INTERACTION BETWEEN NATURAL ENEMIES AND INSECTICIDES USED FOR THE MANAGEMENT OF WESTERN FLOWER THRIPS, FRANKLINIELLA OCCIDENTALIS (PERGANDE) (THYSANOPTERA: THRIPIDAE) IN THREE CULTIVARS OF STRAWBERRY, FRAGARIA X ANANASSA DUCHESNE (ROSACEAE)

Md Touhidur Rahman

B.Sc (Zoology), M.Sc (Zoology)

This thesis is presented for the degree of Doctor of Philosophy of The University of Western Australia

School of Animal Biology

July 2010 ABSTRACT

Interaction between natural enemies and insecticides used for the management of western flower thrips, Frankliniella occidentalis (Pergande) (Thysanoptera: Thripidae) in three cultivars of strawberry, Fragaria x ananassa Duchesne (Rosaceae)

Keywords: Frankliniella occidentalis, Typhlodromips montdorensis, Neoseiulus cucumeris,

Hypoaspis miles, strawberry IPM, host resistance, spinosad, residual toxicity, LT25, resistance

Integrated pest management (IPM) relies on the use of multiple tactics to reduce pest numbers below an economic threshold. One of the challenges for the implementation of IPM is using both insecticides and biological control. This is particularly difficult in horticultural crops where very little damage can be tolerated. Western flower thrips, Frankliniella occidentalis (Pergande) (Thysanoptera: Thripidae) is a worldwide pest of economic importance associated with cultivated crops, ornamentals and weeds. It is considered a major pest of strawberry, Fragaria x ananassa Duchesne (Rosaceae), and can be responsible for substantial yield loss. Insecticides are the main method of control for F. occidentalis in strawberry and other crops. Due to the rapid development of insecticide resistance and the limitations of existing biological control in Australia, there is a need to incorporate insecticides, natural enemies, and resistant host plants to keep the population below an economic threshold. This project sought to (i) evaluate commercial strawberry varieties for feeding and oviposition preferences of F. occidentalis, (ii) assess the compatibility of natural enemies with an insecticide currently used for F. occidentalis control in IPM programs, (iii) assess the effectiveness of the release of multiple species of natural enemies, (iv) determine the residual threshold of an insecticide which controls F. occidentalis effectively whilst having a reduced effect on natural enemies, and (v) assess the compatibility of natural enemies with an increased rate of an insecticide to manage an insecticide-resistant strain.

Frankliniella occidentalis showed a distinctive olfactory preference in a choice trial evaluating the feeding preference of F. occidentalis to strawberry cultivars (Camarosa, Albion and Camino Real). Frankliniella occidentalis was attracted most to Camarosa for both feeding and oviposition, followed by Albion and Camino Real. Frankliniella occidentalis also preferred to feed on fresh leaves to those that had been fed upon by a conspecific. Of the three varieties tested, Camino Real was the least preferred cultivar for oviposition. The development period of F. occidentalis (from eggs to adult emergence) was shortest in Camarosa and longest in Camino Real. Overall, of the three varieties tested, Camarosa appeared the most favourable for F. occidentalis feeding, oviposition and population growth, and the other cultivars might be a better choice for growers seeking to reduce F. occidentalis populations.

Spinosad (Success™; Dow AgroSciences Australia Ltd) is the only insecticide currently registered in Australia that is effective against F. occidentalis and regarded to be compatible in an integrated pest management program. A glasshouse study tested the compatibility of three predatory mite species, Typhlodromips montdorensis (Schicha) (Acari: Phytoseiidae), Neoseiulus cucumeris (Oudemans) (Acari: Phytoseiidae), Hypoaspis miles (Berlese) (Acari: Laelapidae) [commercially available in Australia for thrips management] and spinosad. All three predatory mites appeared to reduce the F. occidentalis population in strawberry plants. The efficacy of predatory mites further improved when combined with spinosad. Spinosad posed no detrimental effect to mites when an interval between spinosad application and the release of predatory mites was maintained. Releases of the two-species combination T. montdorensis and H. miles, or all three species (T. montdorensis, N. cucumeris and H. miles) combined with spinosad applications were more effective in reducing F. occidentalis than single species releases.

In Western Australia, strawberry is grown in low tunnels. Frankliniella occidentalis populations remain low during winter (June-August) and increase during spring (late September) to early summer. Therefore, the use of predatory mites prior to the increase in the F. occidentalis population in spring might be an approach to the management of F. occidentalis populations in the tunnel environment. A field trial revealed that predatory mites could be used to control F. occidentalis in low tunnel-grown strawberry plants in spring. Combined releases of ‘T. montdorensis and H. miles’ or ‘T. montdorensis, N. cucumeris and H. miles’ were most effective against F. occidentalis. Their beneficial effect was further increased when combined with spinosad. It was found that predatory mites performed better when released after a spinosad spray, compared to mites released before a spinosad application.

At the recommended spinosad application rate (80 mL/100 L, 0.096 g a.i./L) to control F. occidentalis in strawberry, residues were toxic to the predatory mites. Thresholds for the contact residual toxicity of spinosad LT25 (lethal time for 25% mortality) were estimated as 4.2 days (101.63h), 3.2 days (77.72) and 5.8 days (138.83 h) for T. montdorensis, N. cucumeris and H. miles respectively. The residual threshold increased when predatory mites were simultaneously fed spinosad-intoxicated F. occidentalis and exposed to residues. Residual thresholds then increased to 5.4 days (129.67), 4 days (95.09), and 6.1 days (146.68 h) for T. montdorensis, N. cucumeris, and H. miles respectively. Spinosad residues were also repellent to predatory mites. According to the standards of the International Organisation for Biological Control (IOBC) the

iii results of this study determined that spinosad is a short-lived chemical to N. cucumeris, and is slightly persistent to H. miles. On the other hand, the recommended rate of spinosad is a short- lived chemical to T. montdorensis, while with twice the recommended rate it was slightly persistent. The residual toxicity trial revealed that T. montdorensis, N. cucumeris and H. miles could be incorporated with a higher application rate of spinosad to combat against a spinosad- resistant strain of F. occidentalis, if the threshold period is maintained. Residual thresholds of twice the recommended rate of spinosad for T. montdorensis, N. cucumeris and H. miles were

6.1 days (146.76 h), 5.3 days (127.85 h), and 6.8 days (162.45 h) [LT25] respectively. Thus, predatory mites can be integrated with a higher application rate of spinosad if required, as long as the above-mentioned interval between application of spinosad and release of predatory mites is maintained.

This study contributes to an emerging body of research aimed at developing an integrated management strategy for F. occidentalis in strawberry crops. Collectively, the findings in this study suggest that existing biological control agents (predatory mites) can be integrated with spinosad for the management of F. occidentalis in glasshouse- and field (low tunnel)-grown strawberry. This management strategy can be further improved by selecting resistant cultivars that are less suitable to F. occidentalis. However, there is scope to further improve the effectiveness of this strategy. This includes further field trials, testing of additional cultivars, and testing different release strategies for the biological control agents.

iv ACKNOWLEDGEMENTS

First and foremost, I would like to extend my sincere gratitude to my supervisors Dr. Helen Spafford at The University of Western Australia (UWA), and Dr. Sonya Broughton at the Department of Agriculture and Food WA, for their advice and supervision. You both provided untiring and unerring guidance, valuable suggestions, and constructive criticism throughout this PhD. You have always made me think critically about every aspect of my work and I am immensely grateful for that. Thank you for making this thesis an enjoyable experience. I specially thank Helen for allowing me to come to Perth to follow my interest in integrated pest management.

Mr Anthony Yewers, of Berry Sweet, Bullsbrook, was kind enough to give me strawberry runners (one of the key components of this project) and allow me to conduct an experiment on his farm. Mr David Cousins, Department of Agriculture and Food WA, helped me to raise and maintain strawberry plants and provided me with valuable information and seedlings. It is my privilege to thank Manchil IPM Services, WA, Biological Services, SA and Beneficial Bug Company, NSW, for providing predatory mites.

I especially thank Kevin Murray, School of Mathematics and Statistics, UWA, for his great help in completing some of the statistical analyses presented in this thesis. I thank Peter Turner and Anna Williams for their friendship and advice, especially in my early days at UWA. I would also like to thank Peter Langland (School of Animal Biology), Sasha Voss (School of Forensic Entomology), Sayed Iftekhar (School of Agriculture Economics), Sharif-Ar-Raffi (School of Plant Biology), and many others at UWA. I benefited from them in many ways. I am also appreciative of my PhD Review Panel Members, Philip Withers (School of Animal Biology, UWA), and James Ridsdill Smith (CSIRO) for their advice, especially during the developmental stage of my project.

My research was funded by UWA, through the University Postgraduate Awards (UPA) and Scholarship for International Research Fees (SIRF). I also acknowledge add hoc scholarship from the School of Animal Biology, UWA, and an ad hoc scholarship from Helen Spafford. Without this financial support, it would not have been possible to complete this project.

Last but not least, I would like to thank my family and friends from Bangladesh. I am immensely grateful to my beloved parents and all family members for their inspiration, wishes and blessings during the entire period of the study. They never let me doubt my ability to achieve my goals. I dedicate this thesis to all my family members, but most especially to my late father, Mr Badiur Rahman. ORGANISATION AND DECLARATIONS

The research presented in this thesis is an original contribution to the integrated management of western flower thrips in glasshouse- and low tunnel-grown strawberry.

This thesis is presented as a series of independent research papers (to be published), preceded by a general introduction chapter, and followed by general discussion. The central chapters of this thesis are written and presented as separate manuscripts, so some repetition of basic information does occur.

I, Touhidur Rahman, carried out the design and conducted the experiments after consultation with my supervisors, Dr Helen Spafford and Dr Sonya Broughton. All statistical analyses performed in this thesis were produced by myself after discussions with and review by my supervisors. I benefited from advice and assistance with the design, laboratory, glasshouse and fieldwork, analyses and writing from my supervisors and others as duly acknowledged.

I, Touhidur Rahman, certify that this thesis does not incorporate, without acknowledgement, any material previously submitted for a degree or diploma in any institute, and that it does not contain any material previously published or written by another person, except where due reference is made in the text.

______Touhidur Rahman TABLE OF CONTENTS

ABSTRACT ii ACKNOWLEDGEMENTS v ORGANIZATION AND DECLARATION vi TABLE OF CONTENTS vii LIST OF FIGURES xi LIST OF TABLES xv

Chapter I: General introduction and literature review Introduction 1 Integrated pest management (IPM) 2 Western flower thrips: origin and distribution 4 Western flower thrips: host range, pest status and damage 5 Chemical control of western flower thrips and insecticide resistance 8 Biological control of western flower thrips 12 Thrips predators 12 Predatory mites 14 Thrips parasitoids 15 Fungi 16 Biological control of western flower thrips in Australia 16 Host-plant resistance 17 Cultural methods of control of western flower thrips 18 IPM programs for control of western flower thrips 18 Outline of this study 20 Brief organisation and structure of this thesis 22 Literature cited 23

Chapter II: Variation in preference and performance of western flower thrips, Frankliniella occidentalis (Pergande) (Thysanoptera: Thripidae) on three strawberry [Fragaria x ananassa Duchesne (Rosaceae)] cultivars Keywords 43 Abstract 43 Introduction 43 Materials and methods 45 Source cultures 45 Strawberry cultivars 45 Western flower thrips (WFT) 46 Experiment 1: Feeding preference of adult WFT 46 Experiment 2: Oviposition preference and performance of WFT on caged plants 47 Experiment 3: Oviposition preference and performance of WFT on leaf discs 48 Egg hatch 48 Larval mortality, pupal mortality and adult emergence rate 48 Developmental time 49 Data analysis 49 Results 50 Experiment 1: Feeding preference of adult WFT 50 Experiment 2: Oviposition preference and performance of WFT on caged plants 54 Experiment 3: Oviposition preference and performance of WFT on leaf discs 56 Discussion 59 Literature cited 61

vii Chapter III: Effect of spinosad and predatory mites (Acari) on western flower thrips, Frankliniella occidentalis (Pergande) (Thysanoptera: Thripidae) in three strawberry cultivars [Fragaria x ananassa Duchesne (Rosaceae)] Keywords 65 Abstract 65 Introduction 65 Materials and methods 68 Source cultures 68 Strawberry cultivars 68 Western flower thrips (WFT) 69 Predatory mites 69 Glasshouse experiment: effect of cultivars and predatory mites with or without 70 spinosad on western flower thrips Data analysis 71 Results 72 Western flower thrips 72 Predatory mites 80 Discussion 81 Literature cited 84

Chapter IV: Single versus multiple releases of predatory mites (Acari) combined with a spinosad application for the management of western flower thrips, Frankliniella occidentalis (Pergande) (Thysanoptera: Thripidae) in strawberry [Fragaria x ananassa Duchesne (Rosaceae)] Keywords 90 Abstract 90 Introduction 91 Materials and methods 92 Source cultures 93 Strawberry cultivar 93 Western flower thrips (WFT) 93 Predatory mites 94 Experiment: effect of single versus multiple species releases of mites combined 94 with spinosad on WFT Data analysis 95 Results 96 Western flower thrips 96 Adults 96 Larvae 98 Predatory mites 100 Discussion 102 Literature cited 104

Chapter V: Use of spinosad and predatory mites (Acari) for the management of western flower thrips, Frankliniella occidentalis (Pergande) (Thysanoptera: Thripidae) in strawberry [Fragaria x ananassa Duchesne (Rosaceae)]: a field study Keywords 109 Abstract 109 Introduction 110 Materials and methods 112 Study site 112 Predatory mites 113 Treatments 113 Pre-treatment sampling 114 Post-treatment sampling 115 Data analysis 115

viii Results 116 Impact of the spray and predatory mite species combinations on WFT adults 117 Flower 117 Fruit 118 Impact of the spray and predatory mite species combinations on WFT larvae 120 Flower 120 Fruit 122 Impact of the spray and mite species combinations on predatory mites 124 Single species 124 Two-species combinations 126 Three-species release 128 Species interactions 129 Discussion 130 Literature cited 135

Chapter VI: Compatibility of spinosad with predaceous mites (Acari) used to control western flower thrips, Frankliniella occidentalis (Pergande) (Thysanoptera: Thripidae) Keywords 141 Abstract 141 Introduction 141 Materials and methods 143 Source cultures 143 Strawberry plants 143 Western flower thrips (WFT) 143 Predatory mites 143 Experiment 1: Direct toxicity of spinosad to WFT and predatory mites 144 Western flower thrips 144 Predatory mites 145 Experiment 2: Residual toxicity of spinosad to WFT and predatory mites 145 Mortality of WFT and mites to spinosad residues (contact) over time 145 Indirect exposure of spinosad to predatory mites via consumption of 146 intoxicated WFT larvae Toxicity of spinosad to predatory mites via consumption of intoxicated 147 WFT larvae and direct exposure to spinosad residues of different ages Experiment 3: Repellency of spinosad to predatory mites (choice test) 148 Data analysis 148 Results 150 Experiment 1: Direct contact toxicity of spinosad to WFT and predatory mites 150 Experiment 2: Residual toxicity of spinosad to WFT and predatory mites 150 Residual (contact) toxicity of spinosad to WFT and predatory mites 150 Indirect exposure of spinosad to predatory mites via consumption of 154 intoxicated WFT larvae Residual toxicity to predatory mites via consumption of spinosad- 157 intoxicated WFT larvae and direct exposure to spinosad residues Experiment 3: Repellency of spinosad to predatory mites (choice test) 159 Discussion 160 Literature cited 162

Chapter VII: Spinosad-resistant western flower thrips, Frankliniella occidentalis (Pergande) (Thysanoptera: Thripidae) can be managed using spinosad and predatory mites (Acari) Keywords 166 Abstract 166 Introduction 166 Materials and methods 168 Source cultures 168

ix Strawberry plants 168 Western flower thrips (WFT) 168 Predatory mites 169 Experiment 1: Direct toxicity of spinosad 169 Western flower thrips 169 Predatory mites 170 Experiment 2: Bioassay of spinosad residual toxicity to predatory mites 170 Experiment 3: Efficacy of predatory mites with spinosad against WFT-resistant 172 strain Data analysis 173 Results 175 Direct toxicity of spinosad to WFT and predatory mites 175 Bioassay of spinosad residual toxicity to predatory mites 176 Efficacy of predatory mites with spinosad against WFT-resistant strain 180 Effect of spinosad and predatory mite releases on WFT adults 180 Effect of spinosad and predatory mite releases on WFT larvae 181 Discussion 183 Literature cited 185

Chapter VIII: General discussion and conclusion General discussion 189 Findings and recommendations 190 Strawberry cultivars distinctively influence western flower thrips’ olfactory and 190 feeding preference and oviposition preference and performance Biological control: multiple species versus single species 190 Combining chemical and biological control 191 IPM of western flower thrips in low tunnel-grown strawberry 192 Control of spinosad-resistant western flower thrips strain 193 Conclusions 194 Literature cited 195

Appendices xvii

x LIST OF FIGURES

Figure captions Page no

Figure 1.1 Distribution map of Frankliniella occidentalis (Pergande) compiled by 5 CAB International in association with the European and Mediterranean Plant Protection Organization (EPPO).

Figure 1.2 (A) Western flower thrips adults, and predatory mites, (B) Typhlodromips 6 montdorensis (Schicha), (C) Neoseiulus cucumeris (Oudemans) and (D) Hypoaspis miles (Berlese).

Figure 1.3 Damage of WFT on strawberry fruit and flower. 7

Figure 2.1 Olfactory preference of WFT adults to strawberry cultivars when offered 51 (A) ungrazed and (B) grazed (B) leaf discs.

Figure 2.2 Simplex plot showing time spent by WFT adults between strawberry 52 cultivars when exposed to ungrazed leaf discs and discs previously grazed by conspecific.

Figure 2.3 Time spent by WFT adults on (A) ungrazed and (B) previously grazed 52 leaf discs.

Figure 2.4 Preference of WFT adults when given a choice between leaf discs 53 exposed to conspecifics (grazed) or ungrazed of three different strawberry cultivars.

Figure 2.5 Time [A = total time, B = time spent feeding] spent by WFT adult on 54 ungrazed and grazed leaf discs.

Figure 2.6 Mean numbers of WFT adults per plant on caged strawberry cultivars at 55 1, 24, and 48 h post-release.

Figure 2.7 Comparison of WFT adult numbers (Y-axis) at different post-release 55 periods (hours) on caged plants.

Figure 2.8 Mean numbers of WFT (A) larvae hatched and (B) adults emerged per 56 plant on caged strawberry cultivars.

Figure 2.9 Comparison of numbers of eggs laid, unhatched eggs, larvae hatched, 57 pupae developed and adults emerged per leaf disc (Y-axis) on three strawberry cultivars.

Figure 2.10 Comparison of the percentage of unhatched eggs (U/E), larvae hatched 57 (L/H), larvae killed (L/K), pupae developed (P), pupae killed (P/K) and adult emerged (A/E) per leaf disc among cultivars.

Figure 2.11 Comparison of survival rate of WFT among strawberry cultivars. 58

Figure 2.12 Egg incubation period (IP), larval period (LP), prepupation period 58 (PPP), pupation period (PP) and total developmental period (TDP) (egg to adult) of WFT in days, among strawberry cultivars.

xi Figure 3.1 Plant covered by a modified cage made from thrips-proof mesh 68 (105µ; Sefar Filter Specialists Pty Ltd., Malaga)

Figure 3.2 Mean numbers of WFT adults per plant treated with spinosad or 73 water and either no predatory mites or one of three species of predatory mites.

Figure 3.3 Effects of predatory mites on number of WFT adults per plant over 74 time (7, 14, or 21 days after release of WFT) sprayed with spinosad on cultivars (A) Camarosa, (B) Camino Real and (C) Albion.

Figure 3.4 Effects of predatory mites on the number of WFT adults per plant 76 over time (7, 14, or 21 days after release of WFT) sprayed with water on strawberry cultivar (A) Camarosa, (B) Camino Real and (C) Albion.

Figure 3.5 Mean numbers of WFT larvae per plant treated with spinosad or 77 water and either no predatory mites or one of three species of predatory mites.

Figure 3.6 Effects of predatory mites on WFT larvae per plant over times (7, 78 14, or 21 days after WFT release) sprayed with spinosad on strawberry cultivar (A) Camarosa, (B) Camino Real and (C) Albion.

Figure 3.7 Effects of predatory mites on WFT larvae per plant over times (7, 79 14, or 21 days after WFT release) sprayed with water on strawberry cultivar (A) Camarosa, (B) Camino Real and (C) Albion.

Figure 3.8 Mean numbers of T. montdorensis (A and B) and N. cucumeris (C 80 and D) per plant in relation to strawberry cultivar and spray treatment

Figure 3.9 Comparison of mean numbers of T. montdorensis and N. 81 cucumeris per plant sprayed with (A) spinosad and (B) water.

Figure 4.1 Comparison of mean number of WFT adults per plant sprayed with 97 either spinosad or water and in the presence of no mites or different mite combinations.

Figure 4.2 Effects of predatory mites on mean number of WFT adults per 98 plant sprayed with (A) spinosad or (B) water.

Figure 4.3 Comparison of mean number of WFT larvae per plant sprayed with 98 either spinosad or water and in the presence of no mites or different mite combinations.

Figure 4.4 Effects of predatory mites on mean number of WFT larvae per 99 plant sprayed with (A) spinosad or (B) water.

Figure 4.5 Comparison of mean number of T. montdorensis and N. cucumeris 101 per plant applied with (A) single-species releases of T. montdorensis and N. cucumeris, (B) double-species releases of T. montdorensis and N. cucumeris, (C) double species releases of T. montdorensis and H. miles and N. cucumeris and H. miles, and (D) triple-species release.

Figure 4.6 Mean numbers of (A) T. montdorensis and (B) N. cucumeris per 102 plant in double-species combinations.

Figure 5.1 Maximum, minimum and average daily air temperature (⁰C), and average 113 daily temperature (⁰C) inside low tunnel (25 September to 20 November 2008).

xii Figure 5.2 Effect of spray treatment and predatory mite species releases on the 117 number of WFT adults/flower in low tunnel strawberry.

Figure 5.3 Influence of predatory mite species combinations on WFT adults per 118 flower over time (X-axis) in (A) water, (B) ‘spinosad then mites’ and (C) ‘mites then spinosad’.

Figure 5.4 Effect of spray treatment and predatory mite species combinations (X- 119 axis) on the number of WFT adults per fruit (Y-axis).

Figure 5.5 Influence of predatory mites on WFT adults per fruit over time (X-axis) 120 in (A) water, (B) ‘spinosad then mites’ and (C) ‘mites then spinosad’.

Figure 5.6 Effect of spray treatment and predatory mite species combinations (X- 121 axis) on the number of WFT larvae per flower (Y-axis).

Figure 5.7 Influence of predatory mites on WFT larvae per flower over time (X-axis) 122 in (A) water, (B) ‘spinosad then mites’ and (C) ‘mites then spinosad’.

Figure 5.8 Effect of spray treatment and predatory mite species combinations (X- 123 axis) on the number of WFT larvae per fruit.

Figure 5.9 Influence of predatory mites on WFT larvae per fruit over time (X-axis) 124 in (A) control (water), (B) ‘spinosad then mites’ and (C) ‘mites then spinosad’.

Figure 5.10 Comparison of mean number of T. montdorensis (Tm) and N. cucumeris 125 (Nc) per flower or fruit, when applied alone (single species).

Figure 5.11 Comparison of mean numbers T. montdorensis (Tm) and N. cucumeris 127 (Nc) applied in double-species combination as (A) ‘T. montdorensis and N. cucumeris’ and (B) T. montdorensis in ‘T. montdorensis and H. miles’ and N. cucumeris in ‘N. cucumeris and H. miles’.

Figure 5.12 Comparison of mean numbers of T. montdorensis (Tm) and N. 128 cucumeris (Nc) when applied as a three-species combination (T. montdorensis, N. cucumeris and H. miles).

Figure 5.13 Comparison of mean number of (A) T. montdorensis and (B) N. 129 cucumeris when applied in double-species combination (X-axis).

Figure 6.1 Diagrammatic representation of the testing arena used for toxicity test. 144

Figure 6.2 Probit mortality of (A) T. montdorensis, (B) N. cucumeris, and (C) H. 154 miles recorded against spinosad residues of different ages (log10 hrs).

Figure 6.3 Probit mortality of (A) T. montdorensis, (B) N. cucumeris, and (C) H. 159 miles recorded against spinosad residues of different ages (log10 hrs).

Figure 7.1 Toxicity of spinosad residues to predatory mites after 96 h post-release 176 exposure period.

Figure 7.2 Relationship of Log10 (hrs) and probit mortality of (A) T. montdorensis, 179 (B) N. cucumeris and (C) H. miles when exposed to twice the recommended rate of spinosad residues with different ages.

Figure 7.3 Mean numbers of WFT adults per plant sprayed with spinosad or water 180

xiii (control) in the presence or absence of predatory mite.

Figure 7.4 Numbers of WFT adults per plant (Y-axis) sprayed with spinosad (A) or 181 water (B) with or without predatory mite.

Figure 7.5 Numbers of WFT larvae per plant with spinosad and water (control) in 182 the presence or absence of predatory mites.

Figure 7.6 Numbers of WFT larvae per plant with spinosad (A) and water (B) in the 183 presence or absence of predatory mites.

xiv LIST OF TABLES

Table captions Page no

Table 1.1 Integration of natural enemies and pesticides registered for glasshouse 3 vegetables in the Netherlands.

Table 1.2 Insecticides that have been tests for efficacy against WFT. 9

Table 1.3 Natural enemies evaluated as biocontrol agents of WFT. 13

Table 1.4 Crops and predatory mites used for WFT control. 15

Table 5.1 Schedules of treatment applications and sampling. 114

Table 6.1 Mean (±SE) corrected mortality (%) at different post-release exposure 150 periods (h) to predatory mites after direct exposure to spinosad (recommended rate, 80 mL/100 L).

Table 6.2 Mean (± SE) corrected mortality (%) of WFT adults and larvae when 151 exposed to spinosad residues aged 2, 12, 24, 48, 72, 96, and 120 h at different post- release exposure periods.

Table 6.3 Residual toxicity (contact) of spinosad to predatory mites at 24, 48, and 72 152 h post release exposure periods. The IOBC classification: 1 = harmless (<25% mortality), 2 = slightly harmful (25-50% mortality), 3 = moderately harmful (51- 75% mortality) and 4 = harmful (>75% mortality). Persistence class: A = short-lived (<5 d), B = slightly persistent (5-15 d).

Table 6.4 Probit analysis (Abbott 1925) of the mortality of adult predatory mites 153 exposed to spinosad residues of different ages.

Table 6.5 Mortality of predatory mites after feeding on spinosad intoxicated WFT 156 larvae at 24 h, 48 h, and 72 h post-release exposure periods. The IOBC classification: 1 = harmless (<25% mortality), 2 = slightly harmful (25-50% mortality), 3 = moderately harmful (51- 75% mortality) and 4 = harmful (>75% mortality).

Table 6.6 Residual toxicity of spinosad to predatory mites at 24 h, 48 h, and 72 h 158 post release exposure periods. Mites were fed spinosad-intoxicated WFT larvae and simultaneously exposed to residue. The IOBC classification: 1 = harmless (<25% mortality), 2 = slightly harmful (25-50% mortality), 3 = moderately harmful (51- 75% mortality) and 4 = harmful (>75% mortality). Persistence class: A = short lived (<5 d), B = slightly persistence (5-15 d).

Table 6.7 Probit analysis (Abbott 1925) of the mortality of adult predatory mites 159 simultaneously exposed to spinosad via consumption of intoxicated WFT larvae and contact with spinosad residues of different ages.

Table 6.8 Mean (±SE) numbers of predatory mites on spinosad- and water-treated 160 strawberry leaf in a choice test (t-test, df = 19).

Table 7.1 Cumulative corrected mortality (%) of spinosad- resistant WFT adults (at 175 96 h post-release exposure period) and larvae (at 72 h post-release-exposure period)

xv when exposed directly to spinosad spray at different rates.

Table 7.2 Residual toxicity of spinosad (twice the recommended rate) to predatory 177 mites at 24 h, 48 h, 72 h, and 96 h post-release exposure periods. Mites were fed spinosad intoxicated WFT larvae and simultaneously exposed to residue. Residual toxicity were classified: 1 = harmless (<25% mortality), 2 = slightly harmful (25- 50% mortality), 3 = moderately harmful (51-75% mortality), and 4 = harmful (>75% mortality). Persistence class: A = short lived (<5 d), B = slightly persistent (5-15 d).

