2010. The snowdrop lectin Galanthus nivalis agglutinin (GNA) and a fusion protein ButaIT/GNA have a differential affect on a pest noctuid Lacanobia oleracea and the ectoparasitoid Eulophus pennicornis. Physiol. Entomol., 35, 334–342.

Wang, J.; Chen, Z.; Du, J.; Sun, Y. & Liang, A. 2005. Novel insect resistance in Brassica napus developed by transformation of chitinase and scorpion toxin genes. Plant Cell Rep., 24, 549–555.

118 Wang, M. B.; Boulter, D. & Gatehouse, J. A. 1992. A complete sequence of the rice sucrose synthase-1(RSS1) gene. Plant Mol. Biol., 19, 881–885.

Wang, W. C.; Menon, G. & Hansen, G. 2003. Development of a novel Agrobacterium- mediated transformation method to recover transgenic Brassica napus plants. Plant Cell Rep., 22, 274–281.

Wang, X. H.; Connor, M.; Smith, R.; Maciejewski, M. W.; Howden, M. E. H.; Nicholson, G. M.; Christie, M. J. & King, G. F. 2000. Discovery and characterization of a family of insecticidal neurotoxins with a rare vicinal disulfide bridge. Nat. Struc. Biol., 7, 505–513.

Wang, X. H.; Connor, M.; Wilson, D.; Wilson, H. I.; Nicholson, G. M.; Smith, R.; Shaw, D.; Mackay, J. P.; Alewood, P. F.; Christie, M. J. & King, G. F. 2001. Discovery and structure of a potent and highly specific blocker of insect calcium channels. J. Biol. Chem., 276, 40306–40312.

Wang, X. H.; Smith, R.; Fletcher, J. I.; Wilson, H.; Wood, C. J.; Howden, M. E. H. & King, G. F. 1999. Structure-function studies of ω-atracotoxin, a potent antagonist of insect voltage-gated calcium channels. Euro. J. Biochem., 264, 488–494.

Waterham, H. R.; Digan, M. E.; Koutz, P. J.; Lair, S. V. & Cregg, J. M. 1997. Isolation of the Pichia pastoris glyceraldehyde-3-phosphate dehydrogenase gene and regulation and use of its promoter. Gene, 186, 37–44.

Whetstone, P. A. & Hammock, B. D. 2007. Delivery methods for peptide and protein toxins in insect control. Toxicon, 49, 576–596.

Wilkinson, J. E.; Twell, D. & Lindsey, K. 1997. Activities of CaMV 35S and nos promoters in pollen: implications for field release of transgenic plants. J. Exp. Botany, 48, 265–275.

Williams, I. S. 1999. Slow-growth, high-mortality – a general hypothesis, or is it? Ecol. Entomol., 24, 490–495.

Windley, M. J.; Escoubas, P.; Valenzuela, S. M. & Nicholson, G. M. 2011. A novel family of insect-selective peptide neurotoxins targeting insect large-conductance calcium- activated K+ channels isolated from the venom of the theraphosid spider Eucratoscelus constrictus. Mol. Pharmacol., 80, 1–13.

Windley, M. J.; Herzig, V.; Dziemborowicz, S. A.; Hardy, M. C.; King, G. F. & Nicholson, G. M. 2012. Spider-venom peptides as bioinsecticides. Toxins, 4, 191–227.

Wittstock, U. & Burow, M. 2010. Glucosinolate breakdown in Arabidopsis: mechanism, regulation and biological significance. Arabidopsis Book, 8, e0134.

Wu, J.; Luo, X.; Wang, Z.; Tian, Y.; Liang, A. & Sun, Y. 2008. Transgenic cotton expressing synthesized scorpion insect toxin AaHIT gene confers enhanced resistance to cotton bollworm (Heliothis armigera) larvae. Biotechnol. Lett., 30, 547–554.

Wu, N. F.; Sun, Q.; Yao, B.; Fan, Y. L.; Rao, H. Y.; Huang, M. R. & Wang, M. X. 2000. Insect-resistant transgenic poplar expressing AaIT gene. Chinese J. Biotechnol., 16, 129–133.

119 Xiao, K.; Zhang, C.; Harrison, M. & Wang, Z. Y. 2005. Isolation and characterization of a novel plant promoter that directs strong constitutive expression of transgenes in plants. Mol. Breeding, 15, 221–231.

Yang, N. S. & Russell, D. 1990. Maize sucrose synthase-1 promoter directs phloem cell- specific expression of Gus gene in transgenic tobacco plants. Proc. Natl. Acad. Sci. USA, 87, 4144–4148.