Table 7.3 Probit analysis (Abbott 1925) of the mortality of predatory mites exposed 179 to spinosad residues (by consumption of intoxicated WFT larvae and simultaneous exposure to residues) of different ages.

xvi CHAPTER I

General introduction and literature review

1.1 Introduction

There are more than 6.3 billion people worldwide (FAO 2004) and the human population continues to expand. Thus, there is an increased need for food and this requires large increases in agricultural production. One of the major threats to human food production is insect pests. It is estimated current food production worldwide is worth about US$1.3 trillion, with insect pests causing crop losses estimated at US$500 billion (FAO 2005). Currently there are 10,000 species of insects considered to be pests of agricultural and horticultural crops (Reuveni 1995). Reuveni (1995) suggests that the total elimination of pests is not possible, but attempts have been made to reduce their detrimental effects on plants. In this regard, chemical control of the pest population has become the principal control tool. Since the first synthetic pesticides were produced in the 1940s, pesticide usage has increased fiftyfold. Currently 2.5 million tons of pesticides (US$20 billion) are used each year to combat agricultural pest species worldwide (Pimentel et al. 1992). Unfortunately, due to a lack of sufficient alternatives and a lack of grower knowledge, synthetic pesticides are often used prophylactically, too heavily or inappropriately, and can result in the development of resistant pest populations (Maredia et al. 1992, Eddleston et al. 2002). Currently more than 700 arthropod pest species have been reported to develop resistance to one or more pesticides (Thacker 2002). Furthermore, pesticides pose detrimental effects to naturally occurring or inundative biological control agents, which can lead to secondary pest outbreaks. Therefore, an estimated US$520 million can be attributed for the additional use of pesticides to control secondary pest populations (Thacker 2002).

In response to growing pest and associated problems and risks with pesticide use, the agricultural industry has been challenged to develop and implement an integrated approach to pest management (Stern et al. 1959). One of the other pest management tactics available for many commodities and pests is biological control. However, the integration of biological control and pesticide use in a pest management program can be problematic. For example, spinosad is a biopesticide but it is highly toxic to the predatory mite Neoseiulus fallacies (Garman) (Villanueva and Walgenbach 2005), and slightly toxic to Amblyseius cucumeris (Oudemans) (Jones et al. 2005). Thus, before implementation in pest management programs, compatibility of chemical and biological control agents need to be assessed.

Chapter I: General introduction and literature review

This thesis presents the results of a comprehensive study to assess the compatibility of chemical and biological control for the management of a major economic pest, western flower thrips (WFT), Frankliniella occidentalis (Pergande) (Thysanoptera: Thripidae) on strawberry Fragaria ananassa Duchesne (Rosaceae). When WFT was first detected in Australia in 1993, there were no biological control agents commercially available and control relied solely on chemicals. Since then, three predatory mite species have become commercially available in Australia. As with control of WFT in other parts of the world, the challenge is to integrate chemical and biological control methods in a way that provides an economic reduction in the pest population. The present project tested the hypothesis that integration of an insecticide (spinosad) with predatory mite releases [Typhlodromips montdorensis (Schicha) (Phytoseiidae), Neoseiulus cucumeris (Oudemans) (Phytoseiidae), and Hypoaspis miles (Berlese) (Laelapidae)] can improve the management of WFT in crops such as strawberry. WFT has developed resistance to multiple insecticides currently in use. Successful integration of chemical and biological control would minimise the use of chemicals. The ultimate aim of this approach is to slow the inevitable evolution of resistance and keep insecticides available to growers. This study also tested whether multiple releases of predatory mites would provide better WFT management than single species releases. In addition, the influence of strawberry cultivar on WFT preference and performance is tested.

1.2 Integrated pest management (IPM)

The term „integrated pest management‟ (IPM) [first appeared in literature in 1967 (Smith and van den Bosch 1967)] is the use of a range of tactics including biological control, host plant resistance, cultural control, physical control and minimal use of pesticides (Kogan 1998, Trumble 1998, Hillocks 2002, Prokopy and Kogan 2003, Feder et al. 2004) to control pest populations. The United Nations Development Program (UNDP) together with the Food and Agriculture Organization (FAO) has initiated global programs for the development and application of IPM in several crops such as rice, cotton, sorghum, millet and vegetable crops (Peshin et al. 2009). Development of IPM strategies emerged in the USA in the 1950s to improve pest management in agriculture. In Australia, IPM programs have been developed and implemented in several crops. Australian examples include the use of classical and inundative releases of biocontrol agents combined with limited use of selective pesticides for the control of scale insect and mite pests in citrus (Smith et al. 1997), conservation of native predatory mites to reduce mite pests in grapes (James and Whitney 1993), predatory mites releases against mite pests, mating disruption and selective insecticide against moth pests in apple (Thwaite 1997), and the management of currant lettuce aphid, Nasonovia ribisnigri (Mosley) in lettuce using Nasonovia-resistant varieties and selective insecticides (McDougall and Creek 2007).

2 Chapter I: General introduction and literature review

IPM adoption has resulted in a decrease in pesticide use from 2.6 billion kg in 2004 (Allan Woodburn Associates 2005) to 1.7 billion kg in 2007 (Agronova 2008) worldwide. In the USA, adoption of IPM strategies have saved US$500 million each year (Peshin et al. 2009); no figures are available for Australia. Despite demonstrated efficacy and cost savings, pesticides remain and will continue to remain the dominant component in many cropping systems (Dhawan and Peshin 2009). This is because zero damage is tolerated by some markets (Nothnagl 2006), alternative control may not be available for some crops and pests, and biological control is not always effective when pest densities are very high (Malezieux et al. 1992). Moreover, biological control is expensive compared to chemical control, even for growers of high value crops (Reuveni 1995). Therefore, one of the biggest challenges of IPM is the use of biological control agents in conjunction with pesticides.

The first successful integration of biological control agents with pesticides dates back to the 1960s, with the use of dimethirimol (fungicide) integrated with releases of the predatory mite, Phytoseiulus persimilis Athias-Henriot (Acari: Phytoseiidae), for control of two spotted mite in cucumber (Reuveni 1995). Since then, the integrated use of pesticides combined with biological control has been practised in many cropping systems (Table 1.1).

Table 1.1 Integration of natural enemies and pesticides registered for glasshouse vegetables in the Netherlands after Reuveni (1995).

Pest Beneficials Acaricides/insecticides Remarks Spider mites Predatory mites Fenbutatin oxide, - hexythiazox, clofentezine Whiteflies Hymenopterous Buprofezin Not effective against parasitoid Bemisia tabaci and some strains of Trialeurodes vaporariorum Teflubezuron, Hydrogen Harmful to anthocorids cyanide Harmful to beneficial insects

Effective against Liriomyza Leafminers Hymenopterous Oxamyl bryoniae only, selective if parasitoid used systemically

- Predatory mites, - Thrips anthocorid bugs Not effective against Aphis Hymenopterous Pirimicarb Aphids gossypii and some strains parasitoid, of Myzus persicae cecidomyiids As above lacewings, Hydrogen cyanide

coccinellids Noctuids As above Egg parasitoid Teflubenzuron

3 Chapter I: General introduction and literature review

However, pesticide applications are often disruptive to natural enemies and consequently pest populations may increase to more damaging levels than before treatment occurred (Croft 1990). Thus, integration of biological control agents with chemical control will not be successful unless natural enemies survive the pesticides being used in a cropping system (Hoy 1985, Hoy and Cave 1985). By limiting exposure of a beneficial to a pesticide, using narrow-spectrum pesticides or biological pesticides, this problem can be reduced (Croft 1990, Greathead 1995). Tauber et al. (1985) stated, "It is probable that the most dramatic increase in the utilisation of biological control in agricultural IPM systems could come through the judicious use of selective pesticides in conjunction with effective natural enemies in specific cropping systems, in specific geographic regions ….”

In order to develop and implement an effective management strategy, it is necessary to have information about the pest, the available control strategies and advantages and disadvantages associated with different control methods. The remainder of this chapter reviews the biology and management of WFT, and the challenges associated with the management of WFT in strawberry crops.

1.3 Western flower thrips: origin and distribution

Of more than 5500 species of thrips (Thysanoptera) identified to date (Morse and Hoddle 2006), a few hundred species, mainly from the family Thripidae, are of economic importance. Of them, a number of species have become key pests in a wide range of agricultural and horticultural crops, including WFT. WFT has a cosmopolitan distribution (Figure 1.1) and was first described in 1895 from specimens collected in California, on apricot, potato, orange and various weeds (Pergande 1895). Pergande placed WFT in the genus Euthrips, but in 1912, Karny (1912) moved this species to the newly erected genus Frankliniella. Until 1960, the distribution of the WFT was supposedly restricted to western North America, Mexico, and Alaska (Bryan and Smith 1956, Nakahara 1997). However, old records indicate the presence of WFT in New Zealand in 1934 (Mound and Walker 1982). Since the 1960s, there has been a dramatic increase in the transport of plant materials throughout the world and WFT has been accidentally moved (Kiritani 2001). In 1966, it was recorded for the first time on chrysanthemum in a glasshouse in Pennsylvania [Greenough et al. (1985) cited by Kirk and Terry (2003)]. Subsequently, WFT was recorded on marijuana in Kansas in 1971, and on African violet in Missouri in 1973, in North Carolina in 1977 and in Louisiana in 1983 [Greenough et al. (1985) cited by Kirk and Terry (2003)]. In the early 1980s, WFT was recorded in glasshouses in Canada, where it caused severe epidemics of Tomato spotted wilt virus (Broadbent et al. 1987).

4 Chapter I: General introduction and literature review

Outside North America, WFT was first recorded in New Zealand in 1934 on lupin (Mound and Walker 1982, Zur Strassen 1986), but was not considered a pest there during that time. It was reported from Hawaii in 1955 (Sakimura 1962). In 1983 WFT entered Europe through the Netherlands (Mantel 1989) and spread quickly across the European continent (Tommasini and Maini 1995). In the late 1980s, WFT was detected in Eastern Africa (Nakahara 1997), Colombia, Costa Rica (Baker and Hurd 1968), and South Africa (Gilmore 1989). More recently it has been recorded from Japan (Hayase and Fukuda 1991), Brazil [1994, (Monterio et al. 2001) cited in Kirk and Terry (2003)], Argentina [1993, (De Santis 1995) cited in Kirk and Terry (2003)] and Chile [1995, (Gonzalez 1996)]. In Australia, WFT was first recorded in 1993 on glasshouse-grown chrysanthemum in Yangebup, Western Australia (Malipatil et al. 1993). WFT damage to strawberry was detected in Albany, Western Australia in 1994 (Steiner and Goodwin 2005)]. This detection was followed by the detection of WFT on strawberry in the Sydney Basin, New South Wales in 1995, the Yarra Valley, Victoria and the Adelaide Hills, South Australia in 1998-1999 (Steiner and Goodwin 2005)]. WFT has since spread to all states except the Northern Territory (Medhurst and Swanson 1999).

Figure 1.1 Distribution map of Frankliniella occidentalis (Pergande) compiled by CAB International in association with the European and Mediterranean Plant Protection Organization (EPPO) (CAB International 1999). ● Present: national record, + Present: subnational record.

1.4 Western flower thrips: host range, pest status and damage

WFT is a worldwide pest (Figure 1.1 and 1.2) of economic importance associated with cultivated crops, ornamental, and wild plants (Lewis 1997b). It is highly polyphagous, adults of WFT having been reported from at least 240 species from 62 different families of plants (Lewis 1997b). Abundance, biology, and developmental stages of thrips can vary under different ecological conditions and thus, species compositions of thrips, their occurrence, abundance, and

5 Chapter I: General introduction and literature review life cycles on plants may differ. Common hosts of WFT include cotton, onion, strawberry, cabbage, lettuce, capsicum (sweet pepper), tomato, chrysanthemum, gerbera, rose, cucumber, eggplant, bean, geranium, apple, nectarines, peach, and table grapes [(Elmore 1949, Hightower and Martin 1956, Allen and Gaede 1963, Oatman and Patner 1969, Zur Strassen 1986, van de Veire 1987, Brødsgaard 1989b, Buxton and Wardlow 1991) cited in Tommasini (2003)]. In Australia, WFT utilises a wide range of host plants including ornamentals, fruit crops, vegetables and weeds (Persley et al. 2009).

Figure 1.2 (A)Western flower thrips adults, and predatory mites, (B) Typhlodromips montdorensis (Schicha), (C) Neoseiulus cucumeris (Oudemans) and (D) Hypoaspis miles (Berlese) [Photos: Sonya Broughton and James Altman].

WFT causes substantial crop losses through (i) direct damage (feeding and oviposition), and (ii) vectoring plant viruses and bacterial diseases (Lewis 1997b). Like all other thrips species, WFT uses its piercing-sucking stylets (Hunter and Ullman 1989) to penetrate epidermal and sub- epidermal cells, causing extensive damage to the tissue of leaves, fruits and petals (Kirk 1997). The degree of feeding damage depends on the plant tissue that is affected, the developmental stage of the plant, and the susceptibility of the cultivars or the plant species attacked (Childers and Achor 1995, Childers 1997). Damage includes deformation and growth reduction of the plant, and silver scars on fruits and leaves (Hunter and Ullman 1989, van Dijken et al. 1994, de Jager et al. 1995). Scars caused by WFT often induce aesthetic injury to fruits and flowers,

6 Chapter I: General introduction and literature review making them less valuable or even unmarketable (Parrella and Jones 1987). Frequent symptoms of WFT damage on ornamentals (e.g. rose, gerbera, chrysanthemum, carnation, geranium, pansy, marigold etc) are streaking, browning, and distortion of leaves, petals and even buds (Chisholm and Lewis 1984, Oetting et al. 1993, Harrewijn et al. 1996, Childers 1997). Feeding on fruits by WFT causes scarring, fruit malformation, and russetting. WFT feeding on immature cucumber causes silvery scarring or malformation (Rosenheim et al. 1990, Shipp et al. 2000a), minute scarring on immature nectarine fruits which can develop into serious surface russetting on mature fruits. Fruits with minor feeding damage may be downgraded at sale, but fruits displaying serious damage are culled (Pearsall 2000). Feeding on the plant foliage by WFT negatively affects leaf size and photosynthesis, which can cause significant yield loss (Welter et al. 1990, Shipp et al. 1998, Shipp et al. 2000a). In addition to feeding damage, WFT can cause yield loss by oviposition. As in all Terebrantia thrips, the female of WFT features a saw-like ovipositor with which it drills holes into the parenchymal tissues of leaves, flowers and fruits, where it deposits kidney-shaped opaque eggs (Brødsgaard 1989b). Oviposition injury by WFT can result in economic loss in some plants, whilst producing no measurable damage to others (Brødsgaard 1989b). WFT causes a central russet area surrounded by a white halo on apples due to oviposition injury (Terry and DeGrandi-Hoffman 1988, Terry 1991). This condition is known as „pansy spot‟. It has also been reported that oviposition injury of WFT causes surface deformation in avocado fruit (Fisher and Davenport 1989), black dark scars surrounded by halos of whitish tissue in grape berries [(Jensen 1973) cited in Childers (1997)], and pale halo surrounded with puncture marks in tomatoes (Salguero Navas et al. 1991). Plant damage is aggravated in dry conditions, especially under glasshouse conditions when heavily infested plants lose moisture rapidly, which can seriously reduce yields and sometimes render crops uneconomic (Lewis 1997a).

Figure 1.3 Damage of WFT on strawberry fruit and flower [Photo by Sonya Broughton and Paul Horne].

In addition to direct damage, WFT transmits tospoviruses such as Chrysanthemum stem necrosis virus, Groundnut ringspot virus, Impatiens necrotic spot virus, Tomato chlorotic spot

7 Chapter I: General introduction and literature review virus and Tomato spotted wilt virus (Whitfield et al. 2005). Of these, TSWV is considered a serious disease in several economically important crops worldwide (Cho et al. 1988). TSWV was first described in 1915 in Australia and has been spread worldwide (McDougall and Tesoriero 2007). Currently, 1090 plants species in 85 families are hosts of TSWV (Parrella et al. 2003). WFT can also cause secondary infections on plants by transmitting pathogenic fungi or bacteria (Lewis 1973, Lewis 1997b).

Although accurate information is difficult to obtain, WFT is thought to cause billions of dollars in yield loss year. For example, in Florida annual crop losses due to WFT and Thrips palmi exceeded US$10 million (Nuessly and Nagata 1995). In the UK, it is estimated that WFT can cause crop losses of up to US$76,000 ha-1 each year in glasshouse-grown cucumbers (Nuessly and Nagata 1995). Annual yield loss attributed to TSWV are US$1 billion alone in the USA (Goldbach and Peters 1994). As a result, growers spend millions of dollars to manage WFT. Between 1987 to 1990, the Finland Government spent US$390,000 to eradicate WFT from glasshouse-grown vegetables and ornamentals (Lewis 1997a)].

In this project, strawberry, Fragaria ananassa Duchesne (Rosaceae) was used as the model host of WFT. In Australia, strawberry is an intensively managed crop cultivated for its fresh, aromatic, red berries, with a gross value of approximately AUD$308 million per year (Anonymous 2009a). WFT is considered a major pest of low tunnel- and greenhouse-grown strawberries and production is often affected by its direct damage (Ullio 2002). Studies have shown that flowers may provide WFT with essential resources, either by serving as a mating site (Rosenheim et al. 1990), or as a source of high-quality food (Trichilo and Leigh 1988). WFT feeding on fruit typically causes direct puncture damage (Tommasini and Maini 1995). Medhurst and Steiner (2001) suggested that WFT damage contributes to „seediness‟ of fruit (Figure 1.3), and that this may be responsible for uneven ripening and yield loss, both of which reduce grower profits (Houlding and Woods 1995). Feeding by WFT on blossoms may also cause stigmas and anthers to turn brown and wither prematurely (Zalom et al. 2001). Feeding by WFT causes significant reduction in flower receptacle size in strawberry (Coll et al. 2007). However, strawberry is not affected by TSWV (Herron et al. 2007).

1.5 Chemical control of western flower thrips and insecticide resistance

Chemical control has become standard practice for WFT management (Contreras et al. 2001). Considerable laboratory and field studies have focused on the toxicity of insecticides to WFT (Table 1.2). In Australia, WFT control currently relies on „older chemistry‟ insecticides including organophosphates (acephate, chlorpyrifos, dichlorvos, dimethoate, endosulfan,

8 Chapter I: General introduction and literature review malathion, methamidophos, methidathion, pyrazophos), carbamates (methiocarb, methomyl) and some newer chemistry insecticides (fipronil, abamectin, spinosad). These are made available to growers under permits issued by the Agricultural Pesticides and Veterinary Medicines Authority (APVMA) (Herron and James 2005), or through product registration.

Table 1.2 Insecticides that have been tested for efficacy against WFT.

Insecticide IPM Class Reference compatibility* Endosulfan Organophosphate 1, 5, 7, 12, 16 Chlorpyrifos Organophosphate 1, 4, 7, 10, 12, 13, 15 Diclorvos Organophosphate 4, 5, 6, 7, 8, 18 Etrimfos Organophosphate 3 Malathion Organophosphate 8, 12 Methamidophos Organophosphate 1, 2, 4, 5, 7, 14, 16

Monocrotophos Organophosphate 1, 13 Omethoate Organophosphate 2, 7, 14 Acephate Organophosphate 20 Naled Organophosphate 20 Sulfotep Organophosphate 20 Formetanate Carbamate 4, 7, 9, 10, 14 Furathiocarb Carbamate 3 Methiocarb Carbamate 4, 10, 11, 12, 13, 14, 17 Methomyl Carbamate 1, 5, 7, 19 Bendiocarb Carbamate 20 Fenoxycarb Carbamate 20 Deltamethrin Pyrethroid 2, 8

Λ-cyhalothrin Pyrethroid 5, 10, 14 Bifenthrin Pyrethroid 20 Cyfluthrin Pyrethroid 20 Fluvalinate Pyrethroid 20 Fenpropathrin Pyrethroid 20 Resemethrin Pyrethroid 20 Abamectin* Yes Macrocyclic lactone 5, 13, 14, 15 Azadiractin (neem) Yes Botanical 20 Nicotine* Yes Botanical 20 Beauveria bassiana* Yes Microbial 20 Spinosad* Yes Spinosyn 20 [1.Hamrick (1987), 2. Bohmer and Eilenbach (1987), 3. Freuler and Benz (1988), 4. Paiter (1990), 5. Bournier (1990), 6. Ramakers (1990), 7. Heungens and Butaye (1990), Heungens et al. (1989), 8. Ribes (1990), 9. Jover et al. (1990), 10. Ferrer et al. (1990), 11. Puiggros et al. (1990), 12. Devesa and Iberica (1990), 13. Gokkes (1991), 14. Grasselly et al. (1991), 15. Nasruddin and Smitley (1991), 16. Bohmer et al. (1992), 17. Pasini et al. (1993), 18. Staay and Uffelen (1988), 19. Heungens (1994)] cited in Tommasini (2003), 20. McDonough et al. (2009)].*IPM compatibility (Biobest 2009)

Although chemical control has been the primary tool for WFT management (Contreras et al. 2001), because of the small size, secretive habit, high reproductive potential of WFT, haplodiploidy and improper use of insecticides, WFT has developed insecticide resistance to several major classes of insecticides (Broadbent and Pree 1997, Jensen 1998, Jensen 2000c,

9 Chapter I: General introduction and literature review

Bielza et al. 2007b, Bielza et al. 2008). It has developed resistance to one or more insecticides in different part of the world (Helyer and Brobyn 1992, Brødsgaard 1994) including Australia (Herron and Gullick 1998, Herron and James 2005). In 1961, the first control failure was reported in Mexico to control WFT on cotton with toxaphene (organochlorine) [Race (1961) cited in Jensen (2000b)]. However, it was not until 30 years after the first report that Robb (1989) carried out an experiment to demonstrate insecticide resistance in WFT. Since then, there have been numerous reports published on the reduction in efficacy of insecticides to control WFT, indicating the presence of insecticide resistance [for example (Brødsgaard 1991, 1994, Robb et al. 1995, Zhao et al. 1995c, Zhao et al. 1995b)]. Strains of WFT have developed resistance to organochlorines, organophosphates, carbamates and pyrethroids and some newer reduced-risk insecticides (Schreiber et al. 1990, Immaraju et al. 1992, Brødsgaard 1994, Robb et al. 1995, Broadbent and Pree 1997, Herron and Gullick 1998, Jensen 1998, Jensen 2000c, Herron and Gullick 2001, Espinosa et al. 2002a, b, Espinosa et al. 2005, Herron and James 2005, Loughner et al. 2005, Ralf Nauen 2005, Maymo et al. 2006, Bielza et al. 2007b, Bielza et al. 2007a, Bielza et al. 2008, Zhang et al. 2008). Immaraju et al. (1992) reported high levels of resistance to pyrethroids (permethrin, bifenthrin, abamectin) and moderate-to-high levels of resistance to methomyl in some populations of WFT. In Europe and Africa, WFT populations have shown resistance to acephate and endosulfan (organophosphates), and methiocarb (carbamate) (Brødsgaard 1994). North American WFT populations showed resistance to diazinon and methomyl (organophosphates), bendiocarb (carbamate) and cypermethrin (pyrethroid) (Zhao et al. 1995a). In Australia, WFT has developed resistance to almost all insecticides currently available under permits issued by the APVMA (Herron and James 2005), except methiocarb and pyrazophos (Herron and James 2005). In Australia, a high level of resistance to pyrethroids (cypermethrin, bifenthin, deltamethrin and fluvalinate) was detected and pyrethroids are no longer recommended for use against WFT (Herron and Gullick 2001). WFT has also been reported to develop resistance to abamectin (Immaraju et al. 1992, Kontsedalov et al. 1998), endosulfan (Brødsgaard 1994), DDT, imidacloprid and amitraz (Zhao et al. 1994, Zhao et al. 1995c, Zhao et al. 1995a, b). Pesticide resistance has become more problematic since cross-resistance has been observed, for example resistance to methiocarb exists in populations of WFT that have never been exposed to methiocarb (Brødsgaard 1994, Jensen 2000a). In addition, some of these insecticides have either direct (e.g. mortality) or indirect (oviposition, longevity and predation) adverse effects on beneficials resulting in pest resurgence (Blümel et al. 1999).

Increasing problems with resistance, the availability of insecticides and high cost of chemicals usage, environmental and health risk and adverse effects on natural enemies have challenged us to develop ways of using insecticides or new insecticide that can be effective against WFT with

10 Chapter I: General introduction and literature review no or minimal effect on natural enemies (Zhang and Sanderson 1990, Jensen 2000c, Shipp et al. 2000b, James 2002, Jones et al. 2005), For example, Chapman et al. (2009) experimented with the release of Trichogramma osttiniae and the use of bio-rational insecticides (spinosad, indoxacarb and methoxyfenozide). They found that these two tactics provide an environmentally sound approach to managing Ostrinia nubilalis (Hübner) (Lepidoptera: Crambidae) in bell peppers, due, in part, to the conservation of generalist predators. Kraiss and Cullen (2008) evaluated the efficiency of pyrethrin, insecticidal soap and mineral oil (known as reduced-risk insecticides) and the predatory coccinellid, Harmonia axyridis (Pallas) (Coleoptera: Coccinellidae), for the management of soybean aphid, Aphis glycine Matsumura (Hemiptera: Aphididae) on North American soybean (Glycine max (L.)). They found that these chemicals could be used for the management of A. glycine in the presence of H. axyridis.

The novel insecticide spinosad (Dow AgroSciences USA) is a macro-cyclic lactone bio- insecticide (Elzen et al. 1998b, a, Ludwig and Oetting 2001). Due to its effective use rate, safety to the environment, mammals and beneficials, the Environmental Protection Agency (USA) classified spinosad as an environmentally and toxicologically reduced-risk chemical (Saunders and Bret 1997). Spinosad is efficacious against several thrips species (Cloyd and Sadof 2001) including WFT (Eger Jr. et al. 1998, Funderburk et al. 2000), and marketing has focused on its favourable environmental profile, emphasising its potential for use in IPM systems (Thompson and Hutchins 1999, Thompson et al. 2000). Because of its effectiveness and safety, spinosad was also awarded the „Presidential Green Chemistry Challenge Award” in 1999 (Anonymous 2009b). Copping (2001) in his “Biopesticide manual” reported that spinosad shows no effects on predatory insects such as ladybirds, lacewings, big-eyed bugs or minute pirate bugs. According to the International Organisation for Biological Control (IOBC), the recommended rate of spinosad is harmless to most predatory arthropods and has been reported to be compatible with predatory mites, which are extensively used biocontrol agents against WFT and spider mites (Bret et al. 1997, Williams 2001, Miles et al. 2003, Williams et al. 2003, Anh et al. 2004, Kim et al. 2005, van Driesche et al. 2006a). Similarly, Bret et al. (1997) reported that spinosad is non-toxic to natural enemies including Orius spp., Chrysopa spp., coccinellids, and predaceous mites. Holt et al. (2006) tested the compatibility of spinosad and the predatory mite Phytoseiulus persimilis Athias-Henriot (Phytoseiidae) for the control of two- spotted spider mites on ivy geraniums and found that it can be used in conjunction with spinosad, causing no obvious detrimental effects to this predator. However, Cote et al. (2004) reported that toxicity of spinosad against natural enemies as variable. van Driesche et al. (2006a) report that direct toxicity testing using fresh residues (2h) of the recommended rate of spinosad for WFT on greenhouse flower crops had no toxic effect on the mite Neoseiulus cucumeris (Oudemans), but lowered the survival rate of Iphiseius degenerans (Berlese).

11 Chapter I: General introduction and literature review

Unfortunately, WFT strains have developed resistance to spinosad in some parts of the world (Loughner et al. 2005, Bielza et al. 2007b, Bielza et al. 2007a, Zhang et al. 2008) including Australia (Herron and James 2005). In August 2008, Dow AgroSciences voluntarily suspended the use and sale of multiple spinosyn insecticides from two counties of Florida for 12 months as WFT has developed resistance (Anonymous 2008). Although spinosad resistance in WFT has been detected in some parts of the world, it still appears to be a promising insecticide, particularly since it can be combined with biological control (Jensen 2000c).

1.6 Biological control of western flower thrips

Use of biological control agents for the management of arthropod pests dates back to the 12th century when farmers in the Oriental region used ants to protect fruit trees from pests (Thacker 2002). Natural enemies have been incorporated into IPM strategies of commercial glasshouses for WFT management with varying degrees of success (Chambers and Sites 1989, Gillespie 1989, Gilkeson 1990, Brødsgaard 2004b, Shipp and Ramakers 2004). Several species of natural enemy have been reported to attack WFT (Table 1.3) (van Driesche et al. 1998), including predators (Riudavets 1995, Sabelis and Van Rijn 1997), parasitoids (Loomans et al. 1997), entomopathogens, particularly fungi (Butt and Brownbridge 1997) and nematodes (Loomans et al. 1997).

1.6.1 Thrips predators

Many arthropods are known to be predators of thrips and belong to several families under orders Hemiptera, Diptera, Neuroptera, Coleoptera, Araneida, and some species of Thysanoptera (Lewis 1973, Ananthakrishnan 1979). However, the most widely employed predators are anthocorid bugs of the genus Orius (Anthocoridae) and phytoseiid mites (Acari) (Riudavets 1995, Sabelis and Van Rijn 1997). The genera Orius Wolf, Anthocoris Fallen, Montandoniola Poppius, Xylocoris Dufour and Scoloposcelis Fieber are predators of several thrips species (Ananthakrishnan and Sureshkumar 1985) in a wide range of field crops, tree crops and ornamentals (Herring 1966) in temperate and Mediterranean areas of North America and Europe. Many Orius species can be effective predators of WFT (Sabelis and Van Rijn 1997) and have been used in greenhouse-grown capsicum, cucumber, and eggplant (Gilkeson 1990, van den Meiracker and Ramakers 1991, Chambers et al. 1993, Bolckmans and Tetteroo 2002) in Europe and the USA. Among Orius spp., substantial studies have been made on O. tristicolor (White), O. insidiosus (Say), O. majusculus (Reuter), O. niger (Wolff), O. laevigatus (Fieber) and O. albidipennis (Reuter) (Gilkeson et al. 1990, van den Meiracker and Ramakers 1991, Dissevelt et al. 1995, Rubin et al. 1996, Brown et al. 1999, Funderburk et al. 2000,

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Ramachandran et al. 2001, Deligeorgidis 2002, Ludwig 2002, Shipp and Wang 2003, Tommasini 2003, Brødsgaard and Enkegaard 2005, Chow et al. 2008). Orius spp can be found on several cultivated and non-cultivated plant species and appear to prey on all different thrips stages (Ramakers 1990, Sabelis and Van Rijn 1997). However, the Orius population and its

Table 1.3 Natural enemies evaluated as biocontrol agents of WFT (van Driesche et al. 1998).

Generalist/ Cropping Natural enemies Mass production Specialist system Predators Heteroptera Orius spp. Generalist GH/Field Some species available expensive Anthocoris nemorum Generalist GH Possible, expensive Geocorus spp. Generalist Field Not yet developed Nabis spp. Generalist Field Not yet developed Dicyphus spp. Generalist GH/Field Not yet developed Macrolophus spp. Generalist GH Possible, reasonable Thysanoptera (predatory thrips) Aeolothrips spp. Generalist Field Not yet developed Acari (predatory mites) Some available Amblyseius / Neoseiulus spp. Generalist GH Possible, cheap Hypoaspis spp. Generalist GH/Field Possible, reasonable Typhlodromips montdorensis Generalist GH Available Iphiseius degenerans Generalist GH/Field Available Various predators Neuroptera, Diptera Specialist GH/Field Some available Parasitoids

Ceranisus spp. Generalist GH/Field Difficult Pathogens (nematodes, fungi) Steinernema spp. Possible, cheap Heterohabditis spp. Generalist GH Possible, cheap Thripinema spp. Unknown GH Not yet developed Verticillium lecanii Generalist GH/Field Possible, cheap Beauveria bassiana Generalist GH Possible, cheap Metarhizium anisopliae Generalist GH Not yet developed Paecilomyces fumosoroseus Generalist GH/Field Generalist GH Not yet developed *GH = Glasshouse effectiveness are strongly influenced by prey type, density and environmental factors, especially temperature (Riudavets 1995). Moreover, some species of Orius are injurious to plants since they probe plant tissue for moisture, or oviposit on growing tips (Riudavets 1995). In addition, Orius spp. often leave the crop or glasshouse (Riudavets 1995).

Geocoris pallens (Stal) and G. atricolor Montandon (Anthocoridae) have been described as predators of WFT (Benedict and Cothran 1980, Riudavets 1995). However, Geocoris spp. are omnivorous and frequently consume plant juices (Benedict and Cothran 1980, Yokoyama

13 Chapter I: General introduction and literature review

1980). Adults and larvae of Nabis alternatus (Parshley) and N. americoferus (Carayon) (Nabidae) appear to predate on WFT in alfalfa and bean (Benedict and Cothran 1980). In the Miridae, Deraecoris tamaninii Wagner and Macrolophus caliginosus Wagner are considered to be WFT predators (Riudavets et al. 1993b, a, Riudavets 1995). Some thrips species in the genus Aeolothrips (Thysanoptera), A. fasiatus L. and A. intermedius Bagnall (Lacasa 1980, Baker 1988) cited in Tommasini (2003)] predate on WFT.

1.6.2 Predatory mites

Worldwide, considerable research has focused on the effectiveness of predatory mites (Acari) against insect and mite pests (Chant 1985, Sengonca and Bendiek 1988, McMurtry and Croft 1997, Sabelis and Van Rijn 1997, Brown et al. 1999, Williams 2001, Berndt 2003, Weintraub et al. 2003, Amano et al. 2004, Berndt et al. 2004a, Berndt et al. 2004b, Colfer et al. 2004, Wiethoff et al. 2004, Jones et al. 2005, Cakmak et al. 2006, Ebssa et al. 2006, Holt et al. 2006, Khan and Morse 2006, Kongchuensin and Takafuji 2006, Rhodes and Liburd 2006, van Driesche et al. 2006b, Fitzgerald et al. 2007, Gerson and Weintraub 2007, Ochiai et al. 2007). Biological control of thrips started with observations of predatory mites preying on Thrips tabaci (Lindeman) in greenhouse crops [Woets (1973) cited in Messelink et al. (2006). However, the first deliberate attempt to control a thrips population was with releases of Neoseiulus barkeri (Hughes) (= Amblyseius mckenziei) (Ramakers 1980). However, the introduction of another indigenous (North European) species, Neoseiulus cucumeris (Oudemans), was more successful (de Klerk and Ramakers 1986). Thereafter, several predatory mite species have been used in WFT management programs in field and greenhouse crops (Table 1.4).

Successful control of WFT with predatory mites varies (Chambers and Sites 1989, Gillespie 1989, Gilkeson 1990, Brødsgaard 2004b, Shipp and Ramakers 2004). Several reasons seem to attribute to the varying degree of success. Firstly, unlike Orius or other thrips predators, predatory mites either prey on larval (e.g. N. cucumeris) or pupal (e.g. H. miles) stages. Secondly, the effectiveness of predatory mites is host-plant-dependent. Brown et al. (1999) reported that the efficacy of N. cucumeris and I. degenerans varied on host plants [(plants: Justicia adhatoda, Saurauia nepaulensis DC., Clivia miniata Lindl., Crambe strigosa L‟Her., Greyia radkloferi Szyszyl, Dietes bicolour Steud., Crotalaria capensis Jacq., Tephrosia grandiflora Pers., Peumus boldus Molina, Limonium spectabile (Svent.), Photinia mussia L., Capsicum annuum L., Dombeya acutangula Cav.]. Riudavets (1995) stated that control of WFT by N. cucumeris was lower on cucumber and eggplant compared to capsicum (sweet pepper). Thirdly, some pesticides applied to control other pests are often harmful to predatory mites

14 Chapter I: General introduction and literature review

(Hassan et al. 1987, Hassan et al. 1988, Kim and Paik 1996, Kim and Seo 2001, Mochizuki 2003, Amano et al. 2004). Fourthly, environmental factors such as high or low temperature, humidity and short photoperiods attribute to the effectiveness of predatory mites (Malezieux et al. 1992, Shipp and van Houten 1997, Kiers et al. 2000, Shipp et al. 2000a, Jacobson et al. 2001). Apart from these factors, WFT control by predatory mites might also fail if WFT densities are very high. Therefore, pesticide applications may be necessary to reduce thrips populations and these may be detrimental to predatory mites (Malezieux et al. 1992).

Table 1.4 Crops and predatory mites used for WFT control (Gerson and Weintraub 2007).

Crop Predator Reference Beans Hypoaspis (Geolaelaps) aculeifer Berndt et al.(2004a) Chrysanthemum Neoseiulus cucumeris Oudemans Skirvin et al. (2006) Stratiolaelaps scimitus (Womersley) Bennison et al. (2002a) H. aculeifer Cucumber N. cucumeris Messelink et al. (2005) Amblyseius swirskii Athias-Henriot S. scimitus Wiethoff et al. (2004) H. aculeifer N. cucumeris Cyclamen De Courcy Williams (2001) Typhlodromips montdorensis Schicha Gerbera Steiner and Goodwin (2002) N. cucumeris Impatiens van Driesche et al. (2006b) N. cucumeris Pepper van Houten et al. (2005) A. swirskii

Iphiseius degenerans Berlese A. andersoni Chant Vänninen and Linnamäki N. cucumeris (2002) Rose T. montdorensis Steiner (2002) Strawberry N. cucumeris Tomatoes Shipp and Wang (2003)

1.6.3 Thrips parasitoids

Very little is known about WFT parasitoids. Three chalcid genera Ceranius, Goetheama, Thripobius (Hymentoptera: Chalcidoidea) and a eulophid Entedonastichus (Eulophidae: Entodontinae) are known to parasitise certain thrips species (Loomans 2003). Only two species of Ceranisus: C. menes (Walker) and C. americensis (Girault) are known to attack and develop on WFT (Loomans et al. 1993, Loomans and van Lenteren 1996, Loomans 2003). However, C. menes and C. americensis showed very limited searching efficiency and low parasitisation rates when released against WFT on roses in greenhouses in the Netherlands (Loomans et al. 1995). Entomopathogenic nematodes (EPNs) in the families Steinernematidae and Heterorhabditidae are potential biological control agents of thrips (Chyzik et al. 1996, Ebssa et al. 2001, Ebssa et al. 2004a, Ebssa et al. 2004b, Belay et al. 2005, Ebssa et al. 2006). The genus Thripinema (=

15 Chapter I: General introduction and literature review

Howwardula) (Tylenchida: Allantonematidae) has been reported to parasitise several species of thrips (Sharga 1932, Nickle and Wood 1964, Greene and Parrella 1993, Tipping et al. 1998). T. nicklewoodi (Siddiqi) is the most abundant pupal parasite of WFT used in commercial greenhouses and is able to maintain WFT populations below economic thresholds (Greene and Parrella 1993, Arthurs and Heinz 2002, Mason and Heinz 2002, Smith et al. 2005). Thripinema nicklewoodi is abundant in greenhouse-grown ornamentals and appears to attack WFT in confined areas (Greene and Parrella 1993, Mason and Heinz 2002). Despite its effectiveness, a commercial product of T. nicklewoodi is not presently available (Arthurs et al. 2003). Temperature also has a significant effect on the success of T. nicklewoodi, and it is most successful between 10 and 20⁰C (Arthurs et al. 2003). Another nematode Steinernema feltiae Filipjev (commercially available in USA) has variable success with control of WFT (Chyzik et al. 1996, Arthurs and Heinz 2006).

1.6.4 Fungi

The fungal species, Zoophthora radicans (Brefeld), Neozygites parvispora (Macleod and Carl) (Keller and West 1983); Entomophthora thripidum Samson (Samson et al. 1979, Ananthakrishnan 1993); Beauveria bassiana Bals. (Dyadechko 1964, Lipa 1985, Ananthakrishnan 1993), Verticillium lecanii (Zimm.) (Nedstam 1991), Metarhizium anisopliae (Metchnikoff), Paecilomyces fumosoroseus (Wize) have all been reported as pathogens of WFT. Mycotal, a product containing V. lecanii is now available in Europe for the control of WFT (Parker 2006). Metarhizium anisopliae, B. bassiana, Lecanicillium muscarium, and Trichoderma viride can provide control of WFT in greenhouse vegetables and ornamental crops (Brownbridge 1995, Butt and Brownbridge 1997, Bradley et al. 1998, Shipp et al. 2002, Ugine et al. 2005, 2007, Gouli et al. 2008). However, a significant drawback of using pathogenic fungi is that they are effective only at narrow temperate regimes (Inglis et al. 1997, Inglis et al. 1999).

1.6.5 Biological control of western flower thrips in Australia

Several Orius species have been found to predate on WFT (Cook 2000). Orius armatus Gross appeared to have a significant impact on WFT in the field (Steiner and Goodwin 2000) and was available in late 2009 for commercial use. Phytoseiid mites have shown some potential for controlling WFT in Australia (Steiner and Goodwin 2000). This includes the native Typhlodromips montdorensis (Schicha) (Figure 1.2B) and Typhlodromus occidentalis (Nesbitt) (Phytoseiidae) (Steiner and Goodwin 2000), both of which have been trialled in greenhouses. T. montdorensis occurs on a wide range of plants including Ageratum sp., cucumber, Datura sp., Eucalyptus sp., strawberry, tomato, Mucuna sp., Oxalis sp., purple bean, green bean, Sechium

16 Chapter I: General introduction and literature review edule (Choko) and Sida acuta (Schicha 1979)Steiner et al. (2003). Typhlodromips montdorensis is widely distributed in unforested coastal areas of Queensland and in irrigated and higher rainfall areas including Biloela and the Atherton Tablelands. It has been reported from New South Wales, South Australia and the Northern Territory (Goodwin and Steiner 1996), and is currently used for WFT and spider mite management (Steiner and Goodwin 1998). Another indigenous species Typhlodromus lailae (Schicha), also appears to be promising candidate for development as a commercial biocontrol agent for WFT (Steiner et al. 2003). Worldwide the most widely employed predatory mite, N. cucumeris (Figure 1.2C) was recently confirmed as occurring in Australia (Steiner and Goodwin 2000), and is being reared commercially in South Australia and Western Australia. Hypoaspis (= Stratiolaelaps ) miles (Berlese) (Laelapidae) (Figure 1.2D) has been available on the Australian market for some time. Although H. miles is primarily used for fungus gnat control, it is effective against WFT (L. Chilman, Manchil IPM Service, Pers. Comm.). Little is known about the parasitoids or pathogens present in Australia that would be suitable for the control of WFT (Steiner and Goodwin 2000).

1.7 Host-plant resistance

Host-plant resistance plays an important role in IPM programs [Smith (1990) cited in Parrella and Lewis (1997)] and is considered a key method for pest control on crops with low economic thresholds (Nothnagl 2006). The quality of host plants and their availability plays an important role in the life cycle of phytophagous insects (White 1969, Wellington 1977, White 1993). Plants vary in their suitability as WFT hosts between species as well as within species (i.e. cultivars) and, as a result, larger infestations occur on some plants than on others (Krik 1997). In addition, development of plant resistance to tospoviruses indirectly reduced direct damage caused by WFT (Ullman 1996). Studies indicate that varietal differences play an important role in WFT management (de Jager et al. 1993, van Dijken et al. 1994, de Jager et al. 1995, de Kogel et al. 1997a, de Kogel et al. 1998, Ohta 2002). Variation in varietal resistance to WFT has been described in several crops including cotton (Trichilo and Leigh 1988), Chrysanthemum (de Jager et al. 1993, van Dijken et al. 1994, Ohta 2002), rose (Gaum et al. 1994), pepper (Maris et al. 2003, Maris et al. 2004), tomato (Kumar et al. 1995), cucumber (Mollema et al. 1995, Soria and Mollema 1995), strawberry (Toshio 2004), cabbage (Stoner and Shelton 1986) and groundnut (Robb and Parella 1989). Ananthakrishnan and Gopichandran (1993) reported that leaf plant morphology such as trichome length and width, density of glandular hairs, the thickness of the leaf cuticle and waxes can have a profound effect on plant acceptability. Brown and Simmonds (2006) mentioned that leaf surface undoubtedly plays a role in the selection and preference of host plants by herbivores. Highly pubescent foliage has been observed to lower the infestation levels of thrips on cultivars of cotton (Rummel and Quisenberry 1979). The

17 Chapter I: General introduction and literature review presence of trichomes on leaf surfaces can provide complex mechanisms enabling the plant to evade thrips infestation (Brown and Simmonds 2006). Trichomes can bioactivate secondary products that may be used by the plant as a chemical method of defence against herbivores (Wagner 1991, Bisio et al. 1999, Roda et al. 2003).

1.8 Cultural methods of control of western flower thrips

Several cultural practices may reduce injury by WFT. Mechanical barriers such as mesh, netting, and plastic sheets (mulch) can contribute to WFT control (Broadbent 1969, Cohen and Marco 1979, Harpaz 1982). Yudin et al. (1991) studied the effect of aluminium polyvinyl netting on WFT numbers in a field of lettuce crop. Berlinger et al. (1993) found that commercial woven screens can reduce the entrance of WFT into greenhouses. Sticky traps can also be used for mass trapping of thrips (Murai 1988, Brødsgaard 1989a, 1993a, b). The use of plastic mulches is a standard cultural practice in many parts of the world. Mulch provides several benefits, including improved retention of irrigation water and soil moisture, conservation of soil applied fertilisers, modulation of soil temperatures, and weed suppression (Terry 1997). Ultra violet (UV) reflective silver mulches have been shown to reduce thrips colonization onto tomato (Scott et al. 1989, Greenough et al. 1990, Brown and J. E. Brown 1992, Kring and Schuster 1992) and pepper (Vos et al. 1995, Reitz et al. 2003). Other cultural practices include the push-pull strategy (Bennison et al. 2002b, Cook et al. 2007), trap/companion crops and intercropping (Bennison et al. 2002b, Matsuura et al. 2005, Kasina et al. 2006, Buitenhuis et al. 2007), habitat manipulation (Nicholls et al. 2000, Groves et al. 2001), irrigation and fertiliser manipulation (Schuch et al. 1998, Chau et al. 2005, Davies Jr. et al. 2005, Chau and Heinz 2006). All can significantly affect the population dynamics of WFT and associated diseases. However, while some of these methods have been readily adopted by growers overseas, this is not the case in Australia.

1.9 IPM programs for control of western flower thrips

Increasing problems with resistance, availability and the high cost of chemicals usage, environmental and health risks, adverse effects on natural enemies and the ineffectiveness of other control strategies have challenged growers to adopt new management strategies (Jensen 2000c, James 2002). As an IPM approach, a combination of cultural, biological, and chemical control appears to be effective for the management of WFT in several crops. For example,

Momol et al. (2004) demonstrated that the combined use of spinosad, actigard and high UV- reflective aluminium mulch effectively reduced TSWV transmitted by WFT as much as 76%, compared to untreated black plastic mulch. Since 1997, Florida pepper growers have been using

18 Chapter I: General introduction and literature review reduced-risk insecticides, O. insidiosus and UV-reflective mulch for the management of WFT (Funderburk et al. 2000, Ramachandran et al. 2001, Reitz et al. 2003). Laboratory and greenhouse experiments of Thoeming and Poehling (2006) illustrated that an integrated approach of soil application of azadirachtin (botanical, neem product) integrated with predatory mites, A. cucumeris and H. aculeifer, is consistently effective against WFT on bean (Phaseolus vulgaris L.) and resulted in efficacies up to 99% in controlling WFT. Soil drenching of neem formulation systemically affects the plant sucking life stages, directly affects the soil-dwelling stages of WFT, while N. cucumeris predate on the larval stage and H. aucleifer predate on the prepupal and pupal stage.

Where multiple biological control agents may be used, it is important to not only evaluate the impact of insecticides on them, but also how they interact with each other. Studies have shown that the timely application of predatory mites and localised application of anthocorid bugs have been very effective against WFT in glasshouse-grown sweet pepper and cucumber (Jacobson 1997, Shipp and Wang 2003, Skirvin et al. 2006). In greenhouse-grown chrysanthemum, a combination of A. cucumeris, Orius sp and V. lecanii has proven to be effective against WFT (Jacobson 1997). Research has shown that a formulation based on B. bassiana, M. anisopliae and L. muscarium can reduce WFT significantly in greenhouse-grown vegetables and floral crops (Brownbridge 1995, Butt and Brownbridge 1997, Bradley et al. 1998, Shipp et al. 2002, Ugine et al. 2005, 2007, Gouli et al. 2008). Premachandra et al. (2003) evaluated the combined application of biocontrol agents and found that the combined use of entomopathogenic nematodes, Heterorhabditis bacteriophora Poinar (Rhabditida: Heterorhabditidae), Steinernema feltiae Filipez (Rhabditida: Steinernematidae) and predatory mite, H. aucleifer can be effective for WFT management.

The use of multiple natural enemies species in pest management programs has been a controversial issue for a long time because of interspecific competition (DeBach and Rosen 1991, Bellows and Hassell 1999). As an integrated approach, there is a growing trend to use two or more species of natural enemies to suppress insect pest populations (Premachandra et al. 2003, Avilla et al. 2004, Blümel 2004, Brødsgaard 2004a, Chau and Heinz 2004, Chow and Heinz 2004, Hoddle 2004, Shipp and Ramakers 2004, Thoeming and Poehling 2006, Chow et al. 2008). Anthocorid bugs of the genus Orius and phytoseiid mites of the genus Amblyseius are commonly used for the control of WFT in greenhouse-grown crops in Europe and North America (Brødsgaard 2004a, Shipp and Ramakers 2004), though resulting benefits for pest suppression were not quantitatively validated (Blockmans and Tetteroo 2002, Skirvin et al. 2006). Some studies support the premise of natural enemy compatibility (Gillespie and Quiring 1992, Wittmann and Leather 1997, Brødsgaard and Enkegaard 2005), whilst others are opposed

19 Chapter I: General introduction and literature review

(Magalhăes et al. 2004, Sanderson et al. 2005). Schausberger and Walzer (2001) demonstrated that interspecific competition may occur when different species of predatory mites are combined together and prey specificity affects the quality and intensity of predator-predator interactions. Others such as Schausberger and Walzer (2001), report that in perennial greenhouse-grown crops, releases of P. persimilis and N. californicus may have complementary effects, and could enhance the biological control of arthropod pests.

In Australia, insecticide applications and the augmentative release of predatory mites are used for WFT management in horticultural crops. However, this approach is not always effective in managing WFT. Moreover, insecticide resistance to WFT is a growing problem in Australia (Herron and James 2005). In order to effectively manage WFT and slow down (if not eliminate) resistance, there is a need to explore strategies that involve the harmonious integration of chemical, biological and other control strategies.

1.10 Outline of this study

The aim of this project was to develop an integrated pest management program for the control of WFT using a range of tactics including resistant varieties, biological control and insecticide. Strawberry [Fragaria ananassa L. (Rosaceae)] was used as the model host of WFT. In Australia, strawberry is an intensively managed crop cultivated for its fresh, aromatic, red berries, with a gross value of approximately AUD$308 million (Anonymous 2009a). WFT is a major pest of low tunnel-, greenhouse- and field-grown strawberries and production is often affected by direct feeding damage (Ullio 2002). Studies have shown that flowers may provide WFT with essential resources, either by serving as a mating site (Rosenheim et al. 1990), or as a source of high-quality food (Trichilo and Leigh 1988). Feeding by WFT on blossoms may cause stigmas and anthers to turn brown and wither prematurely (Zalom et al. 2001), or a significant reduction in flower receptacle size (Coll et al. 2007). WFT feeding on fruit typically causes direct puncture damage (Tommasini and Maini 1995). Medhurst and Steiner (2001) suggest that WFT damage contributes to the „seediness‟ of fruit.

The first section of the present study examined the influence of strawberry cultivars on WFT. The literature suggested that the host-plant resistance plays an important part in the effective management of insect pest populations, especially in crops where low damage is required (de Kogel et al. 1998, Schoonhoven et al. 1998). Studies have indicated that varietal differences plays an important role in WFT management (de Jager et al. 1993, van Dijken et al. 1994, de Jager et al. 1995, de Kogel et al. 1997b, de Kogel et al. 1998, Ohta 2002). In Australia, more than 20 strawberry cultivars are grown. The aim was to explore if strawberry cultivars influence

20 Chapter I: General introduction and literature review feeding and oviposition of WFT. In the present study, it was not possible to evaluate all strawberry cultivars for varietal susceptibility to WFT. Instead, three commonly grown cultivars, Camarosa, Camino Real (Short-day), and Albion (day-neutral) were tested. These cultivars were selected because they are grown throughout Australia.

The remainder of the study explores the possibilities of integrating an IPM-compatible insecticide, spinosad, and three predatory mites, Typhlodromips montdorensis (Schicha) (Phytoseiidae), Neoseiulus cucumeris (Oudemans) (Phytoseiidae) and Hypoaspis miles (Berlese) (Laelapidae) [available in the Australian market for WFT control] as well as effective combinations of these predatory mites. Spinosad is a novel pesticide derived from fermentation of actinomycete Saccharopolyspora spinosa Mertz and Yao. It is a bacterial organism first isolated from a Caribbean soil sample (Sparks et al. 1998), is classified as an environmentally and toxicologically reduced-risk chemical (Cleveland et al. 2002, Thompson et al. 2002) and considered as IPM compatible (Thompson and Hutchins 1999, Thompson et al. 2000). Laboratory and field evaluation of the selectivity of spinosad indicates that it is less toxic to natural enemies including predatory mites than to their prey (Bret et al. 1997, Miles and Dutton 2000, Thompson et al. 2000, Medina et al. 2001, Holt et al. 2006, Kim et al. 2006, Arthurs et al. 2007). However, Cote et al. (2004) reported toxicity results of spinosad against natural enemies as variable. For example, Villanueva and Walgenbach (2005) found it is highly toxic to Neoseiulus fallacies (Garman). Moreover, WFT has developed resistance to spinosad in some locations (Herron and James 2005, Bielza et al. 2007b). Typhlodromips montdorensis, N. cucumeris and H. miles appear to be effective against WFT. However, these mites may not be very effective alone. Moreover, T. montdorensis predate on first and second instar and most effective at average temperatures over 20ºC and require high humidity (Steiner and Goodwin 2002, Hatherly et al. 2004). While N. cucumeris predate only the first instar of WFT and their predation rate is often influenced by alternative food like pollen, which hamper its predation efficiency. Hypoaspis miles is a soil-dwelling predatory mite that predate on the prepupal and pupal stages only. Despite resistance to spinosad and spinosad toxicity on some predatory mites, the literature suggested that the integration of spinosad and predatory mites could be a promising management tactic for WFT in horticulture crops. The aim was to elucidate the compatibility of spinosad and T. montdorensis, N. cucumeris and H. miles and measure the effectiveness against WFT, which in turn might slow down the development of insecticide resistance.

This study assessed more effective methods to release biological control agents in conjunction with insecticide applications to improve the control of WFT. The use of different predatory species singly and in combination against WFT was evaluated. Natural enemies currently

21 Chapter I: General introduction and literature review available for the biological control of WFT forage either on upper plant parts (e.g. N. cucumeris) or in the soil (e.g. H. miles) (Berndt et al. 2004a) and it was hypothesised that combinations of predatory mite species would improve the management of WFT. In the course of this study, differential population growth of WFT on different strawberry varieties was observed. Therefore, the application of pesticides and predators was evaluated on different strawberry varieties.

Thus, this thesis has the following specific objectives:

(i) To determine feeding and oviposition preference and performance of WFT on strawberry cultivars. (ii) To evaluate the effectiveness of a combination of cultivars, spinosad and predatory mite for the management of WFT in glasshouse and low tunnel-grown strawberry. (iii) To investigate the effectiveness of the combined application of predatory mite and spinosad for management of spinosad resistant WFT. (iv) To evaluate the toxicity of spinosad against predatory mite.

1.11 Brief organisation and structure of this thesis

In chapter II, the feeding and oviposition preferences of WFT on strawberry cultivars are evaluated using a choice-test method. Oviposition rate, survival, incubation period, larval period and pupal period associated with different strawberry cultivars are also determined.

In chapter III, an integrated management strategy against WFT in greenhouse-grown strawberry is investigated. This approach included the use of different strawberry cultivars, insecticide (spinosad) and predatory mite releases (T. montdorensis, N. cucumeris and H. miles).

In chapter IV, impact of single versus multiple releases of predatory mites against WFT in greenhouse-grown strawberries is determined. This approach includes the use of insecticide (spinosad) and single or multiple releases of predatory mites (T. montdorensis, N. cucumeris and H. miles). In chapter V, a field study that explores the compatibility of the strategy used in glasshouse (Chapter IV) for the management of WFT is investigated.