Yang, S.; Liu, Z.; Xiao, Y.; Li, Y.; Rong, M.; Liang, S.; Zhang, Z.; Yu, H.; King, G. F. & Lai, R. 2012. Chemical punch packed in venoms makes centipedes excellent predators. Mol. Cell Proteomics, 11, 640–650.

Yang, Y.; Li, Y. & Wu, Y. 2013. Current status of insecticide resistance in Helicoverpa armigera after 15 years of Bt cotton planting in China. J. Econ. Entomol., 106, 375–381.

Yao, B.; Fan, Y.; Zeng, Q. & Zhao, R. 1996. Insect-resistant tobacco plants expressing insect-specific neurotoxin AaIT. Chinese J. Biotechnol., 12, 67–72.

Ye, G. N.; Stone, D.; Pang, S. Z.; Creely, W.; Gonzalez, K. & Hinchee, M. 1999. Arabidopsis ovule is the target for Agrobacterium in planta vacuum infiltration transformation. Plant J., 19, 249–257.

Zalucki, M.; Daglish, G.; Firempong, S. & Twine, P. 1986. The biology and ecology of Heliothis armigera (Hubner) and Heliothis punctigera Wallengren (Lepidoptera, Noctuidae) in Australia - what do we know? Australian J. Zoology, 34, 779–814.

Zhang, P. F.; Chen, P.; Hu, W. J. & Liang, S. P. 2003. Huwentoxin-V, a novel insecticidal peptide toxin from the spider Selenocosmia huwena, and a natural mutant of the toxin: indicates the key amino acid residues related to the biological activity. Toxicon, 42, 15–20.

Zhao, J. H.; Ho, P. & Azadi, H. 2011. Benefits of Bt cotton counterbalanced by secondary pests? Perceptions of ecological change in China. Environ. Monit. Assess., 173, 985–994.

Zhao, J. Z.; Cao, J.; Collins, H. L.; Bates, S. L.; Roush, R. T.; Earle, E. D. & Shelton, A. M. 2005. Concurrent use of transgenic plants expressing a single and two Bacillus thuringiensis genes speeds insect adaptation to pyramided plants. Proc. Natl. Acad. Sci. USA, 102, 8426–8430.

Zhao, J. Z.; Cao, J.; Li, Y.; Collins, H. L.; Roush, R. T.; Earle, E. D. & Shelton, A. M. 2003. Transgenic plants expressing two Bacillus thuringiensis toxins delay insect resistance evolution. Nat. Biotechnol., 21, 1493–1497.

Zhao, Y.; Liu, Q. Z. & Davis, R. E. 2004. Transgene expression in strawberries driven by a heterologous phloem-specific promoter. Plant Cell Rep., 23, 224–230.

Zlotkin, E.; Fishman, Y. & Elazar, M. 2000. AaIT: From neurotoxin to insecticide. Biochimie, 82, 869–881.

Zlotkin, E.; Kadouri, D.; Gordon, D.; Pelhate, M.; Martin, M. F. & Rochat, H. 1985. An excitatory and a depressant insect toxin from scorpion venom both affect sodium conductance and possess a common binding site. Arch. Biochem. Biophys., 240, 877–887.

120 Appendices

Appendix I: Sequence of His6-Hv1a construct in yeast expression vector pPICZαA

Appendix II: Sequence of His6-GNA construct in yeast expression vector pPICZαA

Appendix III: Sequence of His6-Hv1a-GNA construct in yeast expression vector pPICZαA

Appendix IV: Sequence of His6-GNA-Hv1a construct in yeast expression vector pPICZαA

121 Appendix V: Sequence of Arabidopsis thaliana codon optimized ER-Hv1a construct in pMA-T vector

122 Appendix VI: Sequence of Arabidopsis thaliana codon optimized ER-GNA construct in pMA-T vector

123 Appendix VII: Sequence of Arabidopsis thaliana codon optimized ER-CP construct in pMA-T vector

Continued

124

125 Appendix VIII: Sequence of Arabidopsis thaliana codon optimized ER-Hv1a-GNA construct in pMA-T vector

126 Appendix IX: Sequence of Arabidopsis thaliana codon optimized ER-GNA-Hv1a construct in pMA-T vector

127 Appendix X: Sequence of Arabidopsis thaliana codon optimized ER-Hv1a-CP construct in pMA-T vector

Continued

128

129 Appendix XI: Sequence of Arabidopsis thaliana codon optimized ER-CP-Hv1a construct in pMA-T vector

Continued

130

131