In chapter VI, the toxicity of spinosad to predatory mites (T. montdorensis, N. cucumeris and H. miles) is tested via a series of bioassays.

22 Chapter I: General introduction and literature review

In chapter VII, the effectiveness of an IPM management protocol that combines the use of spinosad (bioinsecticide) with predatory mite releases (T. montdorensis, N. cucumeris and H. miles) is assessed against a spinosad-resistant WFT population.

Chapter VIII, discusses this body of work and how it can be used to manage WFT in strawberry grown in glasshouses and low tunnels.

1.12 Literature cited

Agronova. 2008. Agrochemicals-Executive Review. Agronova, UK. Allan Woodburn Associates. 2005. Agrochemicals-executive review, pp. 17, First growth in global agrochemical market for a decade. Allan Woodburn Associates, Orpington, Kent, UK. Allen, W. R., and S. E. Gaede. 1963. The relationship of Lygus bugs and thrips to fruit deformity in strawberries. Journal of Economic Entomology 56: 823-825. Amano, H., Y. Ishii, and Y. Kobori. 2004. Pesticide susceptibility of two dominant phytoseiid mites, Neoseiulus californicus and N. womersleyi, in conventional Japanese fruit orchards (Gamasina: Phytoseiidae). Journal of Acarological Society of Japan 13: 65-70. Ananthakrishnan, T. N. 1979. Biosystematics of Thysanoptera. Annual Review of Entomology 24: 159-183. Ananthakrishnan, T. N. 1993. Binomics of Thrips. Annual Review of Entomology 38: 71-92. Ananthakrishnan, T. N., and N. Sureshkumar. 1985. Anthocorids (Anthocoridae: Heteroptera) as efficient biocontrol agents of thrips (Thysanoptera: Thripidae). Current Science 54: 987-990. Ananthakrishnan, T. N., and R. Gopichandran. 1993. Chemical ecology in thrips host plant interactions. International Science Publisher, New York. Anh, K. S., S. Y. Lee, K. Y. Lee, Y. S. Lee, and G. H. Kim. 2004. Selective toxicity of pesticides and control effects of the two-spotted spider mite, Tetranychus urticae by predatory mite and pesticide mixture on rose. Korean Journal of Applied Entomology 43: 71-79. Anonymous. 2008. Dow AgroSciences temporarily suspends use of spinosyn products in two Florida counties. Dow AgroSciences. Anonymous. 2009a. Strawberry industry strategic plan 2009-2013, pp. 28. Strawberries Australia. Anonymous. 2009b. Spinosad Technical bulletin. http://www.jlgarden.com/uploads/handouts/Spinosad.pdf. Arthurs, S., and K. M. Heinz. 2002. In vivo rearing of Thripinema nicklewoodi (Tylenchida: Allantonematidae) and prospects as a biological control agent of Frankliniella occidentalis (Thysanoptera: Thripidae). Journal of Economic Entomology 95: 668-674. Arthurs, S., and K. Heinz. 2006. Evaluation of the nematodes Steinernema feltiae and Thripinema nicklewoodi as biological control agents of western flower thrips Frankliniella occidentalis infesting chrysanthemum, pp. 141-155, Biocontrol Science and Technology. Arthurs, S., K. M. Heinz, S. Thompson, and P. C. Krauter. 2003. Effect of temperature on infection, development and reproduction of the parasitic nematode Thripinema nicklewoodi in Frankliniella occidentalis. Biocontrol 48. Arthurs, S. P., L. A. Lacey, and E. R. Miliczky. 2007. Evaluation of the codling moth granulovirus and spinosad for codling moth control and impact on non-target species in pear orchards. Biological Control 41: 99-109. Avilla, J., R. Albajes, O. Alomar, C. Castane, and R. Gabarra. 2004. Biological control of whiteflies on vegetable crops, pp. 171-184. In K. M. Heinz, R. G. van Driesche and M. P. Parrella [eds.], Biocontrol in protected culture. Ball Publishing, Batavia, Illinois.

23 Chapter I: General introduction and literature review

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Gouli, S., V. Gouli, M. Skinner, B. Parker, J. Marcelino, and M. Shternhis. 2008. Mortality of western flower thrips, Frankliniella occidentalis, under influence of single and mixed fungal inoculations. Journal of Agriculture Technology 4: 37-47. Grasselly, D., T. Y. Caudal, and M. Trateau. 1991. Lutte chimique contre le thrips Frankliniella occidentalis. Essaia de quelques specialites en laboratoire. Phytoma 433: 54-56. Greathead, D. J. 1995. Natural enemies in combination with pesticides for integrated pest management, pp. 183-197. In R. Reuveni [ed.], Novel approaches to integrated pest management. CRC Press, Boca Raton, Florida. Greene, I. D., and M. P. Parrella. 1993. An entomopathogenic nematode, Thripinema nicklewoodi and an endoparasitic wasp, Ceranisus sp. parasitizing Frankliniella occidentalis in California. IOBC/WPRS Bulletin 16: 47-50. Greenough, D. R., L. L. Black, and W. P. Bond. 1990. Aluminium-surfaced mulch: An approach to control of tomato spotted wilt virus in solanaceous crops. Plant Disease 78: 805-808. Greenough, D. R., L. L. Black, R. N. Story, L. D. Newsom, and W. P. Bond. 1985. Occurrence of Frankliniella occidentalis in Louisiana - a possible cause for the increased incidence of tomato spotted wilt virus. Phytopathology 75: 1362. Groves, R. L., J. F. Walgenbach, J. W. Moyer, and G. G. Kennedy. 2001. Overwintering of Frankliniella fusca (Thysanoptera: Thripidae) on Winter Annual Weeds Infected with Tomato spotted wilt virus and Patterns of Virus Movement Between Susceptible Weed Hosts. Phytopathology 91: 891-899. Hamrick, D. 1987. Be warned: there are no chemicals in the wings for thrips control. Grower Talks 10. Harpaz, I. 1982. Nonpesticidal control of vector-borne diseases, pp. 1-22. In K. F. Harris and K. Maramorosch [eds.], Pathogens, vectors and plant disease. Academic Press, New York. Harrewijn, P., W. F. Tjallingii, and C. Mollema. 1996. Electrical penetration by western flower thrips. Entomologia Experimentalis et Applicata 86 79: 345-353. Hassan, S. A., F. Bigler, H. Boggenschutz, E. Boller, J. Brun, P. Chiverton, P. Edwards, F. Mansour, E. Naton, P. A. Oomen, W. P. J. Overmeer, L. Polgar, W. Rieckmann, L. Samsoe-Petersen, A. Staubli, G. Sterk, K. Tavares, J. J. Tuset, G. Viggiani, and A. G. Vivas. 1988. Results of the fourth joint pesticide testing programme carried out by the IOBC/WPRS-working group "Pesticide and Beneficial Organisms". Journal of Applied Entomology 105: 321-329. Hassan, S. A., R. Albert, F. Bigler, P. Blaisinger, H. Boggenschutz, E. Boller, J. Brun, P. Chiverton, P. Edwards, W. D. Englert, P. Huang, C. Inglesfield, E. Naton, P. A. Oomen, W. P. J. Overmeer, W. Rieckmann, L. Samsoe-Petersen, A. Staubli, J. J. Tuset, G. Viggiani, and G. Vanwetswinkel. 1987. Results of the third joint pesticide testing programme by the IOBC/WPRS-working group "Pesticides and Beneficial Organisms. Journal of Applied Entomology 103: 92-107. Hatherly, I. S., J. S. Bale, and M. R. Worland. 2004. Thermal biology of Typhlodromips montdorensis: implications for its introduction as a glasshouse biological control agent in the UK. Entomologia Experimentalis et Applicata 86 111: 97-109. Hayase, T., and H. Fukuda. 1991. Occurrence of the western flower thrips, Frankliniella occidentalis (Pergande), on cyclamen and its identification (In Japanese). Shokubutsu Boeki 45: 59-61. Helyer, N. L., and P. Brobyn. 1992. Chemical control of western flower thrips (Frankliniella occidentalis Pergande). Annals of Applied Biology 121: 219-231. Herring, J. L. 1966. The genus Orius of the western hemisphere (Hemiptera: Anthocoridae). Annals of Entomological Society of America 54: 1093-1109. Herron, G., M. Steiner, B. Gollnow, and S. Goodwin. 2007. Western flower thrips (WFT) insecticide resistance management plan. New South Wales Department of Primary Indutries. Herron, G. A., and G. Gullick. 1998. Insecticide resistance in western flower thrips in Australia, pp. 164-170. In M. P. Zalucki, R. A. I. Drew and G. C. White [eds.], Sixth

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Staay, M. V. D., and J. V. Uffelen. 1988. Chemical control of western flower thrips. Dichlorvos can also be used for cucumber. Weekblad Groenten en Fruit: 40-43. Steiner, M. 2002. Progress towards integrated pest management for thrips (Thysanoptera: Thripidae) in strawberries in Australia. IOBC/WPRS Bulletin 25: 253-256. Steiner, M., and S. Goodwin. 2000. Development and evaluation of usage protocols for newly developed western flower thrips biocontrol agents, pp. 15-25. In D. Cook [ed.], National strategy for the management of western flower thrips and tomato spotted wilt virus. Department of Agriculture Western Australia, South Perth. Steiner, M. Y., and S. Goodwin. 1998. Development and marketing of an IPM package for western flower thrips, Frankliniella occidentalis (Pergande). Phase II: Evaluation and producing the natural enemies. HRDC/HSNA Report. NSW Agriculture, Gosford, Australia. Steiner, M. Y., and S. Goodwin. 2002. Development of a new thrips predator, Typhlodromips montdorensis (Schicha) (Acari: Phytoseiidae) indigenous to Australia. Bulletin of IOBC/WPRS 25: 245-247. Steiner, M. Y., and S. Goodwin. 2005. Management of western flower thrips, Frankliniella occidentalis (Pergande) (Thysanoptera: Thripidae), in hydroponic strawberry crops: using yellow sticky traps to determine action thresholds. Australian Journal of Entomology 44: 288-292. Steiner, M. Y., S. Goodwin, T. M. Wellham, I. M. Barchia, and L. J. Spohr. 2003. Biological studies of the Australian predatory mite Typhlodromips montdorensis (Schicha) (Acari: Phytoseiidae), a potential biocontrol agent for western flower thrips, Frankliniella occidentalis (Pergande) (Thysanoptera: Thripidae). Australian Journal of Entomology 42: 124-130. Stern, V. M., R. F. Smith, R. van den Bosch, and K. S. Hagen. 1959. The integration of chemical and biological control of the spotted alfalfa aphid. Part I. The integrated control concept. Hilgardia 29: 81-101. Stoner, K. A., and A. M. Shelton. 1986. Studies on resistance to Thrips tabaci in four commercial varieties of cabbage, Cruciferae Newsletter 11:101. Tauber, M. J., M. A. Hoy, and D. C. Herzog. 1985. Biological control in agricultural IPM systems: a brief overview of the current statue and future prospects, pp. 3-9. In M. A. Hoy and D. C. Herzog [eds.], Biological control in agricultural IPM systems. Academic Press, Florida. Terry, L. I. 1991. Frankliniella occidentalis (Thysanoptera: Thripidae) Oviposition in Apple Buds: Role of Bloom State, Blossom Phenology, and Population Density. Environmental Entomology 20: 1568-1576. Terry, L. I. 1997. Host selection, communication and reproductive behaviour, pp. 65-118. In T. Lewis [ed.], Thrips as Crop Pests. CABI University Press, Cambridge, UK. Terry, L. I., and G. DeGrandi-Hoffman. 1988. Monitoring western flower thrips (Thysanoptera: Thripidae) in 'Granny Smith' apple blossom clusters. Canadian Entomologist 120: 1003-1016. Thacker, J. R. M. 2002. An introduction to arthropod pest control. Cambridge University Press, Cambridge. Thoeming, G., and H.-M. Poehling. 2006. Integrating soil-applied azadiractin with Amblyseius cucumeris (Acari: Phytoseiidae) and Hypoaspis miles (Acari: Laelapidae) for the management of Frankliniella occidentalis (Thysanoptera: Thripidae). Environmental Entomology 35: 746-756. Thompson, D. G., B. J. Harris, L. J. Lanteigne, T. M. Buscarini, and D. T. Chartrand. 2002. Fate of spinosad in litter and soils of a mixed conifer stand in the Acacian forest region of New Brunswick. Journal of Agricultural and Food Chemistry 50: 790-795. Thompson, G., and S. Hutchins. 1999. Spinosad. Pesticide Outlook 10: 78-81. Thompson, G. D., R. Dutton, and T. C. Sparks. 2000. Spinosad-a case study: an example from natural products discovery programme. Pest Management Science 56: 696-702. Thwaite, W. 1997. Australia's progress in apple IPM. A review of industry pest management in Australian apple orchards. NSW Agriculture.

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Tipping, C., K. B. Nguyen, J. E. Funderburk, and G. C. Smart. 1998. Thripinema fuscum n. sp (Tylenchida: Allantonematidae), a parasite of the tobacco thrips, Frankliniella fusca (Thysanoptera). Journal of Nematology 30: 232-236. Tommasini, M. G. 2003. Evaluation of Orius species for biological control of Frankliniella occidentalis (Pergande) (Thysanoptera: Thripidae), pp. 215. Wageningen University, Wageningen, The Netherland. Tommasini, M. G., and S. Maini. 1995. Frankliniella occidentalis and other thrips harmful to vegetable and ornamental crops in Europe, pp. 1-42. In A. J. M. Loomans, J. C. Van Lenteren, M. G. Tommasini, S. Maini and J. Riudavets [eds.], Biological Control of Thrips, Wageningen Agricultural University, Wageningen, the Netherlands. Toshio, K. 2004. Varietal difference in the susceptibility of strawberry to western flower thrips Frankliniella occidentalis (Pergande). Kyushu Plant Protection Research 50: 55-61. Trichilo, P. J., and T. F. Leigh. 1988. Influence of resource quality on the reproductive fitness of flower thrips (Thysanoptera: Thripidae). Annals of the Entomological Society of America 81: 64-70. Trumble, J. T. 1998. IPM: overcoming conflicts in adoption. Integrated Pest Management Reviews 3: 195-207. Ugine, T. A., S. P. Wraight, and J. P. Sanderson. 2005. Acquisition of lethal doses of Beauveria bassiana conidia by western flower thrips, Frankliniella occidentalis, exposed to foliar spray residues of formulated and unformulated conidia. Journal of Invertebrate Pathology 90: 10-23. Ugine, T. A., S. P. Wraight, and J. P. Sanderson. 2007. Effects of manipulating spray- application parameters on efficacy of the entomopathogenic fungus, Beauveria bassiana against western flower thrips, Frankliniella occidentalis, infesting greenhouse impatiens crops. Biocontrol Science and Technology 17: 193-219. Ullio, L. 2002. Australia's national strategy for the management of western flower thrips (WFT), Frankliniella occidentalis (Pergande), pp. 687-689. In T. Hietaranta, M.-M. Linna, P. Palonen and P. Parikka [eds.], Proceedings of the fourth International Strawberry Symposium. Acta Horticulturae, MTT Agrifood Research, Finland. Ullman, D. E. 1996. Thrips and tospoviruses; advances and future directions, pp. 310-324. In G. Kuo [ed.], Symposium on the tospoviruses and thrips of floral and vegetable crops. Acta Horticulture, Taichung, Taiwan. van de Veire, M. 1987. Explosieve aantasting en moeilijke bestrijding kenmerk California thrips. Vakblad Bloemisterij 11: 25-27. van den Meiracker, R. A. F., and P. M. J. Ramakers. 1991. Biological control of western flower thrips, Frankliniella occidentalis, in sweet pepper with the anthocorid Orius insidiosus. Mededelingen Facuteit Landbouwkundige Rijksuniversiteit Gent 56: 241- 249. van Dijken, F. R., M. T. A. Dik, B. Gebala, J. de Jong, and C. Mollema. 1994. Western flower thrips (Thysanoptera: Thripidae) effect on chrysanthemum cultivars: plants growth and leaf scarring in nonflowering plants. Journal of Economic Entomology 87: 1312-1317. van Driesche, R. G., S. Lyon, and C. Nunn. 2006a. Compatibility of spinosad with predacious mites (Acari: Phytoseiidae) used to control western flower thrips (Thysanoptera: Thripidae) in greenhouse crops. Florida Entomologist 89: 396-401. van Driesche, R. G., S. Lyon, E. J. Stanek III, B. Xu, and C. Nunn. 2006b. Evaluation of efficacy of Neoseiulus cucumeris for control of western flower thrips in spring bedding crops. Biological Control 36: 203-215. van Driesche, R. G., K. M. Heinz, J. C. Van Lenteren, A. J. M. Loomans, R. Wick, T. Smith, P. Lopes, J. P. Sanderson, M. Daughtrey, and M. Brownbridge. 1998. Western flower thrips: a review of its biological control and other methods. Floral Facts, pp. 30. Ed. Univ. of Massachusetts, Amherst, MA. van Houten, Y. M., M. L. Ostlie, H. Hoogerbrugge, and K. Blockmans. 2005. Biological control of western flower thrips on sweet pepper using the predatory mites Amblyseius cucumeris, Iphiseius degenerans, A. andersoni and A. swirskii. IOBC/WPRS Bulletin 28: 283-286.

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Vänninen, I., and M. Linnamäki. 2002. Performance of Neoseiulus cucumeris as a biocontrol agent of the western flower thrips in cut roses. OILB/WPRS Bulletin 25: 289-292. Villanueva, R. T., and J. F. Walgenbach. 2005. Development, oviposition, and mortality of Neoseiulus fallacis (Acari: Phytoseiidae) in response to reduced-risk Iisecticides. Journal of Economic Entomology 98: 2114-2120. Vos, J. G. M., T. S. Uhan, and R. Sutarya. 1995. Integrated crop management of hot pepper (Capsicum spp) under tropical lowland condition: Effects of rice straw & plastic mulches on crop health. Journal of Crop Protection 14: 445-452. Wagner, G. J. 1991. Secreting glandular trichomes: more than just hairs. Plant Physiology 96: 675-679. Weintraub, P. G., S. Kleitman, R. Mori, N. Shapira, and E. Palevsky. 2003. Control of the broad mite (Polyphagotarsonemus latus (Banks)) on organic greenhouse sweet peppers (Capsicum annuum L.) with the predatory mite, Neoseiulus cucumeris (Oudemans). Biological Control 27: 300-309. Wellington, W. G. 1977. Returning the insect to insect ecology: some consequences for pest management. Environmental Entomology 6: 1-8. Welter, S. C., J. A. Rosenheim, M. W. Johnson, R. F. L. Mau, and L. R. Gusukuma- Minuto. 1990. Effects of Thrips palmi and western flower thrips (Thysanoptera: Thripidae) on the yield, growth and carbon allocation pattern in cucumbers. Journal of Economic Entomology 83: 2092-2101. White, T. C. R. 1969. An index to measure weather-induced stress of trees associated with outbreaks of psyllids in Australia. Ecology 50: 905-909. White, T. C. R. 1993. The inadequate environment: nitrogen and the abundance of animals. Springer Verlag, New York. Whitfield, A. E., D. E. Ullman, and T. L. German. 2005. Tospovirus-Thrips interactions. Annual Review of Phytopathology 43: 459-489. Wiethoff, J., H.-M. Poehling, and R. Meyhöfer. 2004. Combining plant- and soil-dwelling predatory mites to optimise biological control of thrips. Experimental and Applied Acarology 34: 239-261. Williams, M. E. D. C. 2001. Biological control of thrips on ornamental crops: Interactions between the predatory mite Neoseiulus cucumeris (Acari: Phytoseiidae) and western flower thrips, Frankliniella occidentalis (Thysanoptera: Thripidae) on Cyclamen. Biocontrol Science and Technology 11: 41-55. Williams, T., J. Valle, and E. Vinuela. 2003. Is the naturally derived insecticide Spinosad compatible with insect natural enemies? Biocontrol Science and Technology 13: 459- 475. Wittmann, E. J., and S. R. Leather. 1997. Compatibility of Orius laevigatus Fieber (Hemiptera: Anthocoridae) with Neoseiulus (Amblyseius) cucumeris Oudemans (Acari: Phytoseiidae) and Iphiseius (Amblyseius) degenerans Berlese (Acari: Phytoseiidae) in the biocontrol of Frankliniella occidentalis Pergande (Thysanoptera: Thripidae). Experimental and Applied Acarology 21: 523-538. Woets, J. 1973. Integrated control in vegetables under glass in the Netherlands. I.B.O.C./W.P.R.S. Bulletin 1973: 26-31. Yokoyama, V. Y. 1980. Method for rearing G. pallens, a predator in California cotton. Canadian Entomologist 112: 1-3. Yudin, L. S., B. E. Tabashnik, W. C. Mitchell, and J. J. Cho. 1991. Effects of mechanical barriers on distribution of thrips in lettuce. Journal of Economic Entomology 84: 136- 139. Zalom, F. G., P. A. Phillips, N. C. Toscano, and M. Bolda. 2001. Strawberry, Western flower thrips. UC IPM Pest Management Guidelines: Strawberry. UC ANR Publication 3468, Insects and Mites. Davis, California, USA. http://www.ipm.ucdavis.r734301211.html. Zhang, S.-Y., S. Kono, T. Murai, and T. Miyata. 2008. Mechanisms of resistance to spinosad in the western flower thrips, Frankliniella occidentalis (Pergande) (Thysanoptera: Thripidae). Insect Science 15.

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Zhang, Z. Q., and J. P. Sanderson. 1990. Relative toxicity of Abamectin to the predatory mite Phytoseiulus persimilis (Acari: Phytoseiidae) and two spotted spider mite (Acari: Tetranychidae). Journal of Economic Entomology 83: 1783-1790. Zhao, G. Y., W. Liu, and C. O. Knowles. 1994. Mechanisms associated with diazinon resistance in western flower thrips. Pesticide Biochemistry and Physiology 49: 13-23. Zhao, G. Y., W. Liu, and C. O. Knowles. 1995a. Mechanisms conferring resistance of western flower thrips to bendiocarb. Pesticide Science 44: 293-297. Zhao, G. Y., W. Liu, and C. O. Knowles. 1995b. Fenvalerate resistance mechanisms in western flower thrips (Thysanoptera: Thripidae). Journal of Economic Entomology 88: 531-535. Zhao, G. Y., W. Liu, J. M. Brown, and C. O. Knowles. 1995c. Insecticide resistance in field and laboratory strains of western flower thrips (Thysanoptera: Thripidae). Journal of Economic Entomology 88: 1164-1170. Zur Strassen, R. 1986. Frankliniella occidentalis (Pergande 1895), ein nordamerikanischer Fransenflügler (Thysanoptera) als neuer Bewohner europächshäuser. Nachrichtenblat Deutscher Pflanzenschutzdienst 38: 86-88.

42 CHAPTER II

Variation in preference and performance of western flower thrips, Frankliniella Occidentalis (Pergande) (Thysanoptera: Thripidae) on three strawberry [Fragaria x ananassa Duchesne (Rosaceae)] cultivars

Keywords: Frankliniella occidentalis, feeding, oviposition, preference, performance, developmental period, survival rate, strawberry

Abstract

Western flower thrips, Frankliniella occidentalis (Pergande) (Thysanoptera: Thripidae) is considered a major pest of strawberry, Fragaria x ananassa Duchesne (Rosaceae), causing substantial yield loss through direct feeding on flowers and fruit. Frankliniella occidentalis damage can be influenced by host-plant resistance or tolerance. This study determined whether three commercial strawberry cultivars (Albion, Camarosa, and Camino Real) differed in their suitability to F. occidentalis. Determination of resistance of strawberry cultivars to F. occidentalis was based on the level of olfactory response, feeding damage, ovipositional preference and host suitability for reproduction. Frankliniella occidentalis adults preferred to feed on Camarosa compared to Albion and Camino Real. However, if leaves had previously been fed on by conspecifics, there was no difference in feeding preference. Camarosa was the most preferred cultivar for oviposition. More eggs were laid by WFT on Camarosa than either on Albion or Camino Real. More larvae hatched and adults emerged from Camarosa than either Albion or Camino Real. The percentage of unhatched eggs, larvae and pupae that died was highest on Camino Real. The survival rate was highest and lowest on Camarosa and Camino Real respectively. The egg incubation, prepupation, pupation and total developmental periods (egg to adult) was shorter on Camarosa than on Albion or Camino Real. The larval period was longer on Camarosa (4.43 ± 0.12 days) than Albion (4.26 ± 0.16 days) and Camino Real (4.06 ± 0.14 days). Overall, the data indicate that Camarosa was most favourable for WFT population growth. The other cultivars may be a better choice for WFT management.

2.1 Introduction

Western flower thrips (WFT), Frankliniella occidentalis (Pergande) (Thysanoptera: Thripidae), is a serious pest of agricultural and horticultural crops worldwide. Although, it was first discovered in 1895 in California, USA (Pergande 1895), during 1970s, WFT has spread worldwide, causing substantial damage to a wide range of crops . (Lewis 1973, Shipp et al.

Chapter II: Preference and performance

1991, Robb and Parrella 1995, Tommasini and Maini 1995, Lewis 1997a, b, 1998, Kindt et al. 2003, Kirk and Terry 2003). In Australia, WFT was first recorded in 1993 on chrysanthemum in a glasshouse in Yangebup, Western Australia (Malipatil et al. 1993). Since then it has spread to all states except the Northern Territory (Medhurst and Swanson 1999). In Australia, WFT is considered an economic pest of ornamentals, vegetables and fruits (Persley et al. 2009).

Strawberry, Fragaria ananassa Duchesne (Rosaceae) is an intensively managed crop cultivated for its fresh, aromatic, red berries. In Australia, the strawberry industry has grown steadily over the last few years, with a gross value of approximately AUD$308 million per annum (Anonymous 2009). However, strawberry production is affected by direct damage caused by WFT (Cook 2000, Ullio 2002, Steiner and Goodwin 2005) and WFT is considered a major pest in greenhouse-, low tunnel- and open field-grown strawberries (Coll et al. 2007). WFT feeding on strawberry flowers and fruits causes fruit deformation, uneven ripening, early withering of stigmas and anthers, and reduction in flower receptacle size leading to yield loss (Buxton and Easterbrook 1988, Houlding and Woods 1995, Medhurst and Steiner 2001). Chemical control of WFT is considered the principal control strategy (Broughton and Herron 2009). However, efficient chemical control of WFT is difficult because of their cryptic feeding behaviour, high mobility, soil-dwelling life stages, and short generation time combined with high fertility (Jensen 2000b). In addition, WFT has developed resistance to all major insecticide classes in different parts of the world (Helyer and Brobyn 1992, Immaraju et al. 1992, Brødsgaard 1994, Zhao et al. 1994, Zhao et al. 1995, Jensen 2000b, a, c, Bielza et al. 2007), including Australia (Herron and Gullick 1998, Herron and James 2005). In Australia, resistance to newer insecticides such as spinosad (Success™), three years after it was first introduced is disturbing, given that it is widely use as an IPM-compatible insecticide (Herron and James 2005). Spinosad resistance has been detected in populations in NSW, Queensland (Herron and James 2005) and Western Australia (S. Broughton, Pers. Comm.) and other parts of the world (Bielza et al. 2007). Alternative approaches for the management of WFT are clearly necessary.

Host-plant resistance is considered a key method for pest control in many crops with low economic thresholds (Schoonhoven et al. 1998). Differences in varietal resistance to thrips have been described in several crops including cotton (Trichilo and Leigh 1988), chrysanthemum (de Jager et al. 1993, Ohta 2002), rose (Gaum et al. 1994), pepper (Maris et al. 2003, Maris et al. 2004), tomato (Kumar et al. 1995a), cucumber (Soria and Mollema 1995), strawberry (Toshio 2004), cabbage (Stoner and Shelton 1986) and groundnut (Robb and Parella 1989). Some cultivars can compensate for thrips feeding and influence oviposition, and therefore thrips development (Kumar et al. 1995a). For example, de Kogel et al. (1998) found that feeding, reproduction and adult survival of WFT were reduced on resistant chrysanthemum cultivars

44 Chapter II: Preference and performance

compared to susceptible cultivars. The development of plant resistance to tospoviruses can also indirectly reduce direct thrips damage (Ullman 1996).

In Australia, growers have access to several strawberry cultivars (e.g. Albion, Camarosa, Camino Real, Ventana, Gaviota, Festival, Kiewa). Evaluation of cultivars tends to be based on yield, flavour, sweetness and shelf-life (Phillips and Reid 2008), rather than varietal susceptibility to WFT. My preliminary observations of WFT on strawberry cultivars suggest that leaf feeding damage varied with cultivar. If strawberry cultivars can influence WFT feeding, oviposition and development, then it could be used by industry as part of an IPM program. Only three cultivars were evaluated [Camarosa (short-day), Camino Real (short-day) and Albion (day-neutral)], since runners of these cultivars were available at the beginning of the project. In this study, I seek to assess the: (i) variation in feeding preference and performance of WFT on strawberry cultivars, (ii) effect of fresh (not previously exposed to conspecifics) and damaged (previously exposed to conspecifics) leaves on preference and performance of WFT, (iii) oviposition preference on strawberry cultivars, and (iv) developmental time and survival.

2.2 Material and methods

Experiments were conducted in a controlled temperature (CT) room (25±1⁰C temperature, 50- 60% RH and 16:8 h L:D regime) at the University of Western Australia (UWA), Crawley. The above conditions were considered most suitable for WFT oviposition (Marullo and Tremblay 1993). At 16.6-36.6ºC the total development period of WFT is 7-13 days (Robb 1989). Experiments were conducted from August to September 2006.

2.2.1 Source cultures

2.2.1.1 Strawberry cultivars

Three commercial strawberry cultivars [Fragaria x ananassa Duchesne (Rosaceae)] (Albion, Camarosa and Camino Real) were used in this study. Strawberry runners were propagated in pots (32.5l x 32.5w x 40.5h cm) containing potting mix (Baileys Fertilisers, Rockingham, WA) in glasshouses at the Department of Agriculture and Food WA (DAFWA) and UWA. All pots were fitted with sprinklers with automatic timers. Plants were watered every third day in summer, and once a week in winter and spring. A liquid fertilizer (Thrive®, Yates, Australia; NPK: 12.4: 3: 6.2; rate: 5mL/2 L water) was applied once a month. All plants were covered with a thrips cage (105µ; 45 x 35 cm)

45 Chapter II: Preference and performance

made from mesh (Sefar Filter Specialists Pty Ltd., Malaga, WA) and supported by a steel-rod stand. Prior to experimentation, plants were moved from the glasshouse into a CT room.

2.2.1.2 Western flower thrips (WFT)

WFT were reared on Calendula, Calendula officinalis L. (Asteraceae), planted in plastic pots (50 x 100 mm) with potting mix and kept in insect-proof Perspex cages (500 x 420 x 400 mm H x D x W), with 105 µ mesh net fitted on a Nylex tote box (432 x 320 x 127 mm; Blyth Enterprises Pty Ltd, Australia). Plants were kept in a glasshouse or net house at UWA from July 2006 to November 2008. All pots were fitted with sprinklers with automatic timers. The plants were watered every third day in summer and once a week during winter and spring. WFT adults were released at the base of each caged plant. Every second week, adults were collected from the caged plants and released onto new potted plants to ensure continuous availability. To obtain uniformly aged WFT, adults (20 individuals) were collected and released onto fresh plants and allowed to lay eggs for 24 h. After 24 h, all adults were removed with an aspirator. Plants were checked daily for adult emergence and adults that emerged on the same day were used in trials.

2.2.2 Experiment 1: Feeding preference of adult WFT

Twenty-four hours prior to the experiment, potted strawberry plants (six plants of each cultivar, total n = 18) were brought to the CT room. Flowers were removed from the plants with a sharp blade, because WFT shows a preference for flowers (Bailey 1933). Plants of each cultivar were divided into two groups, and covered with thrips cages as described above. One group of plants was kept free from WFT (ungrazed), whilst the other group received 50 WFT adults per plant and allowed to feed for 3-4 hours (grazed). To promote feeding, WFT adults were kept without food 24 h prior to release. WFT adults were then removed from the plants with an aspirator.

From the above plants, leaf discs were cut from the leaves for use in the experiment. To prepare leaf discs for trials, leaves were detached with a scalpel. Leaf discs (2.5 cm diam.) were cut using a cork borer (1450-9, MET-APP Pty Ltd, Victoria, Australia) and placed onto moistened filter paper in a Petri dish (150 x 15 mm), axial side down. Leaf discs were placed randomly in a circle by maintaining the same distance between discs. Leaf discs were marked with cultivar code (AL, CA and CR for Albion, Camarosa and Camino Real respectively), and additionally coded ‘F’ (ungrazed leaves) or ‘PI’ (grazed leaves). A combination of ungrazed and grazed leaf discs were placed at the bottom of the Petri dish equidistant apart. Combinations included: (i)

ALF + CRF + CAF, (ii) ALPI + CRPI + CAPI, (iii) ALF + ALPI, (iv) CRF + CRPI and (v) CAF +

46 Chapter II: Preference and performance

CAPI. Each combination was replicated 20 times. WFT adults (3 d old) were collected with an aspirator and kept in Petri dishes (150 x 15 mm) for 24 h to acclimatise them with the experimental arena. No food was provided; except cotton ball soaked in 10% sugar solution was placed inside the Petri dish.

After 24 h, individual thrips were transferred to the centre of an experimental Petri dish. The top of the Petri dish was then covered with mesh (mesh net, 105 µ), and sealed with parafilm (Parafilm M®, Micro Analytix Pty Ltd). Thrips were observed constantly for 30 mins under a stereomicroscope (20-x magnification illuminated by 12v/10w bulbs) and their behaviour recorded. Behavioural events recorded were: (1) olfactory preference, (2) total time spent on a cultivar, and (3) time spent feeding on a cultivar. The first movement of an individual onto a leaf disc after release was considered to be an olfactory preference. Feeding preferences were recorded by measuring times spent feeding (time between probing sylets into the leaf tissue til withdrawn) on the leaf disc of each cultivar. Of the 30 minutes observation, time spent feeding as well as total time spent on each leaf disc was recorded. If an individual thrips did not move from its initial position within two minutes after release, it was discarded and replaced with a new individual. For each individual, a new Petri dish and leaf disc was used.

2.2.3 Experiment 2: Oviposition preference and performance of WFT on caged plants

The experiment was conducted in a CT room as previously described. Three strawberry plants, one of each cultivar (4-5 leaves/plant) that had not been previously exposed to WFT, were placed in a circle on a laboratory bench such that each pot touched. Three-day-old adult WFT females were collected from the colony and placed in a refrigerator for approximately two minutes. Cold anaesthetised WFT adults were then transferred to a filter paper in a Petri dish. For each group of three plants, 20 WFT adults were placed in the centre of the plants and thrips were allowed to disperse. The plants were then covered with a modified cylindrical thrips cage (open both ends), ensuring that the cage did not touch either pot or plant parts which was supported by a cylindrical stand. The cage was 150w x 45h cm in dimension made from mesh net (105 μ). The bottom of the cage was taped onto the bench with masking tape. The top end of the cage was folded and closed with a paper clip for easy handling. Plants were checked at one, 24 and 48 h post-release and adult numbers were recorded on each plant. If previously released WFT were not found, a hand-held battery-powered magnifying glass (4x bifocal magnifier) was used. After 48 h, all adults were removed from the plants with an aspirator. Plants were then checked twice a day (0800 and 1900 h) for one week with a magnifying glass for larvae. No newly hatched larvae were found after five days. Each day, newly hatched larvae were removed with a fine brush, counted and transferred to new plants of the same cultivar.

47 Chapter II: Preference and performance

Individual plants with WFT larvae were covered with a mesh net cage (45 x 35 cm) as described above and . These plants were checked twice daily (0800 and 1900 h) for three weeks, and any emerged adults were counted and removed from the plant. The experiment was repeated 10 times (replicates).

2.2.4 Experiment 3: Oviposition preference and performance of WFT on leaf discs

The experiment was conducted in a CT room. Three leaf discs, one from each cultivar, were prepared from fresh strawberry leaves as described in section 2.2.2. They were placed adaxial side up, two mm apart and equidistant from each other on a moistened filter paper at the centre of a Petri dish (150 x 15 mm). Prior to the experiment, 20 adult females (5-6 d old, females) [on average after three days post-emergence, WFT females start laying eggs at 20-25°C (Marullo and Tremblay 1993)] from the same cohort were collected with an aspirator and kept in Petri dishes for 24 h to acclimatise with the experimental arena. After 24 h, females were released at the centre of the Petri dish and allowed to oviposit for 24 h. The Petri dish was covered with mesh net (105 µ), and the edges sealed with parafilm (15 μm) (de Kogel et al. 1997). After 24 h, WFT adults were removed and each leaf disc was transferred onto moistened filter paper in separate Petri dishes and covered with parafilm as above. Petri dishes were checked under a stereomicroscope (20x magnification) twice daily for one week [at 20-25ºC, eggs hatch within 2-4 days (Pfleger et al. 1995)]. Numbers of hatched larvae were recorded per leaf disc of each cultivar for seven days, though in this trial, no larva hatched after five days. After each count, larvae were removed from the leaf disc and transferred to fresh leaf discs of the respective cultivar. The trial was repeated 20 times (replicates).

2.2.4.1 Egg hatch

After seven days, leaf discs were used to count the numbers of unhatched eggs (if any). Because WFT adults lay eggs inside the leaf tissue by probing their ovipositor, it is not possible to count any unhatched eggs directly. In order to count unhatched inside the leaf tissue, leaf discs of each cultivar were boiled into separate beakers (250 ml) with distilled water into a microwave oven and heated for three minutes (700 watt) (de Kogel et al. 1997). Leaf discs were soaked in methyl red, so that eggs were clearly seen under a stereomicroscope (40 x). Numbers of unhatched in each leaf discs were counted.

2.2.4.2 Larval mortality, pupal mortality and adult emergence rate

To determine the effect of strawberry cultivars on larval mortality, pupal mortality and adult emergence rate, newly hatched larvae (within 12 h of hatching) were collected each day from

48 Chapter II: Preference and performance

leaf discs and transferred to new leaf discs (Ø = 2.5 cm) of the same cultivar. Each disc was placed in a separate Petri dish on slightly moistened filter paper and covered with clear plastic film. Fresh leaf discs were added daily. To reduce handling stress, larvae were allowed to move from the old to the new leaf disc. Once all larvae had moved, the older disk was checked under a magnifying glass, and then discarded. Leaf discs were checked twice daily under a binocular stereomicroscope (20x magnification) until all larvae had either pupated or died. Any dead larvae were removed from the leaf disc. Pupae were checked until adult emergence. If a WFT adult did not emerge, the pupa was considered dead.

2.2.4.3 Developmental time

To determine the influence of strawberry cultivar on egg, larval, prepupal, and pupal period, and total development time (egg to adult emergence), the duration of each period was measured for 40 larvae (replicates) on the three cultivars. Since WFT laid eggs inside leaf tissue, the exact measurement of the incubation period was not possible. However, as WFT adults were allowed to lay eggs for 24 h and leaf discs were checked at 12 h intervals, the incubation period was considered to be the period between half the 24 h period and either the 12 h or 24 h count (depending on when eggs hatched). Leaf discs were changed every second day.

2.2.5 Data analysis

When each thrips chose a leaf disc to move onto, this was scored as the olfactory preference of WFT adults between cultivars. The differences in olfactory preference by WFT adults between cultivars were analysed by a chi-square goodness of fit test. Similarly, olfactory preference between ungrazed and grazed leaf disc within each cultivar was also scored and analysed with a chi-square test. Feeding preference of WFT between cultivars was determined by the amount of time spent feeding on leaf disc of each cultivar. In terms of the amount of time spent feeding, adults exhibited several feeding behaviours such as approaching to leaf disc, stand-still, feeding, resting, walking. However, only the total time spent feeding and the total time that WFT remained on leaf discs was analysed. The time spent by adults among cultivars was expressed as a proportion. To determine adult preference, the total time and the time spent feeding was subjected to Compositional Data Analysis separately for each cultivar combination (Aitchison 1986, Aebischer and Robertson 1993). Least square mean difference with 5% significance level was used to separate means.

In the caged trial, the effect of cultivar on adult numbers per plant at one, 24 and 48 h post release were analysed by separate one-way ANOVAs [independent variable: cultivar, response

49 Chapter II: Preference and performance

variable: WFT adult numbers per plant of each cultivar). The difference in adult numbers between post-release periods for each cultivar was analysed by repeated measures ANOVA. The influence of cultivars on mean numbers of larvae hatched and adults emerged was analysed by two separate one-way ANOVAs. Similarly, the effect of strawberry cultivars on numbers of eggs laid, unhatched eggs, larvae hatched, larvae killed, pupae developed, pupae killed and adults emerged were compared with a series of one-way ANOVAs. If ANOVA results were significant, means were separated by least square means difference (alpha = 0.05).

In the leaf disc trial, the survival of the immature stages and the overall survival rate were calculated as follows: No of larvae hatched Larval hatching rate (%) = 100 No of eggs No of thrips prepupae Pupation rate (%) = 100 No. of thrips larvae tested No. of thrips adults Emergence rate (%) = 100 No. of thrips prepupae No. of thrips adults Survival rates (%) = 100 No of thrips eggs

The influence of strawberry cultivars on larval hatching rate, pupation rate, adult emergence rate and total survival rate (eggs to adult emergence) was analysed with separate one-way ANOVA. Cultivars effect on eggs, larval and pupal mortality was analysed by separate ANOVAs. If ANOVAs were significant, means were separated by least square mean difference (α = 0.05).

Data subjected to ANOVAs were transformed for homogeneity using √(x+0.5) (Zar 1999). Data were reversed transformed for presentation. Statistical analyses were conducted using SAS software 9.1 (SAS 2002-2003).

2.3 Results

2.3.1 Experiment 1: Feeding preference of adult WFT

2 WFT adults showed an olfactory preference for cultivar (χ 2 = 6.34, P = 0.04) leaves not previously exposed to conspecifics. Camarosa was the most preferred cultivar, followed by 2 2 Albion (χ 1 = 3.66, P = 0.03) and Camino Real (χ 1 = 5.76, P = 0.01) (Figure 2.1A). However, there was no significant difference in olfactory preference of WFT adults between Camino Real 2 and Albion (χ 1 = 0.14, P = 0.71) (Figure 2.1A). When offered leaves that had previously been

50 Chapter II: Preference and performance

2 grazed on by conspecifics, adult preference did not differ (χ 2 = 1.03, P = 0.60) among cultivars (Figure 2.1B).

When offered fresh leaf discs, the total time (Wilks λ = 0.47, F2, 17 = 9.54, P = 0.0017) and the time spent feeding (Wilks λ = 0.38, F2, 17 = 13.99, P = 0.0003) differed between cultivars (Figure 2.2). Total time spent and time spent feeding onto each cultivar was longest on Camarosa and shortest on Camino Real (Figure 2.3). The proportion of time spent feeding did not differ between Albion and Camino Real. When offered grazed leaf discs, the total time (λ =

0.51, F2, 17 = 8.28, P = 0.0031) and time spent feeding (Wilks λ = 0.60, F2, 17 = 5.66, P = 0.0131) differed with cultivar (Figure 2.2). In both cases, adults spent the most time and the highest proportion of their time feeding on Camarosa and lowest on Camino Real (Figure 2.3). No differences were detected between Camarosa and Albion.

** A 60 *

40

NS 20

0 60 B NS

40 NS Percentage of WFT (%) of adults Percentage NS

20

0 Camarosa Albion Camino Real

Figure 2.1 Olfactory preference of WFT adults to strawberry cultivars when offered (A) ungrazed and (B) grazed leaf discs. **Significant at P = 0.01, *P = 0.05, NS Not-significant at P = 0.05.

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Figure 2.2 Simplex plot showing time spent by WFT adults between strawberry cultivars when exposed to ungrazed leaf discs (A = total time, B = time spent feeding) and discs previously grazed by conspecifics (C = total time, D = time spent feeding). AL = Albion, CA = Camarosa, CR = Camino Real. In each figure, each dot represents individual WFT adult.

Albion Camarosa Camino Real A 100 c 80 b

SE) 60  40 a 20 a b a 0 100 B 80 b 60 b 40 b Time spent by WFT adult (%) (Mean (%) adult WFT by spent Time a a a 20

0 Total time spent Time spent feeding

Figure 2.3 Time spent by WFT adults on (A) ungrazed and (B) previously grazed leaf discs. Within group, means with different letters differed significantly (LS means, α =0.05).

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WFT showed olfactory preference for ungrazed leaf discs over grazed leaf discs for all three 2 cultivars and difference was significant [Figure 2.4; Albion (χ 1 = 6.24, P = 0.013), Camarosa 2 2 (χ 1 = 9.75, P = 0.0018) and Camino Real (χ 1 = 3.50, P = 0.04)].

When offered a choice between leaf discs from ungrazed and grazed leaves, WFT spent the most time on ungrazed discs compared to grazed discs for all cultivars (Figure 2.5; Albion: λ =

0.09, F1, 8 = 85.43, P < 0.0001; Camarosa: λ = 0.10, F1, 8 = 69.40, P < 0.0001 ; Camino Real: λ

= 0.12, F1, 8= 65.23, P < 0.0001). WFT adults also spent more time feeding on ungrazed leaf discs compared to grazed leaf discs, regardless of cultivar (Figure 2.5).

Ungrazed Grazed

80 * ** * 60

40

20

Percentage of WFT adults (%) adults of WFT Percentage 0 Albion Camarosa Camino Real

Figure 2.4 Preference of WFT adults when given a choice between leaf discs exposed to conspecifics (grazed) or ungrazed of three different strawberry cultivars. **Significant at P = 0.01, *Significant at P = 0.05.

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Ungrazed Grazed 100 A

80

60 SE)

 40

20

0 100 B 80

60 Time spent (%) (Mean Time (%) spent 40

20

0 Albion Camarosa Camino Real

Figure 2.5 Time [A = total time, B = time spent feeding] spent by WFT adult on ungrazed and grazed leaf discs. For each cultivar, means were significantly different (α = 0.05). In each cultivar, WFT adults spent longer times on ungrazed leaf discs compared to grazed leaf discs.

2.3.2 Experiment 2: Oviposition preference and performance of WFT on caged plants

More adults were found on Camarosa and the least on Camino Real when counted at one hour

(F2, 27 = 9.21, P = 0.0009), 24 h (F2, 27 = 23.01, P < 0.0001) and 48 h (F2, 27 = 42.01, P < 0.0001) post release (Figure 2.6). Differences in preferences were not observed between Albion and Camino Real until 24 hours post-release, with more adults on Albion than Camino Real (Figure 2.6).

In each cultivar, WFT adult numbers changed over times from its initial numbers (Figure 2.7). For Albion, WFT adult numbers per plant gradually decreased from initial numbers over time, though the difference (Figure 2.7; F2, 27 = 0.90, P = 0.4201) was not significant. In Camarosa, WFT adult numbers gradually increased from initial numbers at one hour post-release and was highest at 48 h post-release, the difference was significant (Figure 2.7; F2, 27 = 4.89, P = 0.015). WFT adult numbers per plant in Camino Real gradually decreased from its initial numbers; the difference was significant (Figure 2.7; F2, 27 = 3.76, P = 0.0363).

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The mean number of larvae hatched (F2, 27 = 45.83, P < 0.0001) and adults emerged (F2, 27 = 77.91,P < 0.0001) were highest on Camarosa (larvae hatched = 39.80 ± 2.11, adults emerged = 20.27 ± 1.49) and lowest on Camino Real (larvae hatched = 15.00 ± 1.31, adults emerged = 4.40 ± 0.89) (Figure 2.8). Moreover, significantly more larvae hatched and adults emerged on Albion compared to Camino Real (Figure 2.8).

Albion Camarosa Camino Real 18

SE) c  c b 12

a b 6 a b a a

Number of WFT adults (Mean adults WFT of Number 0 1h 24h 48h

Figure 2.6 Mean numbers of WFT adults per plant on caged strawberry cultivars at 1, 24, and 48 h post-release. Means with different letters differed significantly (LS mean, α = 0.05).

1h 24h 48h 20 SE)  b 15 a a

10 a a a 5 c b a

Numbers of WFT adults (Mean adults WFT of Numbers 0 Albion Camarosa Camino Real

Figure 2.7 Comparison of WFT adult numbers (Y-axis) at different post-release periods (hours) on caged plants. Means within cultivar with different letters differed significantly (LS means, α = 0.05).

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50 A c 40

30 b 20 a SE)  10

0 25 c B 20 Number ofNumber WFT (Mean 15

10 b a 5

0 Albion Camarosa Camino Real

Figure 2.8 Mean numbers of WFT (A) larvae hatched and (B) adults emerged per plant on caged strawberry cultivars. Means with different letters differed significantly (LS means, α = 0.05).

2.3.3 Experiment 3: Oviposition preference and performance of WFT on leaf discs

The mean numbers of eggs laid (F2, 57 = 8.74, P = 0.0005), unhatched eggs (F2, 57 = 17.93, P <

0.0001), larvae hatched (F2, 57 = 38.044, P < 0.0001), pupae developed (F2, 57 = 51.89, P <

0.0001) and adults emerged (F2, 57 = 45.03, P < 0.0001) differed significantly between cultivars (Figure 2.9). The greatest numbers of eggs were laid on Camarosa and the lowest on Camino Real. There were no statistically significant differences in egg lay between Albion and Camino Real. The least unhatched eggs were recorded on Camarosa, and more on Camino Real. Numbers of larvae hatched, pupae developed and adults emerged were highest and lowest on Camarosa and Camino Real respectively (Figure 2.9). The number of pupae that developed and adults that emerged did not differ between Albion and Albion (p >0.05).

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Albion Camarosa Camino Real 25 b SE)  20 c a a 15 b b b 10 b a b a 5 a a a a

Numberof WFT (Mean 0 Total eggs Unhatch eggs Larvae Pupae Adults

Figure 2.9 Comparison of numbers of eggs laid, unhatched eggs, larvae hatched, pupae developed and adults emerged per leaf disc (Y-axis) on three strawberry cultivars. Within each group (X-axis), means with different letters differed significantly (LS means, α = 0.05).

The development rate of WFT was different between cultivars (Figure 2.10 and 2.11). Mean percentage of larvae hatched (F2, 57 = 106.70, P < 0.0001), pupae developed (F2, 57 = 7.01, P <

0.0001), adults emerged (F2, 57 = 3.78, P = 0.0284) and survival rate (eggs to adults) (F2, 57 = 24.45, P < 0.0001) was highest on Camarosa and lowest on Camino Real. There were no significant differences between Albion and Camino Real. The mean percentage of unhatched eggs (F2, 57 = 104.48, P < 0.0001), larvae (F2, 57 = 7.04, P = 0.0019) and pupae (F2, 57 = 11.13,P < 0.0001) that died was lowest on Camarosa and highest on Camino Real. There were no differences between Albion and Camino Real.

Albion Camarosa Camino Real 100

SE) b

 c 80 b a a b 60 c a b b a a b b b 40 a

a a 20 Percentage of WFT (%) (Mean (%) of WFT Percentage 0 U/E L/H L/K P P/K A/E

Figure 2.10 Comparison of the percentage of unhatched eggs (U/E), larvae hatched (L/H), larvae died (L/K), pupae developed (P), pupae died (P/K) and adult emerged (A/E) per leaf disc among cultivars. Means within groups with different letters differed significantly (LS means, α = 0.05).

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60

SE) b 

40

a 20 a

Survival rate of WFT (Mean WFT of rate Survival 0 Albion Camarosa Camino

Figure 2.11 Comparison of survival rate of WFT among strawberry cultivars. Means with different letters differed significantly (LS means, α = 0.05).

The mean egg incubation period (F (2, 117) = 4.45, P = 0.0137), larval (F (2, 117) = 3.65, P =

0.0290), prepupal (F (2, 117) = 3.19, P = 0.0447), pupal (F (2, 117) = 14.76, P < 0.0001) and total developmental (F (2, 117) = 4.61, P = 0.0119) periods differed between cultivars (Figure 2.12). Mean egg incubation, prepupation, pupation and total developmental periods were lowest on Camarosa and highest on Camino Real. There were no differences between Albion and Camarosa. Mean larval duration was also highest and lowest on Camarosa and Camino Real respectively, but did not differ between Camarosa and Albion.

Albion Camarosa Camino Real 15

b a a SE) in days in SE)  10

5 ab b a a a b a b b a a b Developmental period (Mean period Developmental 0 IP LP PPP PP TDP

Figure 2.12 Egg incubation period (IP), larval period (LP), prepupation period (PPP), pupation period (PP) and total developmental period (TDP) (egg to adult) of WFT in days, among strawberry cultivars. Means within group with different letters differed significantly (LS means, α = 0.05).

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2.4 Discussion

Host-plant resistance in the present study is defined as a reduction in insect performance. The results confirm that WFT shows a preference for particular strawberry cultivars. When given a choice between three cultivars, Camarosa was the most preferred and Camino Real the least preferred cultivar. This result was confirmed in leaf disc and whole plant choice tests. In leaf disc trials, adults showed an olfactory and feeding preference for Camarosa. Adults spent the most time and the highest proportion of their time feeding on Camarosa and the lowest on Camino Real. In choice tests with whole plants, adults actively dispersed from the less preferred cultivars. Whilst no published information is available on strawberry, Maris et al. (2004) found that WFT adults dispersed from one capsicum cultivar to another cultivar in laboratory trials. This suggests that given a choice, WFT adults can move from one host to another host that may be more suitable for feeding or oviposition. Leaf condition (i.e. leaves grazed by conspecifics vs ungrazed leaves) also strongly influenced adult preference. Adults preferred ungrazed leaves over leaves that had been previously grazed by conspecifics. When offered previously grazed leaves, no cultivar preference was detected. This result agrees with Delphia et al. (2007) who reported that higher numbers of WFT were found on unwounded (fresh) tobacco plants (Nicotiana tabacum L) than plants wounded by conspecific WFT. Delphia et al. (2007) found that wounded tobacco plants released higher amounts of volatile chemicals than unwounded plants. Whether a similar mechanism is operating in strawberry is not known.

The quality of specific cultivars may influence thrips performance (Brødsgaard 1987, Brodbeck et al. 2002). van Lenteren and Noldus (1990) suggest that a higher fecundity rate, faster developmental rate and higher survival rate indicate better host-plant suitability. On resistant cucumber and chrysanthemum cultivars for example, reproduction of WFT is reduced (de Kogel et al. 1997, de Kogel et al. 1998). Soria and Mollema (1995) similarly found that unsuitable cucumber varieties severely reduced the population growth of WFT. To determine whether WFT females chose a strawberry cultivar that offered a possible reproductive advantage, ovipositional preferences were measured in plant (cage) and leaf disc trials. Ovipositional preference was determined by counting the number of larvae that hatched and the number of unhatched eggs. In the cage more larvae developed and adults emerged on Camarosa (larvae hatched = 39.80 ± 2.11, adult emerged = 20.27 ± 1.49) compared to Camino Real (larvae hatched = 15.00 ± 1.31, adult emerged = 4.40 ± 0.89). In leaf disc trials, 50.72% of eggs successfully developed to adult on Camarosa and 15.46% of eggs developed to adult on Camino Real. The egg incubation, prepupation, pupation and total developmental periods (egg to adult) were also determined. If adults fail to emerge, the population of the next generation is affected, which could delay WFT from reaching damaging levels. Whilst all cultivars supported the

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development of eggs to adult, the percentage of adults that emerged and the total development period varied. On Camarosa, 50.72% of eggs developed to adult and the total developmental period was 11.19 ± 0.22 days. In Albion, 21.31% of eggs developed to adult and the total developmental period was 11.20 ± 0.26 days, while in Camino Real, 15.46% eggs successfully developed to adults and the total developmental period was 12.03 ± 0.23 days.

The mechanism or mechanisms that may be involved in the selection by WFT of Camarosa over the two other cultivars is not known, and was beyond the scope of this study. Possible mechanisms include the release of plant volatiles (Bernays and Chapman 1994, Dobson 1994), the presence of toxic metabolites, the absence of, or suboptimal amounts of essential nutrients that are required for insect growth, or the presence of enzymes that inhibit food digestion thus reducing nutrient utilisation (Saxena 1985). For example, Yang et al. (1993) and Snook et al. (1994) report that wild peanut species (Arachis sp) sustained less damage from Frankliniella fusca (Hinds) than cultivated peanut (Arachis hypogea L.). They suggest that this may be due to differences in culticular lipids, especially phenolic acid and acetates (general digestibility inhibitors).

Physical characteristics such as leaf morphology may be also important (Ananthakrishnan and Gopichandran 1993, Kumar et al. 1993, Kumar et al. 1995a, Kumar et al. 1995b). Epidermal appendages such as trichomes can provide a physical barrier against insects. Bioactive secondary products produced by glandular trichomes may also be used as a chemical method of defence (Wagner 1991, Bisio et al. 1999, Roda et al. 2003). Two types of trichomes, simple trichomes and uniseriate glandular trichomes occur in strawberry (Steinite and Levinsh 2003). However the affect of trichomes on WFT on strawberry cultivars is not known. However, Rummel and Quisenberry (1979) reported that infestation of thrips on cotton cultivars varies because of trichome density in different cotton cultivars. The presence of wax on the leaf surface may also play an important role in the selection of hosts by herbivorous insects (Eigenbrode 1996). More than 200 reports on resistance to arthropod pests in vegetables have shown that tolerance was involved in about 10% of the cases, whereas the remaining cases were equally attributed to either antixenosis (inability of a plant to serve as a host) or antibiosis (kill insects or reduce plant digestibility) (Schoonhoven et al. 1998).

In conclusion, feeding and oviposition preference and performance of WFT were affected by strawberry cultivars. Before implementing the present findings in the field, further research should be carried out to determine the seasonal variation in preference and performance on strawberry cultivars. The leaf morphology and nutritional contents of each cultivar should also be assessed to determine the basis for thrips preference, which could then be incorporated into a

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screening program for strawberry cultivars. Furthermore, whether WFT injury could cause economic damage on least preferred cultivar needs to be assessed.

3.5 Literature cited

Aebischer, N. J., and P. A. Robertson. 1993. Compositional analysis of habitat use from animal radio-tracking data. Ecology 74: 1313-1325. Aitchison, J. 1986. The Statistical analysis of compositional data. Chapman & Hall, New York. Ananthakrishnan, T. N., and R. Gopichandran. 1993. Chemical ecology in thrips host plant interactions. International Science Publisher, New York. Anonymous. 2009. Strawberry industry strategic plan 2009-2013, pp. 28. Strawberries Australia. Bailey, S. F. 1933. The biology of the bean thrips. Hilgardia 7: 467-522. Bernays, E. A., and R. F. Chapman. 1994. Host-plant selection by phytophagous insects. Chapman & Hall, New York. Bielza, P., V. Quinto, J. Contreras, M. Torne, A. Martin, and P. J. Espinosa. 2007. Resistance to spinosad in the western flower thrips, Frankliniella occidentalis (Pergande), in greenhouses of southeastern Spain. Pest Management Science 63: 682- 687. Bisio, A., A. Corallo, P. Gastaldo, G. Romussi, G. Ciarallo, N. Fontana, N. Tommasi de, and P. Profumo. 1999. Glandular hairs and secreted materials in Salvia blepharophylla Brandegee ex Epling grown in Italy. Annals of Botany 83: 441-452. Brodbeck, B. V., J. E. Funderburk, J. Stavisky, P. C. Andersen, and J. Hulshof. 2002. Recent advances in the nutritional ecology of Thysanoptera, or the lack thereof, pp. 145-153. In R. Marullo and L. Mound [eds.], Thrips and tospoviruses: Proceedings of the 7th International Symposium on Thysanoptera. Australian National Insect Collection (ANIC), Canberra, Australia. Brødsgaard, H. F. 1987. Frankliniella occidentalis (Thysanoptera: Thripidae)-a new pest in Danish glasshouses. A review. Tidsskr. Planteavl. 93: 83-91. Brødsgaard, H. F. 1994. Insecticide resistance in Europe and African strains of western flower thrips (Thysanoptera: Thripidae) tested in a new residue-on-glass test. Journal of Economic Entomology 87: 1141-1146. Broughton, S., and G. A. Herron. 2009. Management of western flower thrips, Frankliniella occidentalis (Pergande) (Thysanoptera: Thripidae) on strawberries. General and Applied Entomology 38: 37-41. Buxton, J. H., and M. A. Easterbrook. 1988. Thrips as a probable cause of severe fruit distortion in late-season strawberries. Plant Pathology 37: 278-280. Coll, M., S. Shakya, I. Shouster, Y. Nenner, and S. Steinberg. 2007. Decision-making tools for Frankliniella occidentalis management in strawberry: consideration of target markets. Entomologia Experimentalis et Applicata 122: 59-67. Cook, D. 2000. National strategy for the management of western flower thrips and tomato spotted wilt virus, pp. 167. Department of Agriculture and Food Western Australia, South Perth. Western Australia. de Jager, C. M., R. P. T. Butot, T. J. de Jong, P. G. L. Klinkhamer, and E. van der Meijden. 1993. Population growth and survival of western flower thrips Frankliniella occidentalis (Pergande) (Thysanoptera: Thripidae) on different chrysanthemum cultivars. Two methods for measuring resistance. Journal of Applied Entomology 115: 519-525. de Kogel, W. J., M. van der Hoek, and C. Mollema. 1997. Oviposition preference of western flower thrips for cucumber leaves from different positions along the plant stem. Entomologia Experimentalis et Applicata 82: 283-288.

61 Chapter II: Preference and performance

de Kogel, W. J., M. van der Hoek, M. T. A. Dik, F. R. van Dijken, and C. Mollema. 1998. Variation in performance of western flower thrips populations on a susceptible and a partially resistant chrysanthemum cultivar. Euphytica 103: 181-186. Delphia, C. M., M. C. Mescher, and C. M. de Moraes. 2007. Induction of plant volatiles by herbivores with different feeding habits and the effects of induced defences on host- plant selection by thrips. Journal of Chemical Ecology 33: 997-1012. Dobson, H. F. 1994. Floral volatiles in insect biology, pp. 47-81. In E. A. Bernays [ed.], Insect- plant interactions. CRC Press, Boca Raton, Florida. Eigenbrode, S. D. 1996. Plant surface waxes and insect behaviour, pp. 201-221. In G. Kerstiens [ed.], Plant cuticles. BIOS Scientific Publishers Ltd, Oxford. Gaum, W. G., J. H. Gilimore, and K. L. Pringle. 1994. Resistance of some rose cultivars to the western flower thrips, Frankliniella occidentalis (Thysanoptera: Thripidae). Bulletin of Entomological Research 84: 487-492. Helyer, N. L., and P. Brobyn. 1992. Chemical control of western flower thrips (Frankliniella occidentalis Pergande). Annals of Applied Biology 121: 219-231. Herron, G. A., and G. Gullick. 1998. Insecticide resistance in western flower thrips in Australia, pp. 164-170. In M. P. Zalucki, R. A. I. Drew and G. G. White [eds.], Sixth Australian Applied Entomological Research Conference. The Australian Entomological Society, The University of Queensland, Brisbane, Australia. Herron, G. A., and T. M. James. 2005. Monitoring insecticide resistance in Australian Frankliniella occidentalis Pergande (Thysanoptera: Thripidae) detects fipronil and spinosad resistance. Australian Journal of Entomology 44: 299-303. Houlding, B., and B. Woods. 1995. Mite and insect pests of strawberries. Farmnote 71/1995. Western Australia Department of Agriculture, South Perth, WA, Australia. Immaraju, J. A., T. D. Paine, J. A. Bethke, K. L. Robb, and J. P. Newman. 1992. Western flower thrips (Thysanoptera: Thripidae) resistance to insecticides in coastal California greenhouses. Journal of Economic Entomology 85: 9-14. Jensen, S. E. 2000a. Mechanisms associated with methiocarb resistance in Frankliniella occidentalis (Thysanoptera: Thripidae). Journal of Economic Entomology 93: 464-471. Jensen, S. E. 2000b. Insecticide resistance in the western flower thrips, Frankliniella occidentalis. Integrated Pest Management Reviews 5: 131-146. Jensen, S. E. 2000c. Insecticide resistance in the western flower thrips, Frankliniella occidentalis, pp. 101, Department of Life Science and Chemistry. Roskilde University, Roskilde, Denmark. Kindt, F., N. N. Joosten, D. Peters, and W. F. Tjallingii. 2003. Characterisation of the feeding behaviour of western flower thrips in terms of electrical penetration graph (EPG) waveforms. Journal of Insect Physiology 49: 183-191. Kirk, W. D. J., and L. I. Terry. 2003. The spread of the western flower thrips Frankliniella occidentalis (Pergande). Agricultural and Forest Entomology 5: 301-310. Kumar, N. K. K., D. E. Ullman, and J. J. Cho. 1993. Evaluation of Lycopersicon germ plasm for tomato spotted wilt tospovirus resistance by mechanical and thrips transmission. Plant Disease 77. Kumar, N. K. K., D. E. Ullman, and J. J. Cho. 1995a. Resistance among Lycopersicon species to Frankliniella occidentalis (Thysanoptera: Thripidae). Journal of Economic Entomology 88: 1057-1065. Kumar, N. K. K., D. E. Ullman, and J. J. Cho. 1995b. Frankliniella occidentalis (Thysanoptera: Thripidae) landing and resistance to tomato spotted wilt tospovirus among Lycopersicon accessions with additional comments on Thrips tabaci (Thysanoptera: Thripidae) and Trialeurodes vaporariorum (Homoptera: Aleyrodidae). Environmental Entomology 24: 513-520. Lewis, T. 1973. Thrips: Their Biology, Ecology, Evolution and Economic Importance. Academic Press, London and New York. Lewis, T. [ed.] 1997a. Thrips as Crop Pests. CAB International, Wallingford, Oxon, U.K. Lewis, T. 1997b. Pest thrips in perspective, pp. 1-13. In T. Lewis [ed.], Thrips as crop pests. CAB International, Wallingford.

62 Chapter II: Preference and performance

Lewis, T. 1998. Pest thrips in perspective, pp. 385-390, Proceedings The 1998 Brighton Conference- Pest and Diseases. British Crop Protection Council, Brighton, UK. Malipatil, M. B., A. C. Postle, J. A. Osmelak, M. Hill, and J. Moran. 1993. First record of Frankliniella occidentalis (Pergande) (Thysanoptera: Thripidae) in Australia. Australian Journal of Entomology 32: 378. Maris, P. C., N. N. Joosten, D. Peters, and R. W. Goldbach. 2003. Thrips Resistance in Pepper and Its Consequences for the Acquisition and Inoculation of Tomato spotted wilt virus by the Western Flower Thrips. Phytopathology 93: 96-101. Maris, P. C., N. N. Joosten, R. W. Goldbach, and D. Peters. 2004. Decreased preference and reproduction, and increased mortality of Frankliniella occidentalis on thrips-resistant pepper plants. Entomologia Experimentalis et Applicata 113: 149-155. Marullo, R., and E. Tremblay. 1993. Le specie italiane del genere Frankliniella Karny. Inf. Fitopat 11: 37-44. Medhurst, A., and B. Swanson. 1999. WFT insecticide management plans: capsicum, cucumber, lettuce, ornamentals, Strawberries and tomato, pp. 16-27, Western Flower Thrips Newsletter, December 1999. National Strategy for the Management of Western Flower Thrips and Tomato Spotted Wilt Virus. Medhurst, A., and M. Y. Steiner. 2001. Western Flower Thrips and Strawberries. National Strategy for the Management of WFT & TSWV, East Melbourne, Victoria, Australia. Ohta, I. 2002. Host plant resistance in Japanese chrysanthemums against Frankliniella occidentalis (Thysanoptera: Thripidae) during the non-flowering stage. Applied Entomology and Zoology 37: 271-277. Pergande, T. 1895. Observation on certain Thripidae. Insect Life 7: 390-395. Persley, D., M. Sharman, J. Thomas, I. Kay, S. Heisswolf, and L. McMichaell. 2009. Thrips and tospoviruses: a management guide. Department of Primary Industries and Fisheries (DPI&F). Pfleger, F. L., M. E. Ascerno, and R. Wawrzynski. 1995. Tomato spotted wilt virus. Minnesota Commercial Flower Growers Bulletin 44: 1-7. Phillips, D., and A. Reid. 2008. New strawberry varieties for WA - trial results in 2005, 2006 and 2007, pp. 9. Department of Agriculture and Food Western Australia, Kensington, South Perth WA. Robb, K., and M. Parella. 1989. An integrated approach to preventing western flower thrips and TSWV in the greenhouse. Grower Talks 53: 26-32. Robb, K. L. 1989. Analysis of Frankliniella occidentalis (Pergande) as a pest of floricultural crops in California greenhouses, pp. 135pp. University of California, Riverside, CA, California. Robb, K. L., and M. P. Parrella. 1995. IPM of western flower thrips, pp. 365-370. In B. L. Parker, M. Skinner and T. Lewis [eds.], Thrips biology and management. Plenum Press, New York. Roda, A. L., N. J. Oldham, A. Svatos, and I. T. Baldwin. 2003. Allometric analysis of the induced flavonols on the leaf surface of wild tobacco (Nicotiana attenuata). Phytochemistry 62: 527-536. Rummel, D. R., and J. E. Quisenberry. 1979. Influence of thrips injury on leaf development and yield of various cotton types. Journal of Economic Entomology 72: 706-709. SAS 2002-2003. SAS 9.1 computer program, version 9.1. By SAS, Cary, NC, USA. Saxena, K. N. 1985. behavioural basis of plant susceptibility to insects. Insect Science and Its Application 6: 303-313. Schoonhoven, L. M., T. Jermy, and J. J. A. van Loon. 1998. Insect-Plant Biology. Chapman & Hall, London. Shipp, J. L., G. J. Boland, and L. A. Shaw. 1991. Integrated pest management of disease arthropod pests of greenhouse vegetable crop in Ontario: Current status and future possibilities. Canadian Journal of Plant Science 71: 887-914. Snook, M. E., R. E. Lynch, A. K. Culbreath, and C. E. Costello. 1994. 2,3-Di-(E)-caffeoyl- (2R, 3R)-(+)-tartaric acid in terminals of peanut (Arachis hypogaea L.) varieties with different resistances to late leaf spot disease (Cercosporidium personatum) (Berk. and M. A. Deighton) and the insects tobacco thrips (Frankliniella fusca (Hinds))and potato

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leafhopper (Empoasca fabae (Harris). Journal of Agriculture and Food Chemistry 42: 1572-1574. Soria, C., and C. Mollema. 1995. Life-history parameters of western flower thrips on susceptible and resistant cucumber genotypes. Entomologia Experimentalis et Applicata 74: 177-184. Steiner, M. Y., and S. Goodwin. 2005. Management of western flower thrips, Frankliniella occidentalis (Pergande) (Thysanoptera: Thripidae), in hydroponic strawberry crops: using yellow sticky traps to determine action thresholds. Australian Journal of Entomology 44: 288-292. Steinite, I., and G. Levinsh. 2003. Possible role of trichomes in resistance of strawberry cultivars against spider mite. Acta Universitatis Latviensis 662: 59-65. Stoner, K. A., and A. M. Shelton. 1986. Studies on resistance to Thrips tabaci in four commercial varieties of cabbage, Cruciferae Newsletter 11:101. Tommasini, M. G., and S. Maini. 1995. Frankliniella occidentalis and other thrips harmful to vegetable and ornamental crops in Europe, pp. 1-42. In A. J. M. Loomans, J. C. Van Lenteren, M. G. Tommasini, S. Maini and J. Riudavets [eds.], Biological Control of Thrips, Wageningen Agricultural University, Wageningen, the Netherlands. Toshio, K. 2004. Varietal difference in the susceptibility of strawberry to western flower thrips Frankliniella occidentalis (Pergande). Kyushu Plant Protection Research 50: 55-61. Trichilo, P. J., and T. F. Leigh. 1988. Influence of resource quality on the reproductive fitness of flower thrips (Thysanoptera: Thripidae). Annals of the Entomological Society of America 81: 64-70. Ullio, L. 2002. Australia's national strategy for the management of western flower thrips (WFT), Frankliniella occidentalis (Pergande), pp. 687-689. In T. Hietaranta, M.-M. Linna, P. Palonen and P. Parikka [eds.], Proceedings of the fourth International Strawberry Symposium. Acta Horticulturae, MTT Agrifood Research, Finland. Ullman, D. E. 1996. Thrips and tospoviruses; advances and future directions, pp. 310-324. In G. Kuo [ed.], Symposium on the tospoviruses and thrips of floral and vegetable crops. Acta Horticulture, Taichung, Taiwan. van Lenteren, J. C., and L. P. J. J. Noldus. 1990. Whitefly- plant relationship: behavioural and ecological aspects, pp. 47-89. In D. Gerling [ed.], Whiteflies: their bionomics, pest status and management. Intercept, Andover, UK. Wagner, G. J. 1991. Secreting glandular trichomes: more than just hairs. Plant Physiology 96: 675-679. Yang, G., K. E. Espelie, J. W. Todd, A. K. Culbreath, R. N. Pittman, and J. W. Demski. 1993. Cuticular lipids from wild and cultivated peanuts and the relative resistance of these peanut species to fall armyworms and thrips. Journal of Agriculture and Food Chemistry 41: 814-818. Zar, J. H. 1999. Biostatistical Analysis. Prentice Hall International, Upper Saddle River, New Jersey. Zhao, G. Y., W. Liu, and C. O. Knowles. 1994. Mechanisms associated with diazinon resistance in western flower thrips. Pesticide Biochemistry and Physiology 49: 13-23. Zhao, G. Y., W. Liu, J. M. Brown, and C. O. Knowles. 1995. Insecticide resistance in field and laboratory strains of western flower thrips (Thysanoptera: Thripidae). Journal of Economic Entomology 88: 1164-1170.

64 CHAPTER III

Effect of spinosad and predatory mites (Acari) on western flower thrips, Frankliniella occidentalis (Pergande) (Thysanoptera: Thripidae) in three strawberry cultivars [Fragaria x ananassa Duchesne (Rosaceae)]

Keywords: Frankliniella occidentalis, Typhlodromips montdorensis, Neoseiulus cucumeris, Hypoaspis miles, spinosad, strawberry cultivars

Abstract

Western flower thrips, Frankliniella occidentalis (Pergande) (Thysanoptera: Thripidae) is a major pest of strawberry, Fragaria ananassa Duchesne (Rosaceae) in Australia. Spinosad (Success™, Dow AgroSciences Australia Ltd) is the only insecticide currently registered in Australia that is efficacious against F. occidentalis, and regarded to be compatible in an integrated pest management program. This study sought to determine whether three predatory mite species, Typhlodromips montdorensis (Schicha) (Phytoseiidae), Neoseiulus cucumeris (Oudemans) (Phytoseiidae) and Hypoaspis miles (Berlese) (Laelapidae) could be used with spinosad and least-preferred cultivars for the management of F. occidentalis. Typhlodromips montdorensis and N. cucumeris attack first instar thrips whilst H. miles feeds on thrips pupae. In the glasshouse, three strawberry cultivars (Camarosa, Camino Real, and Albion) were sprayed once with spinosad (80 mL/100 L rate, 0.096 g a.i./L) or water (control). Thrips adults were released onto plants 24 h after spraying and predatory mites released six days later. Spinosad significantly reduced thrips numbers compared to water. Typhlodromips montdorensis reduced thrips numbers as did N. cucumeris and H. miles. Spinosad had no effect on predatory mites. The number of H. miles could not be counted directly, but the numbers of thrips in treatments with H. miles were lower than those in treatments without the mite. Thrips numbers were lowest on Camino Real followed by Albion and Camarosa. These results suggest that the use of Camino Real with spinosad applications can be used to reduce initial F. occidentalis thrips numbers, followed by releases of predatory mites.

3. 1 Introduction

Western flower thrips (WFT), Frankliniella occidentalis (Pergande) (Thysanoptera: Thripidae) is regarded one of the most important economic pests worldwide (Brødsgaard and Albajes 1999, Jones et al. 2002, Kirk and Terry 2003), causing extensive crop losses of agricultural and horticultural crops (Lewis 1998). With its piercing-sucking mandibles, WFT penetrates Chapter III: Effect of spinosad, cultivar and predatory mites epidermal and sub-epidermal cells, causing extensive damage; this includes deformation and growth reduction of the plant, and silver scars on fruits and leaves (Hunter and Ullman 1989, Ullman et al. 1989, van Dijken et al. 1994, de Jager et al. 1995). In addition to direct damage, WFT cause indirect damage by transmitting tospoviruses (German et al. 1992, Wijkamp et al. 1996, Ullman et al. 1997). Since its detection in 1993 in Western Australia (Malipatil et al. 1993), WFT has spread to each state and territory, except the Northern Territory and has become a major pest in several crops including strawberry, Fragaria x ananassa Duchesne. In Australia, strawberry production has grown steadily in the last few years; however, its production is often affected by direct damage by WFT. This includes fruit deformation, uneven ripening, early withering of stigmas and anthers, and reduction in flower receptacle size leading to yield loss (Medhurst and Steiner 2001, Zalom et al. 2001, Coll et al. 2007). Similar damage occurs in low tunnel-, glasshouse- and field-grown strawberries (Ullio 2002).

Whilst insecticides are the main control method (Herron and Cook 2002), because of its small size, secretive habit, high reproductive potential, and ability to develop resistance to insecticide, WFT can be difficult to control (Jensen 2000). It has developed resistance to several major classes of chemicals (Brødsgaard 1994, Broadbent and Pree 1997, Jensen 1998, Jensen 2000) in different parts of the world. Spinosad™(Dow AgroSciences, USA) is a mixture of tetracyclic- macrolide compounds and is classified as a reduced-risk bioinsecticide (Sparks et al. 1998). It has been used in over 180 different crops to control a wide range of pests, and is effective against WFT (Funderburk et al. 2000). Spinosad is relied on heavily by growers in Australia, but WFT has already begun to develop resistance to this chemical (Herron and James 2005). WFT resistance to insecticides appears to be a serious threat to its effective management; therefore, additional management strategies are clearly necessary.

Overseas, predatory mites (Acari) are used to manage WFT in field and glasshouse crops (Chant 1985, van Lenteren and Woets 1988, McMurtry and Croft 1997). Until recently, Australian growers had no effective biological control options for WFT due to quarantine restrictions prohibiting their importation (Steiner and Goodwin 2001). Recently four predatory mite species have become available to Australian growers. Of them, two species are natives: Typhlodromips montdorensis (Schicha) and Typhlodromus occidentalis (Schicha) (Phytoseiidae) (Steiner and Goodwin 2000). Neoseiulus cucumeris (Oudemans) (Phytoseiidae) and Hypoaspis miles (Berlese) (Laelapidae) are native to New Zealand and were recently confirmed as occurring in Australia. Of the four mite species, only T. montdorensis, N. cucumeris and H. miles have been used for WFT management (Anonymous 2006). However, augmentative releases of predatory mites are not always sufficient to manage WFT in crops with a low economic threshold such as strawberry (Gillespie and Ramey 1988, Bakker and Sabelis 1989, Gillespie 1989).

66 Chapter III: Effect of spinosad, cultivar and predatory mites

It is possible to integrate biological and chemical control when mites are used as natural enemies (Reuveni 1995, Kongchuensin and Takafuji 2006). However, there are concerns about the detrimental effects of insecticides on predatory mites. The negative impact of pesticides have been reported on several predatory mite species including Phytoseiulus persimilis Athias- Henriot, Typhlodromus pyri (Scheuten), Amblyseius finlandicus (Oudemans), A. potentillae (Garman), Stratiolaelaps scimitus (Womersley) and Neoseiulus womersleyi (Schicha) (Hassan et al. 1987, Hassan et al. 1988, Kim and Paik 1996, Bowie et al. 2001, Kim and Seo 2001, Amano et al. 2004, Cabrera et al. 2004, Li et al. 2006, van Driesche et al. 2006). Pesticides can affect behaviour, survival, reproduction, attack-rate, and prey handling time (Li et al. 2006, van Driesche et al. 2006).

Applications of insecticide followed by releases of natural enemies are one proposed solution to this problem. Bentz and Neal (1995) reported that an initial application of a natural insecticide [derived from Nicotiana gossei] followed by the release of the hymenopteran parasitoid, Encarsia formosa Gahan (Aphelinidae) is effective for the management of whitefly in glasshouse-grown tomato. An initial application of Acramite® (miticide) followed by the release of Neoseiulus californicus (McGregor) (Phytoseiidae) was reported by Rhodes and Liburd (2006) as an effective strategy to control spider mite in glasshouse- and field-grown strawberry. Spraying of the selective pesticide pyriproxyfen, an insect growth regulator, followed by the release of the predatory bug, Orius sauteri (Poppius) (Anthocoridae), is common practice for the management of spider mite in eggplant in Japan (Nagai 1990).

Spinosad can provide effective control of many insect pests, while generally having reduced toxicity to natural enemies (Brunner et al. 2001, Elzen 2001, Villanueva and Walgenbach 2005b). However, selectivity of spinosad on predators is under review (Pietrantonio and Benedict 1999, Williams et al. 2003). It is regarded to have low to moderate toxicity to predatory mites (Pietrantonio and Benedict 1999, Ludwig and Oetting 2001) and the toxicity varies from species to species (Williams et al. 2003, Cote et al. 2004, Jones et al. 2005). van Driesche et al. (2006) reported that fresh residues of the recommended rate of spinosad for WFT on glasshouse flower crops had no toxic effect on N. cucumeris, while it lowered the survival of Iphiseius degenerans (Berlese). Holt et al. (2006) reported that an application of spinosad for the control of two-spotted spider mite had no effect on the predatory mite P. persimilis, one day before or after its release. Spinosad is highly toxic to Neoseiulus fallacis (Garman), the most abundant predaceous mite in North Carolina apple orchards that prey on the mite pests Panonychus ulmi (Koch) and Tetranychus urticae Koch (Tetranychidae) (Villanueva and Walgenbach 2005a). Despite its detrimental effect on some species, spinosad can be integrated with biological control for WFT management (Funderburk et al. 2000, Ludwig

67 Chapter III: Effect of spinosad, cultivar and predatory mites and Oetting 2001) if a period of time between spray and release is maintained (Jones et al. 2005, Khan and Morse 2006, Kongchuensin and Takafuji 2006).

The integration of spinosad and commercially available predatory mites in Australia for the management of WFT has not been evaluated. Given that cultivar has an effect on thrips survival (Chapter 2), the objective of this study was to investigate the effectiveness of commercially available predatory mites [T. montdorensis, N. cucumeris, and H. miles] and the influence of strawberry cultivars (Camarosa, Camino Real and Albion), with or without a spinosad application on WFT.

3.2 Materials and Methods

The trial was conducted in a glasshouse (25±2⁰C, 60-70% RH, 16:8 L: D regimes) at The University of Western Australia (UWA) from September to November 2007.

3.2.1 Source cultures

3.2.1.1 Strawberry cultivars

Strawberry [Fragaria x ananassa Duchesne (Rosaceae)] cv Camarosa and Camino Real (short-day), and Albion (day-neutral) were used in experiments. Strawberry runners were obtained from a commercial grower in June 2006, and propagated in pots (32.5l x 32.5w x 40.5h cm) with potting mix (Baileys Fertilisers, Rockingham, WA) in

Figure 3.1 Plant covered by a modified cage made from thrips-proof mesh (105µ; Sefar Filter Specialists Pty Ltd., Malaga, W. Australia).

68 Chapter III: Effect of spinosad, cultivar and predatory mites glasshouses at the Department of Agriculture and Food Western Australia (DAFWA) and UWA. All potted plants were covered with thrips-proof cages (45 x 35 cm) made from mesh net (105µ; Sefar Filter Specialists Pty Ltd., Malaga, WA; Figure 3.1) and supported by steel- rod stands. At the beginning, all runners propagated were checked daily for any insects including WFT and any disease symptoms. Plants with any insect or with disease symptoms were removed from the glasshouses. This was done to ensure healthy plants used in experiments in later stages. All pots were fitted with a sprinkler operated with automatic timers. During summer, plants were watered every third day, while in winter and spring watering was once a week. A liquid fertiliser (Thrive®, Yates, Australia; NPK: 12.4: 3: 6.2; rate: 5mL/2 L water) was applied once a month.

3.2.1.2 Western flower thrips (WFT)

WFT was reared on calendula, Calendula officinalis L. (Asteraceae), planted in plastic pots (50x100 mm) with potting mix (Baileys Fertilisers, Rockingham, WA) kept in an insect-proof Perspex cage (500 mm high, 420 mm deep and 400 mm wide) with 105 µ mesh net fitted on a Nylex tote box (320mm x 420mm; Blyth Enterprises Pty Ltd, Australia). Plants with WFT were kept in a glasshouse at UWA from July 2006 to November 2008. All pots were fitted with sprinklers operated with automatic timers. Plants were watered every third day during summer and once a week in winter and spring. Every second week, adults were collected from caged plants and released onto new potted plants at the base to ensure the continuous availability of WFT.

To obtain uniformly aged WFT for this trial, adults (20 individuals) were collected and released onto fresh caged plants and allowed to lay eggs for 24 h. After 24 h, all adults (20 individuals) were removed with a small aspirator. Plants were then checked daily for larvae. The newly hatched larvae were removed, and released onto a strawberry leaf that was placed on a moistened filter paper in a Petri dish (150 x 15 mm). The time and date of collection were recorded. Larvae that hatched on the same day were grouped together and maintained on strawberry leaves in Petri dishes until pupation. Strawberry leaves were changed as required. Pupae were maintained on a moistened filter paper in a Petri dish until adult emergence. Adults that emerged on the same day were used in this trial.

3.2.1.3 Predatory mites

Predatory mites [T. montdorensis, N. cucumeris and H. miles] used in the study were obtained from commercial suppliers (Biological Services, SA; Chilman IPM Services,

69 Chapter III: Effect of spinosad, cultivar and predatory mites

WA; and Beneficial Bug Company, NSW). Mites were provided in plastic buckets containing vermiculite. Trials were conducted immediately upon receipt of mites.

3.2.2 Glasshouse experiment: effect of cultivars and predatory mites with or without spinosad on western flower thrips

In order to evaluate the effect of cultivar and efficacy of predatory mites with or without spinosad on WFT, an experiment with a split plot design with factors: cultivar, spray treatment and predatory mite was carried out in a glasshouse (25 ± 2 ⁰C, 60-70% RH) at UWA. Eighty potted strawberry plants of each cultivar (Camarosa, Camino Real and Albion) at the 2-3 leaf stage (2-3 weeks old) were divided into two groups. One group of plants was sprayed with a spinosad solution (80 mL/100 L rate, 0.096 g a.i./L) using a hand-held atomiser (Hills Sprayers, BH220063) until run-off. The other group of plants was sprayed with distilled water (control). All plants were then covered with a thrips-proof cage as described in 3.2.1.1. The bottom end of the cage was secured with tape. The top end of the cage was closed with a rubber band.

WFT adults and predatory mites were collected with an aspirator from the stock culture, and kept in separate glass vials. On the appropriate release date, opened vials with WFT or predatory mites were placed at the base of the plant. Fifteen WFT adults (2 d old) were released onto each plant one day after spraying. Each group of sprayed plants from each cultivar with WFT adults was further divided into four groups (treatments), with 10 plants per treatment: (i) No mites, (ii) T. montdorensis, (iii) N. cucumeris, and (iv) H. miles. Six mites were released on each plant-receiving mites six days after the spray treatment. Each plant was considered a replicate. During experimental periods, experimental plants were kept flowerless. Plants were checked daily and any blooms developed were removed by a pair of sharp scissors.

Twenty-four h after WFT release, WFT adults and larvae on each plant were counted daily for three weeks. Each day, all leaves of each plant were checked thoroughly with a hand-held illuminated magnifying glass [50mm (2") diameter 2x power with 4x bifocal magnifier] and numbers of WFT adults and larvae were recorded. WFT counts were made between 0600 to 0800 h.

In order to determine the number of predatory mites (T. montdorensis and N. cucumeris) per plant, plants were cut at the base at the end of the trial and immediately placed in a container (500 ml) with 80% ethyl alcohol. Later, plant materials were sieved with a

70 Chapter III: Effect of spinosad, cultivar and predatory mites double-layer sieve (mesh size 105 µ) and checked under a stereomicroscope, and the numbers of T. montdorensis and N. cucumeris per plant were recorded. To determine the numbers of H. miles (soil dwelling), the plants were cut as described above, but the top soil (2-3 cm) from pots was also collected and placed immediately into a container with 80% ethyl alcohol. Later, the plant materials and the soil were sieved as mentioned above and were checked under stereomicroscope for H. miles. Unfortunately, no H. miles were recovered.

All trial pots were fitted with a sprinkler so that water did not reach the leaves and upper parts of the plant. This was intended to avoid any wash out of WFT or mites. Although the sprinkler was adjusted so that water could not reach plant leaf or upper parts, as an extra precaution, the watering time was set for the afternoon (1900 h) and plants were watered once per week.

3.2.2 Data analysis

To determine the effect of cultivars (Camarosa, Camino Real and Albion) and predatory mites (no mites, T. montdorensis, N. cucumeris and H. miles) with or without spinosad on WFT over time (days), adults and larvae were analysed separately with a repeated measures ANOVA (independent fixed variables: cultivars, spray treatments, predatory mites, sampling days; random variable: plant number; response variables: adults, larvae) (Proc Mixed Procedure). The data from days seven, 14, and 21 were used (end data point of each week). Due to significant interaction of cultivars, spray and predatory mites over time (days) on WFT adults and larvae, additional ANOVAs (repeated measures, Proc Mixed Procedure) were conducted for each cultivar separately (Quinn and Keough 2002). Because there was a significant interaction of spray and predatory mites over time, further ANOVAs (repeated measures, Proc Mixed Procedure) were conducted for each spray and each cultivar. Due to the number of post hoc multiple tests, an adjustment to the significance level was made [α = 0.00833= (0.05/3*2)]. If ANOVAs were significant, means were separated using least square means (SAS 2002-2003).

The influence of cultivars and spray on predatory mites (T. montdorensis and N. cucumeris) was determined by two-way ANOVAs (Proc Mixed Procedure) separately for T. montdorensis and N. cucumeris. The difference in mean numbers of T. montdorensis and N. cucumeris was assessed using two-sample t-test (Proc ttest Procedure) separately for spray treatments in each cultivar.

71 Chapter III: Effect of spinosad, cultivar and predatory mites

Square root was applied to meet the assumption of homogeneity of variances (Zar 1999). Data were reverse transformed for presentation. All statistical analyses were performed using Statistical Package SAS 9.1(SAS 2002-2003). Figures were constructed using Graphpad Prism 5.0 (GraphPad 2007).

3.3 Results

3.3.1 Western flower thrips

The numbers of WFT adults per plant were influenced by predatory mites (T. montdorensis, N. cucumeris, and H. miles) and cultivar (Camarosa, Camino Real and Albion) in the presence or absence of spinosad, and these effects changed over time

(Interaction: F = 9.2612, 432, P < 0.0001). Consequently, a series of ANOVAs for each cultivar (Camarosa: F 6, 144 = 9.75, P < 0.0001; Camino Real: F 6, 144 = 4.29, P = 0.0005;

Albion: F 6, 144 = 13.02, P < 0.0001) were performed (Appendix, 3.1). As interactions in these three-way ANOVAs were also significant, the effects of predatory mites on WFT adults were determined by further ANOVAs for each spray treatment in each cultivar (Appendix 3.1).

Before presenting the more detailed results, some general trends were apparent. Overall, mean numbers of WFT adults per plant were lower on plants treated with spinosad (19.95 ± 1.73) compared to water (32.96 ± 1.74) across the different mite treatments (Figure 3.2). The number of WFT on the plants with mites (T. montdorensis: 21.47 ± 1.66, N. cucumeris: 24.33 ± 1.86, H. miles 26.89 ± 2.02) was lower than those without mites (33.14 ± 1.56). There were fewest WFT on Camino Real (19.89 ± 0.94), followed by Albion (21.66 ± 1.06); most WFT were found on Camarosa (24.84 ± 1.25).

72 Chapter III: Effect of spinosad, cultivar and predatory mites

Spinosad Water 50 SE) 

40

30

20

10

0 Number of WFT adults per plant (Mean plant per adults WFT of Number No mites Tm Nc Hm

Figure 3.2 Mean numbers of WFT adults per plant treated with spinosad or water and either no predatory mites or one of three species of predatory mites. Within each group (X-axis), means were separated by LS means (α = 0.05). Tm = T. montdorensis, Nc = N. cucumeris, Hm = H. miles.

At day seven, the number of WFT adults on any of the cultivars in the spinosad-treated plants did not differ (Figure 3.3). At days 14 and 21 on plants sprayed with spinosad, predatory mite species reduced the number of WFT adults over time, regardless of cultivar (Camarosa F6,72 = 28.22, P < 0.0001, Camino Real F6,72 = 39.04, P < 0.0001,

73 Chapter III: Effect of spinosad, cultivar and predatory mites

No mites Tm Nc Hm 80 A

60 d c 40 c b a b a a 20 a a a a SE)

 0 80 B

60

40 d c c 20 b a b a a a a a ab

0 80 C Number of WFT adults per plant (Mean plant per adults WFT of Number 60

d 40 c c b b b a 20 a a a a a

0 7D 14D 21D

Figure 3.3 Effects of predatory mites on number of WFT adults per plant over time (7, 14, or 21 days after release of WFT) sprayed with spinosad on cultivars (A) Camarosa, (B) Camino Real and (C) Albion. Means with different letters within each day were significantly different from others (LS means, α = 0.00835). Tm = T. montdorensis, Nc = N. cucumeris, Hm = H. miles.

Albion F6, 72 = 19.00, P < 0.0001, Figure 3.3). For all three cultivars, plants with no mites had the highest numbers of WFT adults and differed significantly from the rest (Figure 3.3). The number of WFT adults was lowest when plants received T. montdorensis. However, on day 14, the numbers of WFT adults were not different on Camarosa (Figure 3.2A) or Camino Real (Figure 3.2B) with T. montdorensis and N. cucumeris, but were on Albion plants (Figure 3.2C). By day 21 there was a distinct

74 Chapter III: Effect of spinosad, cultivar and predatory mites difference between the numbers of WFT adults on all cultivars with each of the different mite species (Figure 3.3).

The response of WFT adults to the cultivars and predatory mites on those plants sprayed with water was similar to those on plants sprayed with spinosad. There was no difference in the mean number of WFT adults among any of the treatments on day seven (Figure 3.4), and mites reduced the number of thrips adults across all cultivars (Figure 3.3). There was, however, a difference in the effect of predatory mites on the number of

WFT adults across all three cultivars over time: Camarosa (F 6, 72 = 22.90, P < 0.0001),

Camino Real (F 6, 72 = 19.7, P < 0.0001) and Albion (F 6, 72 = 83.33, P < 0.0001) (Appendix 3.1, Figure 3.4). On days 14 and 21, plants of each cultivar with T. montdorensis had the lowest numbers of WFT adults. However, on day 14 in Camarosa and Camino Real, WFT adult numbers were not different between T. montdorensis- and N. cucumeris-treated plants, but did differ in Albion. Similarly, on day 14, WFT adult numbers were not different between Camarosa and Camino Real plants that received N. cucumeris and H. miles, but differed on Albion. By day 21 the mean numbers of WFT adults were different in all predatory mite treatments with a consistent trend for the least number of thrips being on the plants with T. montdorensis, followed by N. cucumeris and then H. miles (Figure 3.4 A, B, and C).

Overall, the number of WFT larvae per plant was lowest on plants sprayed with spinosad (20.67 ± 1.12) compared with those that received water only (31.38 ± 1.57) across the different mite treatments (Figure 3.5). The number of WFT larvae on the plants with mites (20.85 ± 1.14, 22.88 ± 1.19 and 27.59 ± 1.45 for T. montdorensis, N. cucumeris and H. miles respectively) was lower than those without mites (32.77 ± 1.60). There were fewest WFT larvae on Camino Real (22.41 ± 1.14), followed by Albion (26.53 ± 1.32). The highest number of WFT larvae was on Camarosa plants (29.13 ± 1.58). There was a significant interaction between the cultivars, spray, and predatory mites over time (days) (F 12, 432 = 2.31, P = 0.0073) that influenced WFT larvae numbers. Consequently, further ANOVAs for each cultivar were performed (Appendix 3.2).

75 Chapter III: Effect of spinosad, cultivar and predatory mites

No mites Tm Nc Hm 80 d A c 60 b c a 40 b a ab a a a 20 a

SE) 0  80 B

60 d c b 40 c a ab b a a a a 20 a

0 80 C

Number of WFT adults per plant (Mean plant per adults WFT of Number d 60 c b d a 40 c a b a a a a 20

0 7D 14D 21D

Figure 3.4 Effects of predatory mites on the number of WFT adults per plant over time (7, 14, or 21 days after release of WFT) sprayed with water on strawberry cultivar (A) Camarosa, (B) Camino Real and (C) Albion. Means with different letters within each day were significantly different from others (LS means, α = 0.00835). Tm = T. montdorensis, Nc = N. cucumeris, Hm = H. miles.

76 Chapter III: Effect of spinosad, cultivar and predatory mites

Spinosad Water 50 SE) 

40

30

20

10

0 Number of WFT larvae per plant (Mean plant per larvae WFT of Number No mites Tm Nc Hm

Figure 3.5 Mean numbers of WFT larvae per plant treated with spinosad or water and either no predatory mites or one of three species of predatory mites. Within each group (X-axis), means were significantly different (LS means, α = 0.05). Tm = T. montdorensis, Nc = N. cucumeris, Hm = H. miles.

An interaction of spray and predatory mites treatment and time was evident in each cultivar (Camarosa: F 6, 144 = 3.78, P = 0.001; Camino Real: F 6, 144 = 2.5,P = 0.022;

Albion: F 6, 144 = 2.76, P = 0.011); therefore, the impact of mite treatment and time on the number of WFT larvae was evaluated separately by an ANOVA for each spray in each cultivar (Appendix 3.2).

In those plants sprayed with spinosad, the number of WFT larvae was reduced in each cultivar over time (days) when predatory mites were present (F 6, 72 = 8.77Camarosa

7.25Camino Real 5.71Albion, P < 0.0001) (Appendix 3.2, Figure 3.6). In each cultivar, plants that received T. montdorensis had the lowest numbers of WFT larvae per plant, except in Albion on day seven, when the N. cucumeris-treated plant had the lowest numbers of WFT larvae (Figure 3.6C). However, on days seven and 14 in Camarosa (Figure 3.6A), on day 14 in Camino Real (Figure 3.6B) and on day seven in Albion (Figure 3.6C), mean numbers of WFT larvae were not different between the plants that received T. montdorensis and N. cucumeris. On the other hand, in each cultivar, WFT larvae numbers were highest on plants that received ‘no mites’. However, on day seven, in Camarosa as well as in Albion, there was no difference in WFT larvae between plants that received ‘no mites’ and H. miles (Figure 3.6).

77 Chapter III: Effect of spinosad, cultivar and predatory mites

No mites Tm Nc Hm 60 A

40 c d c b b b b a a a 20 a a SE)

 0 60 B

40

c d c 20 d c b b a a a a b

0 60 C Number of WFT larvae per plant (Mean plant per larvae WFT of Number

40 d d c c b b b b a 20 a a a

0 7D 14D 21D

Figure 3.6 Effects of predatory mites on WFT larvae per plant over times (7, 14, or 21 days after WFT release) sprayed with spinosad on strawberry cultivar (A) Camarosa, (B) Camino Real and (C) Albion. Means with different letters within each day were significantly different from others (LS means, α = 0.00835). Tm = T. montdorensis, Nc = N. cucumeris, Hm = H. miles.

Similar to the spinosad treatment, WFT larvae numbers per plant were significantly influenced by predatory mites (Camarosa: F 6, 72 = 3.99, P = 0.0017; Camino Real: F 6,

72 = 3.87, P = 0.0021; Albion: F 6, 72 = 3.59, P = 0.0036) (Appendix 3.2, Figure 3.7). Typhlodromips montdorensis-treated plants had the lowest numbers of WFT larvae, except on day seven in Albion, when the N. cucumeris-treated plants had the lowest numbers of WFT. However, on day 7 in Camarosa and Albion and on day 14 in Camino

78 Chapter III: Effect of spinosad, cultivar and predatory mites

Real, there was no difference in WFT larvae between plants that received T. montdorensis and N. cucumeris. On the other hand, at all times in each cultivar, mean numbers of WFT larvae were highest on plants that received ‘no mites’.

No mites Tm Nc Hm 60 d A c d b 40 c c a b b a a a 20 SE)

 0 60 B

d 40 c c b b c a a a b b 20 a

0 60 C

Number of WFT larvae per plant (Mean plant per larvae WFT of Number b d c 40 c a a c b b a a a 20

0 7D 14D 21D

Figure 3.7 Effects of predatory mites on WFT larvae per plant over times (7, 14, or 21 days after WFT release) sprayed with water on strawberry cultivar (A) Camarosa, (B) Camino Real and (C) Albion. Means with different letters within each day were significantly different from others (LS means, α = 0.00835). Tm = T. montdorensis, Nc = N. cucumeris, Hm = H. miles.

79 Chapter III: Effect of spinosad, cultivar and predatory mites

3.3.2 Predatory mites

The numbers of T. montdorensis and N. cucumeris per plant were unaffected by spinosad (F T. montdorensis = 3.57 1, 54, P = 0.06; F N. cucumeris = 0.052 1, 54, P = 0.116) or cultivar (F T. montdorensis = 1.11 2, 54, P = 0.336; F N. cucumeris = 0.51 2, 54, P = 0.604) (Figure 3.78). In both spray treatments, T. montdorensis numbers were higher than N. cucumeris

(Figure 3.9), but the difference was not significant in Camarosa (tspinosad = 0.78 18, P =

0.44; twater = 0.22 18, P = 0.83), Camino Real (tspinosad = 1.64 18, P = 0.12; twater = 1.33

18, P = 0.20) nor in Albion (tspinosad = 1.85 18, P = 0.08; twater = 1.61 18 P = 0.13).

A B 25 20 SE)  15 10 5 0

C D 25 20 15 10

Number of predatory mites (Mean 5 0 Camarosa Camino Real Albion Spinosad Water

Figure 3.8 Mean numbers of T. montdorensis (A and B) and N. cucumeris (C and D) per plant in relation to strawberry cultivar and spray treatment.

80 Chapter III: Effect of spinosad, cultivar and predatory mites

T. montdorensis N. cucumeris 25 A 20 SE)

 15

10

5

0 25 B 20

15

10

Number of predatory mites per plant (Mean plant per mites predatory of Number 5

0 Camarosa Camino Real Albion

Figure 3.9 Comparison of mean numbers of T. montdorensis and N. cucumeris per plant sprayed with (A) spinosad and (B) water.

3.4 Discussion

This study suggests that it is possible to apply spinosad and subsequently release predatory mites (T. montdorensis, N. cucumeris and H. miles) with a resultant reduction in WFT adults and larvae, and little or no negative effect on natural enemies. Control of WFT can be further improved by the use of resistant cultivars. As was presented in Chapter 2, it appears that strawberry cultivars had a significant influence on WFT numbers, with Camino Real being least favourable to WFT and Camarosa being most favourable. As in Chapter 2, this study did not explore the specific factors or mechanisms that may be responsible for differences in WFT adults. However, this study does demonstrate that the varieties interact with other tactics for control of WFT.

All three predatory mites were more successful in reducing WFT numbers in cultivar Camino Real than Albion or Camarosa, although the actual number of predatory mites (T. montdorensis and N. cucumeris) did not differ amongst cultivars. Brown et al. (1999) tested the predation efficacy of predatory mites, N. cucumeris and Iphiseius degenerans (Berlese) (Acari: Phytoseiidae) against WFT on 12 plant families and found that the WFT predation rate of these

81 Chapter III: Effect of spinosad, cultivar and predatory mites mites was different on different plant species. The variation in predation efficacy might be due to differences in plant architecture (Kareiva and Sahakian 1990), surface texture (Kareiva and Sahakian 1990) and plant chemistry (Price et al. 1980). The susceptibility of herbivores to predators is often related to the nutritional quality of plants on which the herbivores are feeding (Price et al. 1980). WFT numbers were low on the cultivar Camino Real, even in the absence of mites. The difference in WFT numbers on different cultivar-mite combinations suggests that cultivars not only influence WFT numbers, but also influence the effectiveness of predatory mites.

Regardless of cultivar, all three mite species reduced the numbers of WFT adults and larvae, but T. montdorensis appears to be the most effective species in suppressing WFT, followed by N. cucumeris and H. miles. The efficiency of natural enemies in a pest management program often varies from species to species of both predator and prey (Chyzik et al. 1996, Berndt et al. 2004a, Berndt et al. 2004b), and the reasons are likely be multi-factorial. These may include differences in prey preference, predation rate, distribution and population development (Chyzik et al. 1996, Berndt et al. 2004b). Bakker and Sabelis (1989) reported that N. cucumeris is able to predate on only the smallest thrips, which effectively limits them to attacking first instar WFT larvae. Similarly, T. montdorensis feeds on first instar WFT larvae. The other mite species tested, H. miles, inhabits the top soil layer (at about 1.3 cm depth) and preys only on WFT pupae (Glockemann 1992), although some studies suggest that this mite also preys on late second instar larvae (Berndt 2002, Berndt 2003). Berndt et al. (2004b) found that the predatory mite, Hypoaspis aculeifer (Canestrini) (Acari: Laelapidae) is more effective against WFT soil dwelling stages compared to Stratiolaelaps (=Hypoapsis) miles (Berlese) (Acari: Laelapidae), mainly because the predation rate of H. aculeifer is higher than S. miles. Similarly, Brødsgaard (1989) and van Houten et al. (1995) reported that N. cucumeris consumed higher numbers of WFT larvae compared to Amblyseius barkeri (Swirskii) (Acari: Phytoseiidae). Neoseiulus cucumeris is reported to feed on an average of six WFT larvae (first instar) per day (Zilahl- Balogh et al. 2007), whilst T. montdorensis feeds on 7-14 larvae per day (Steiner et al. 2003). Rhodes and Liburd (2006) also reported variation in performance of predatory mites against two-spotted spider mites (Tetranychus urticae) in field-grown strawberry. Rhodes and Liburd (2006) found that Neoseiulus californicus and Phytoseiulus persimilis both significantly reduced T. urticae, though their performance varied from each other. Phytoseiulus persimilis took longer (one week) than N. californicus to bring two-spotted spider mites under control (<10 mite per leaflet, ETL). Wiethoff et al. (2004) also found that the efficacy of predatory mites against WFT varied. In cucumber, N. cucumeris is more successful in reducing WFT numbers than H. aculeifer (Wiethoff et al. 2004).

82 Chapter III: Effect of spinosad, cultivar and predatory mites

The distribution of predatory mites often influences their effectiveness in pest management programs. Typhlodromips montdorensis is a generalist predator and has the ability to distribute rapidly on different plant parts (Steiner and Goodwin 1998, 2001) which may give this predator an advantage in reducing WFT over other predatory mites. However, it has been reported that the within-plant distribution of N. cucumeris is uneven and it prefers the lower part of the plant, while WFT prefer to remain on the upper part of the plants (cucumber) (Messelink et al. 2006). Variation in population development rate among different predatory mites might also account for the variation in effectiveness (Messelink et al. 2006). The present results indicate that T. montdorensis numbers were slightly, but not significantly, higher than N. cucumeris. However, even this small difference might contribute to a difference in effectiveness of WFT suppression. This study did not explore the specific factors that influence the effectiveness of predatory mites in suppressing WFT; nonetheless, all three predatory mites provide some control of WFT. Typhlodromips montdorensis seems to be the most effective. However, to maximise the efficiency of predatory mites in WFT management, future research is needed to evaluate factors that influence predatory mite efficiency.

In the present study, the numbers of WFT were lower in treatments with one of the mite species and when spinosad was applied. Spinosad was not detrimental when applied before mite releases. This suggests that spinosad had no adverse effects on predatory mites at least seven days after insecticide application at the recommended rate. Thoeming and Poehling (2006) reported that an application of the botanical azadirachtin (NeemAzal-U, 17% azadirachtin) with two predatory mite species reduced the numbers of WFT by up to 99% without causing any significant harm to predatory mites. The previous findings of Ludwig and Oetting (2001) and Ludwig (2002) indicated that the combined application of spinosad and the predatory anthocorid bug, Orius insidiosus Say (Heteroptera: Anthocoridae) significantly reduced WFT numbers in glasshouse potted chrysanthemums and marigold, compared to the control (without spinosad or Orius). Ludwing and Oetting (2001), Ludwig (2002), and Funderburk et al. (2000) demonstrate that spinosad had no or little effect on O. insidiosus. However, laboratory studies by Elzen et al. (1998) and Pietrantonia and Benedict (1999) suggest that spinosad has low toxicity to O. insidiosus. Similarly, Kongchuensin and Takafuji (2006) reported that there was a significant negative effect of spinosad on eggs and the immature stage of the predatory mite N. longispinosus, if exposed to fresh residues within 48 hrs. However, there was no or very little negative influence of spinosad on adults, eggs or the immature stage of N. longispinosus, if exposed seven days after spinosad is applied (Kongchuensin and Takafuji 2006).

In conclusion, the management of WFT can be improved by releasing predatory mites after a spinosad application without any apparent detrimental effect. Moreover, again, Camino Real

83 Chapter III: Effect of spinosad, cultivar and predatory mites appears to be the least preferred cultivar to WFT. Of the three predatory mite species, T. montdorensis in conjunction with spinosad appears to be the most effective combination for the management of WFT. However, because of the low economic threshold of WFT in strawberry, single species releases of mites such as T. montdorensis may not be sufficient to suppress WFT. Multiple species release of predatory mites (Wiethoff et al. 2004, Premachandra et al. 2005) combined with a spinosad application could be a better strategy for WFT management. Thus, it is worthwhile to test the compatibility of T. montdorensis, N. cucumeris, and H. miles in multiple releases with spinosad. In addition, increasing the numbers of predatory mites (decreasing the ratio of predatory mites to WFT) and using a shorter period between spray application and predatory mite release could further enhance WFT control. However, a shorter time lapse between application and predatory mite release could be toxic to predatory mites. Thus, a study is required to evaluate the residual toxicity of spinosad to predatory mites. This study was a small-scale study (plants covered with mesh net) and so the conclusions that can be drawn at this point are somewhat limited. A large-scale experiment in either the glasshouse, low tunnel environment or both is needed to confirm the present findings before any recommendation to commercial growers can be made.

3.5 Literature cited

Amano, H., Y. Ishii, and Y. Kobori. 2004. Pesticide susceptibility of two dominant phytoseiid mites, Neoseiulus californicus and N. womersleyi, in conventional Japanese fruit orchards (Gamasina: Phytoseiidae). Journal of Acarological Society of Japan 13: 65-70. Anonymous. 2006. Typhlodromus occidentalis. Biological Services, Loxton, Australia. Bakker, F. M., and M. W. Sabelis. 1989. How larvae of Thrips tabaci reduce the attack success of phytoseiid predators. Entomologia Experimentalis et Applicata 50: 47-51. Bentz, J.-A., and J. W. Neal Jr. 1995. Effect of a natural insecticide from Nicotiana gossei on the whitefly parasitoid Encarsia formosa (Hymenoptera: Aphelinidae). Journal of Economic Entomology 88: 1611-1615. Berndt, O. 2002. Entomopathogenic nematodes and soil-dwelling predatory mites: suitable antagonists for enhanced biological control of Frankliniella occidentalis (Pergande) (Thysanoptera: Thripidae)?, pp. 128, Von dem Fachbereich Gartenbau. University of Hannover, Hannover, Germany. Berndt, O. 2003. Entomopathogenic nematodes and soil-dwelling predatory mites: suitable antagonists for enhanced biological control of Frankliniella occidentalis (Pergande) (Thysanoptera: Thripidae)?, pp. 128, Von dem Fachbereich Gartenbau. University of Hannover, Hannover, Germany. Berndt, O., R. Meyhofer, and H.-M. Poehling. 2004a. The edaphic phase in the ontogenesis of Frankliniella occidentalis and comparison of Hypoaspis miles and Hypoaspis aculeifer as predators of soil-dwelling thrips stages. Biological Control 30: 17-24. Berndt, O., H.-M. Poehling, and R. Meyhofer. 2004b. Predation capacity of two predatory laelapid mites on soil-dwelling thrips stages. Entomologia Experimentalis et Applicata 112: 107-115. Bowie, M. H., S. P. Worner, O. E. Krips, and D. R. Penman. 2001. Sublethal effects of esfenvalerate residues on pyrethroid resistant Typhlodromus pyri (Acari: Phytoseiidae) and its prey Panonychus ulmi and Tetranychus urticae (Acari: Tetranychidae). Experimental and Applied Acarology 25: 311-319.

84 Chapter III: Effect of spinosad, cultivar and predatory mites

Broadbent, A. B., and D. J. Pree. 1997. Resistance to insecticides in populations of Frankliniella occidentalis (Pergande) (Thysanoptera: Thripidae) from greenhouses in the Niagara region of Ontario. Canadian Entomologist 129: 907-913. Brødsgaard, H. F. 1989. Frankliniella occidentalis (Thysanoptera; Thripidae) - a new pest in Danish glasshouse. Tidsskr. Planteavl. 93: 83-91. Brødsgaard, H. F. 1994. Insecticide resistance in Europe and African strains of western flower thrips (Thysanoptera: Thripidae) tested in a new residue-on-glass test. Journal of Economic Entomology 87: 1141-1146. Brødsgaard, H. F., and R. Albajes. 1999. Insects and mite pests, pp. 48-60. In R. Albajes, M. L. Guillino, J. C. Van Lenteren and Y. Elad [eds.], Integrated pest management in greenhouse crops. Kluwer Academic press, Netherlands. Brown, A. S. S., M. S. J. Simmonds, and W. M. Blaney. 1999. Influence of species of host plants on the predation of thrips by Neoseiulus cucumeris, Iphiseius degenerans and Orius laevigatus. Entomologia Experimentalis et Applicata 92: 283-288. Brunner, J. F., J. E. Dunley, M. D. Doerr, and E. H. Beers. 2001. Effect of pesticides on Colpoclypeus florus (Hymenoptera: Eulophidae) and Trichogramma platneri (Hymenoptera: Trichogrammatidae), parasitoids of Leafrollers in Washington. Journal of Economic Entomology 94: 1075-1084. Cabrera, A. R., R. A. Cloyd, and E. R. Zaborski. 2004. Effects of Greenhouse Pesticides on the Soil-Dwelling Predatory Mite Stratiolaelaps scimitus (Acari: Mesostigmata: Laelapidae) Under Laboratory Conditions. Journal of Economic Entomology 97: 793- 799. Chant, D. A. 1985. The Phytoseiidae, pp. 3-32. In W. Helle and M. W. Sabelis [eds.], Spider mites 1B. Elsevier, Amsterdam. Chyzik, R., I. Glazer, and M. Klein. 1996. Virulence and efficacy of different entomopathogenic nematode species against western flower thrips (Frankliniella occidentalis). Phytoparasitica 24: 103-110. Coll, M., S. Shakya, I. Shouster, Y. Nenner, and S. Steinberg. 2007. Decision-making tools for Frankliniella occidentalis management in strawberry: consideration of target markets. Entomologia Experimentalis et Applicata 122: 59-67. Cote, K. W., P. B. Schultz, and E. E. Lewis. 2004. Using acaricides in combination with Phytoseiulus persimilis Athias-Henriot to suppress Tetranychus urticae Koch populations. Journal of Entomological Science 39: 267-274. de Jager, C. M., R. P. T. Butot, and J. A. Guldemond. 1995. Genetic variation in chrysanthemum for resistance to western flower thrips and Thrips tabaci, pp. 403-406. In B. L. Parker, M. Skinner and T. Lewis [eds.], Thrips biology and management. Plenum Press, New York. Elzen, G. W. 2001. Lethal and Sublethal Effects of Insecticide Residues on Orius insidiosus (Hemiptera: Anthocoridae) and Geocoris punctipes (Hemiptera: Lygaeidae). Journal of Economic Entomology 94: 55-59. Elzen, G. W., P. J. Elzen, and E. G. King. 1998. Laboratory toxicity of insecticide residues to Orius insidiosus, Geocoris punctipes, Hippodamia convergens and Chrysoperla carnea. Southwestern Entomologist 23: 335-342. Funderburk, J. E., J. Stavisky, and S. Olson. 2000. Predation of Frankliniella occidentalis (Thysanoptera: Thripidae) in field peppers by Orius insidiosus (Hemiptera: Anthocoridae). Environmental Entomology 29: 376-382. German, T. L., D. E. Ullman, and J. W. Moyer. 1992. Tospoviruses: Diagnosis, Molecular Biology, Phylogeny, and Vector Relationships. Annual Review of Phytopathology 30: 315-348. Gillespie, D. R. 1989. Biological control of thrips (Thysanoptera: Thripidae) on greenhouse cucumber by Amblyseius cucumeris. Entomophaga 34: 185-192. Gillespie, D. R., and C. A. Ramey. 1988. Life history and cold storage of Amblyseius cucumeris (Acarina: Phytoseiidae). Journal of Entomological Society of British Columbia 85: 71-76. Glockemann, B. 1992. Biological control of Frankliniella occidentalis on ornamental plants using predatory mites. EPPO Bulletin 22: 397-404.

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GraphPad 2007. GraphPad Prism, GraphPhad Software Inc. computer program, version 5.0. By GraphPad. Hassan, S. A., F. Bigler, H. Boggenschutz, E. Boller, J. Brun, P. Chiverton, P. Edwards, F. Mansour, E. Naton, P. A. Oomen, W. P. J. Overmeer, L. Polgar, W. Rieckmann, L. Samsoe-Petersen, A. Staubli, G. Sterk, K. Tavares, J. J. Tuset, G. Viggiani, and A. G. Vivas. 1988. Results of the fourth joint pesticide testing programme carried out by the IOBC/WPRS-working group "Pesticide and Beneficial Organisms". Journal of Applied Entomology 105: 321-329. Hassan, S. A., R. Albert, F. Bigler, P. Blaisinger, H. Boggenschutz, E. Boller, J. Brun, P. Chiverton, P. Edwards, W. D. Englert, P. Huang, C. Inglesfield, E. Naton, P. A. Oomen, W. P. J. Overmeer, W. Rieckmann, L. Samsoe-Petersen, A. Staubli, J. J. Tuset, G. Viggiani, and G. Vanwetswinkel. 1987. Results of the third joint pesticide testing programme by the IOBC/WPRS-working group "Pesticides and Beneficial Organisms. Journal of Applied Entomology 103: 92-107. Herron, G. A., and D. F. Cook. 2002. Initial verification of the resistance management strategy for Frankliniella occidentalis (Pergande) (Thysanoptera: Thripidae) in Australia. Australian Journal of Entomology 41: 187-191. Herron, G. A., and T. M. James. 2005. Monitoring insecticide resistance in Australian Frankliniella occidentalis Pergande (Thysanoptera: Thripidae) detects fipronil and spinosad resistance. Australian Journal of Entomology 44: 299-303. Holt, K. M., G. P. Opit, J. R. Nechols, and D. C. Margolies. 2006. Testing for non-target effects of spinosad on twos-potted spider mites and their predator Phytoseiulus persimilis under greenhouse conditions. Experimental and Applied Acarology 38: 141- 149. Hunter, W. B., and D. E. Ullman. 1989. Analysis of mouthpart movements during feeding of Frankliniella occidentalis (Pergande) and Frankliniella schultzei Tryborn (Thysanoptera: Thripidae). International Journal of Insect Morphology and Embryology 18: 161-172. Jensen, S. E. 1998. Acetyl cholinesterase activity associated with methiocarb resistance in a strain of western flower thrips, Frankliniella occidentalis (Pergande). Pesticide Biochemistry and Physiology 61: 191-200. Jensen, S. E. 2000. Insecticide resistance in the western flower thrips, Frankliniella occidentalis. Integrated Pest Management Reviews 5: 131-146. Jones, T., C. Scott-Dupree, R. Harris, and B. Harris. 2002. Spinosad: an effective biocide for inclusion in integrated pest management programs for Frankliniella occidentalis Pergande (Thysanoptera: Thripidae) on greenhouse cucumbers. IOBC/WPRS Bulletin 25: 119-122. Jones, T., C. Scott-Dupree, R. Harris, L. Shipp, and B. Harris. 2005. The efficacy of spinosad against the western flower thrips, Frankliniella occidentalis, and its impact on associated biological control agents on greenhouse cucumbers in southern Ontario. Pest Management Science 61: 179-185. Kareiva, P., and R. Sahakian. 1990. Tritrophic effects of a simple architectural mutation in pea plants. Nature 345: 433-434. Khan, I., and J. G. Morse. 2006. Impact of citrus thrips chemical treatments on the predatory mite Euseius tularensis. Journal of Applied Entomology 130: 386-392. Kim, S. S., and C. H. Paik. 1996. Comparative toxicity of fenpyroximate to the predatory mite, Amblyseius womersleyi Schicha and the Kanzawa spider mite, Tetranychus kanzawai Kishida (Acarina: Phytoseiidae, Tetranychidae). Applied Entomology and Zoology 31: 369-377. Kim, S. S., and S. G. Seo. 2001. Relative toxicity of some acaricides to the predatory mite, Amblyseius womersleyi and the twos-potted spider mite, Tetranychus urticae (Acari: Phytoseiidae, Tetranychidae). Applied Entomology and Zoology 36: 509-514. Kirk, W. D. J., and L. I. Terry. 2003. The spread of the western flower thrips Frankliniella occidentalis (Pergande). Agricultural and Forest Entomology 5: 301-310.

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Kongchuensin, M., and A. Takafuji. 2006. Effects of Some Pesticides on the Predatory Mite, Neoseiulus longispinosus (Evans) (Gamasina: Phytoseiidae). Journal of Acarology Society of Japan 15: 17-27. Lewis, T. 1998. Pest thrips in perspective, pp. 385-390, Proceedings The 1998 Brighton Conference- Pest and Diseases. British Crop Protection Council, Brighton, UK. Li, D.-X., J. Tian, and Z.-R. Shen. 2006. Effects of pesticides on the functional response of predatory thrips, Scolothrips takahashii to Tetranychus viennensis. Journal of Applied Entomology 130: 314-322. Ludwig, S. W. 2002. Impact of spinosad on Orius insidiosus populations on greenhouse Marigolds, pp. 3, First floriculture industry research and scholarship trust. Texas A & M Agricultural Research and Extension Centre Overton, Tx and Kelli Hoover, Department of Entomology, The Pennsylvania State University, Pennsylvania. Ludwig, S. W., and R. D. Oetting. 2001. Effect of spinosad on Orius insidiosus (Hemiptera: Anthocoridae) when used for Frankliniella occidentalis (Thysanoptera: Thripidae) control on greenhouse pot chrysanthemums. Florida Entomologist 84: 311-313. Malipatil, M. B., A. C. Postle, J. A. Osmelak, M. Hill, and J. Moran. 1993. First record of Frankliniella occidentalis (Pergande) (Thysanoptera: Thripidae) in Australia. Australian Journal of Entomology 32: 378. McMurtry, J. A., and B. A. Croft. 1997. Life-styles of phytoseiid mites and their roles in biological control. Annual Review of Entomology 42: 291-321. Medhurst, A., and M. Y. Steiner. 2001. Western Flower Thrips and Strawberries. National Strategy for the Management of WFT & TSWV, East Melbourne, Victoria, Australia. Messelink, G. J., S. E. F. van Steenpaal, and P. M. J. Ramakers. 2006. Evaluation of phytoseiid predators for control of western flower thrips on greenhouse cucumber. Biocontrol 51: 753-768. Nagai, K. 1990. Effects of a juvenile hormone mimic material, 4-phenoxyphenyl (RS)-2-(2- pyridyloxy) propyl ether, on Thrips palmi Karni (Thysanoptera: Thripidae) and its predator Orius sp (Hemiptera: Anthocoridae). Applied Entomology and Zoology 25: 199-204. Pietrantonia, P. V., and J. H. Benedict. 1999. Effect of new cotton insecticide chemistries, tebufenozide, spinosad and chlorfenapyr, on Orius insidiosus and two Cotesia species. Southwestern Entomologist 24: 21-29. Pietrantonio, P. V., and J. H. Benedict. 1999. Effect of new cotton insecticide chemistries, tebufenozide, spinosad and chlorfenapyr on Orius insidiosus and two Cotesia species. Southwestern Entomologist 24: 21-29. Premachandra, D. W. T. S., C. Borgemeister, and H.-M. Poehling. 2005. Effects of Neem and Spinosad on Ceratothripoides claratris (Thysanoptera: Thripidae), an Important Vegetable Pest in Thailand, Under Laboratory and Greenhouse Conditions. Journal of Economic Entomology 98: 438-448. Price, P. W., C. E. Bouton, P. Gross, B. A. McPheron, J. N. Thompson, and A. E. Weis. 1980. Interactions among three trophic levels: influence of plants on interaction between insect herbivores and natural enemies. Annual Review of Ecology and Systematics 11: 41-65. Quinn, G. P., and M. J. Keough. 2002. Experimental design and data analysis for biologists. University Press, Cambridge. Reuveni, R. [ed.] 1995. Novel approaches to integrated pest management. Lewis Publishers, Boca Raton, Florida. Rhodes, E. M., and O. E. Liburd. 2006. Evaluation of predatory mites and acramite for control of two-spotted spider mites in strawberries in North Central Florida. Journal of Economic Entomology 99: 1291-1298. SAS 2002-2003. SAS 9.1 computer program, version 9.1. By SAS, Cary, NC, USA. Sparks, T. C., G. D. Thomson, H. A. Kirst, M. B. Hertlein, L. L. Larson, T. V. Worden, and S. T. Thibault. 1998. Biological activity of the spinosyn, new fermentation derived insect control agents, on tobacco budworm (Lepidoptera: Noctuidae) larvae. Journal of Economic Entomology 91: 1277-1283.

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Steiner, M., and S. Goodwin. 2000. Development and evaluation of usage protocols for newly developed western flower thrips biocontrol agents, pp. 15-25. In D. Cook [ed.], National strategy for the management of western flower thrips and tomato spotted wilt virus. Department of Agriculture Western Australia, South Perth. Steiner, M. Y., and S. Goodwin. 1998. Development and marketing of an IPM package for western flower thrips, Frankliniella occidentalis (Pergande). Phase II: Evaluation and producing the natural enemies. HRDC/HSNA Report. NSW Agriculture, Gosford, Australia. Steiner, M. Y., and S. Goodwin. 2001. Development and marketing of an IPM package for western flower thrips, Frankliniella occidentalis. Phase III: Development and evaluation of usage protocols for newly developed western flower thrips biocontrol agents. HRDC report. NSW Agriculture, Gosford, Australia. Steiner, M. Y., S. Goodwin, T. M. Wellham, I. M. Barchia, and L. J. Spohr. 2003. Biological studies of the Australian predatory mite Typhlodromips montdorensis (Schicha) (Acari: Phytoseiidae), a potential biocontrol agent for western flower thrips, Frankliniella occidentalis (Pergande) (Thysanoptera: Thripidae). Australian Journal of Entomology 42: 124-130. Thoeming, G., and H.-M. Poehling. 2006. Integrating soil-applied azadiractin with Amblyseius cucumeris (Acari: Phytoseiidae) and Hypoaspis miles (Acari: Laelapidae) for the management of Frankliniella occidentalis (Thysanoptera: Thripidae). Environmental Entomology 35: 746-756. Ullio, L. 2002. Australia's national strategy for the management of western flower thrips (WFT), Frankliniella occidentalis (Pergande), pp. 687-689. In T. Hietaranta, M.-M. Linna, P. Palonen and P. Parikka [eds.], Proceedings of the fourth International Strawberry Symposium. Acta Horticulturae, MTT Agrifood Research, Finland. Ullman, D. E., J. L. Sherwood, and T. L. German. 1997. Thrips as vectors of plant pathogens, pp. 539-565. In T. Lewis [ed.], Thrips as Crop Pests. CAB International, Wallingford, U. K. Ullman, D. E., D. M. Westcot, W. B. Hunter, and R. F. L. Mau. 1989. Internal anatomy and morphology of Frankliniella occidentalis (Pergande) (Thysanoptera: Thripidae) with special reference to interactions between thrips and tomato spotted wilt virus. International Journal of Insect Morphology and Embryology 18: 289-310. van Dijken, F. R., M. T. A. Dik, B. Gebala, J. de Jong, and C. Mollema. 1994. Western flower thrips (Thysanoptera: Thripidae) effect on chrysanthemum cultivars: plants growth and leaf scarring in nonflowering plants. Journal of Economic Entomology 87: 1312-1317. van Driesche, R. G., S. Lyon, and C. Nunn. 2006. Compatibility of spinosad with predacious mites (Acari: Phytoseiidae) used to control western flower thrips (Thysanoptera: Thripidae) in greenhouse crops. Florida Entomologist 89: 396-401. van Houten, Y. M., P. van Stratum, J. Bruin, and A. Veerman. 1995. Selection for non- diapause in Amblyseius cucumeris and Amblyseius barkeri and exploration of the effectiveness of selected strains for thrips control. Entomologia Experimentalis et Applicata 77: 289-295. van Lenteren, J. C., and J. Woets. 1988. Biological and Integrated Pest control in Greenhouses. Annual Review of Entomology 33: 239-269. Villanueva, R. T., and J. F. Walgenbach. 2005a. Development, oviposition, and mortality of Neoseiulus fallacis (Acari: Phytoseiidae) in response to reduced-risk Insecticides. Journal of Economic Entomology 98: 2114-2120. Villanueva, R. T., and J. F. Walgenbach. 2005b. Development, Oviposition, and Mortality of Neoseiulus fallacis (Acari: Phytoseiidae) in Response to Reduced-Risk Insecticides. Journal of Economic Entomology 98: 2114-2120. Wiethoff, J., H.-M. Poehling, and R. Meyhöfer. 2004. Combining plant- and soil-dwelling predatory mites to optimise biological control of thrips. Experimental and Applied Acarology 34: 239-261.

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Wijkamp, I., R. Goldbach, and D. Peters. 1996. Propagation of tomato spotted wilt virus in Frankliniella occidentalis does neither result in pathological effects nor in transovarial passage of the virus. Entomologia Experimentalis et Applicata 81: 285-292. Williams, T., J. Valle, and E. Vinuela. 2003. Is the naturally derived insecticide Spinosad compatible with insect natural enemies? Biocontrol Science and Technology 13: 459- 475. Zalom, F. G., P. A. Phillips, N. C. Toscano, and M. Bolda. 2001. Strawberry, Western flower thrips. UC IPM Pest Management Guidelines: Strawberry. UC ANR Publication 3468, Insects and Mites. Davis, California, USA. http://www.ipm.ucdavis.r734301211.html. Zar, J. H. 1999. Biostatistical Analysis. Prentice Hall International, Upper Saddle River, New Jersey. Zilahl-Balogh, G. M. G., J. L. Shipp, C. Cloutier, and J. Brodeur. 2007. Predation by Neoseiulus cucumeris on western flower thrips, and its oviposition on greenhouse cucumber under winter vs. summer conditions in a temperate climate. Biological Control 40: 160-167.

89 CHAPTER IV

Single versus multiple releases of predatory mites (Acari) combined with a spinosad application for the management of western flower thrips, Frankliniella occidentalis (Pergande) (Thysanoptera: Thripidae) in strawberry, Fragaria x ananassa Duchesne (Rosaceae)

Keywords: Frankliniella occidentalis, Typhlodromips montdorensis, Neoseiulus cucumeris, Hypoaspis miles, spinosad, single species releases, multiple species release, strawberry

Abstract

Western flower thrips, Frankliniella occidentalis (Pergande) (Thysanoptera: Thripidae) is a major pest of strawberry in Australia. Three predatory mites, Typhlodromips montdorensis (Schicha) (Phytoseiidae), Neoseiulus cucumeris (Oudemans) (Phytoseiidae) and Hypoaspis miles (Berlese) (Laelapidae) are commercially available in Australia for thrips management. This study sought to determine the effect of single species versus multiple species release of these predatory mites in combination with an insecticide on F. occidentalis. Spinosad is currently registered in Australia and is efficacious against thrips and regarded to be compatible in an IPM program. In the glasshouse, strawberry plants (cv Camino Real) were sprayed once with either spinosad at the recommended rate (80 mL/100 L rate, 0.096 g a.i./L) or water (control). Frankliniella occidentalis adults were released onto plants 24 h after spraying, and mites were released six days later. Mites were released as single-species, two-species, or three- species combinations. Spinosad significantly reduced F. occidentalis numbers compared to the control (water). Typhlodromips montdorensis, N. cucumeris and H. miles significantly reduced F. occidentalis numbers compared to the control (no mites). Spinosad had no effect on mites, as their numbers (T. montdorensis and N. cucumeris) were higher at the end of the trial than when initially released. Numbers of Typhlodromips montdorensis and N. cucumeris did not differ between spinosad and water-treated plants. As H. miles is a soil-dwelling mite, their numbers could not be counted. Mites released in combination with spinosad were more effective at reducing thrips numbers than individual applications of predatory mites alone: ‘T. montdorensis and H. miles’ was the most effective combination. The effectiveness against F. occidentalis was not different between releases of ‘T. montdorensis and H. miles’ and ‘T. montdorensis, N. cucumeris and H. miles’. When T. montdorensis and N. cucumeris were released as single species separately, there was no significant difference in their numbers, though T. montdorensis numbers were relatively higher than N. cucumeris. When released as a double-species combination, there were significantly more T. montdorensis than N. cucumeris. In the triple- Chapter IV: Effects of cultivar and predatory mites species combination, T. montdorensis and N. cucumeris numbers were not significantly different. The results suggest that spinosad followed by releases of either ‘T. montdorensis and H. miles’ or ‘T. montdorensis, N. cucumeris and H. miles’ can reduce thrips number effectively.

4. 1 Introduction

Western flower thrips (WFT), Frankliniella occidentalis (Pergande) (Thysanoptera: Thripidae) is an important economic pest worldwide (Brødsgaard and Albajes 1999, Jones et al. 2002, Kirk and Terry 2003), causing extensive crop losses (Lewis 1998). With its piercing-sucking mandibles, WFT penetrates epidermal and subepidermal cells causing extensive damage: this includes deformation and growth reduction of the plant, and silver scars on fruits and leaves (van Dijken et al. 1994, de Jager et al. 1995). In addition to direct damage, WFT also causes indirect damage by transmitting tospoviruses (Wijkamp et al. 1996, Ullman et al. 1997). Since its detection in 1993 in Western Australia (Malipatil et al. 1993), WFT has spread to each state and territory, except the Northern Territory and become a major pest in several crops including strawberry. Strawberry [Fragaria x ananassa Duchesne (Rosaceae)] is an intensively managed crop cultivated for its fresh, aromatic, red berries. In Australia, the strawberry industry has grown steadily over the last few years with a gross value of approximately $AUD308 million per annum (Anonymous 2009). However, strawberry production is often hampered by direct damage caused by WFT in low tunnel, open fields, and glasshouses (Ullio 2002).

Whilst insecticides are the main control method (Herron and Cook 2002), because of its small size, secretive habit, high reproductive potential, and ability to develop resistance to insecticide, WFT can be difficult to control (Jensen 2000). WFT has also developed resistance to several major classes of chemicals (Brødsgaard 1994, Jensen 1998, Jensen 2000) throughout the world.

As an integrated approach, there is a growing trend to use two or more species of natural enemies to suppress insect pest populations (Premachandra et al. 2003, Avilla et al. 2004, Blümel 2004, Brødsgaard 2004, Chau and Heinz 2004, Chow and Heinz 2004, Hoddle 2004, Shipp and Ramakers 2004, Thoeming and Poehling 2006, Chow et al. 2008). Overseas, phytoseiid mites are used to manage WFT in field and glasshouse crops (Chant 1985, van Lenteren and Woets 1988, McMurtry and Croft 1997). For example, anthocorid bugs of the genus Orius and phytoseiid mites of the genus Amblyseius are commonly used for the control of WFT in glasshouse-grown crops in Europe and North America (Brødsgaard 2004, Shipp and Ramakers 2004), but resulting benefits have not quantitatively validated (Blockmans and Tetteroo 2002, Skirvin et al. 2006). Some of the studies support the premise of biological control agent compatibility (Gillespie and Quiring 1992, Wittmann and Leather 1997,

91 Chapter IV: Effects of cultivar and predatory mites

Brødsgaard and Enkegaard 2005), while, others oppose this view (Magalhăes et al. 2004, Sanderson et al. 2005). Schausberger and Walzer (2001) demonstrated that interspecific competition may occur when different species of predatory mites are combined together and prey specificity can influence the quality and intensity of predator-predator interactions. Schausberger and Walzer (2001) reported that in perennial, glasshouse-grown crops, the release of Phytoseiulus persimilis Athias-Henriot and Neoseiulus californicus (McGrgor) (Acari: Phytoseiidae) released to control carmine spider mite, Tetranychus cinnabarinus (Tetranychidae), could have complementary effects. However, the augmentative release of predatory mites in single or multiple releases is not always sufficient to manage WFT in crops with the low economic threshold (Gillespie and Ramey 1988, Bakker and Sabelis 1989, Gillespie 1989), particularly when low damage is required.

Until recently, Australian growers had no effective biological control options for WFT due to quarantine restrictions prohibiting their importation (Steiner and Goodwin 2001). Recently four predatory mite species have become available to Australian growers, two of which are natives: Typhlodromips montdorensis (Schicha) and Typhlodromus occidentalis (Schicha) (Phytoseiidae) (Steiner and Goodwin 2000). Neoseiulus cucumeris (Oudemans) (Phytoseiidae) and Hypoaspis miles (Berlese) (Laelapidae), native to New Zealand, were recently confirmed as occurring in Australia, despite no record of deliberate introduction. Typhlodromips montdorensis, N. cucumeris and H. miles are commercially available and have been used for WFT management (Anonymous 2006).

The compatibility of these predatory mites has not been tested, nor has their effect in conjunction with insecticides currently used for WFT management in Australia been evaluated. Spinosad™ (Dow AgroSciences, USA), a mixture of tetracyclic-macrolide compounds, has been classified as a reduced-risk bio-insecticide (Sparks et al. 1998) and is the primary insecticide used in Australia for WFT control. However, impact of spinosad on predators is varied (Williams et al. 2003, Cote et al. 2004, Jones et al. 2005, Villanueva and Walgenbach 2005, van Driesche et al. 2006). Thus, the objectives of this study were to (i) investigate the effectiveness of commercially available predatory mites [T. montdorensis, N. cucumeris and H. miles] with or without spinosad against WFT, and (ii) determine the effectiveness of single versus combined release of predatory mites for the management of WFT.

4.2 Materials and methods

The experiment was conducted in a glasshouse (25 ± 2⁰C, 60-70% RH, 16: 8 L: D cycle) at the University of Western Australia (UWA) from November 2007 to January 2008.

92 Chapter IV: Effects of cultivar and predatory mites

4.2.1 Source cultures

4.2.1.1 Strawberry cultivar

Strawberry [Fragaria ananassa Duchesne (Rosaceae)] cv Camino Real (short-day length cultivar) was used in this study. In previous experiments (Chapters 2 and 3), Camino Real was found to be less preferred to WFT and was used in this experiment. Strawberry runners were obtained from a commercial grower in June 2006, and propagated in pots (32.5l x 32.5w x 40.5h cm) containing potting mix (Baileys Fertilisers, Rockingham, WA) in glasshouses at the Department of Agriculture and Food Western Australia (DAFWA) and UWA. All potted plants were covered with a modified thrips cage (45 x 35 cm) made from thrips-proof mesh net (105µ, Sefar Filter Specialists Pty Ltd., Malaga, WA; see Chapter 3) and supported by quadrate steel-rod stands. All pots were fitted with sprinklers. The plants were watered every third day. A liquid fertiliser (Thrive®, Yates, Australia; NPK: 12.4: 3: 6.2; rate: 5mL/2 L water) was applied once a month.

4.2.1.2 Western flower thrips (WFT)

WFT were initially collected from calendula, Calendula officinalis L. (Asteraceae) at DAFWA, South Perth and reared on calendula in pots (50x100 mm). Pots were kept in thrips proof Perspex cages (500 x 420 x 400 mm H x D x W), fitted with 105 µ mesh net. The cage fitted on top of a Nylex tote box (Blyth Enterprises Ptd ltd, Australia; 320 x 420 mm). Cages were kept in a glasshouse and tunnel houses at UWA from July 2006 to November 2008. Plants were watered as described above. Every second week, adults were collected from caged plants using an aspirator and released onto new potted calendula plants to ensure the continuous availability of WFT during study periods.

To obtain uniformly aged WFT, 20 adults were collected from the colony, released onto fresh caged plants, and allowed to lay eggs for 24 h. After 24 h, the adults were removed with a small aspirator. The plants were checked daily for larvae emergence. Newly hatched larvae were removed and released onto a strawberry leaf on a moistened filter paper in a Petri dish (150 x 15 mm). The leaf petiole was covered with cotton, soaked in a 10% sugar solution to extend the life of the leaf. The top of the Petri dish was covered with thrips-proof mesh (105 µ) and the edges of the Petri dish and mesh were sealed with paraffin film (Parafilm M®, Micro Analytix Pty Ltd)] and kept in a controlled temperature room (25±1⁰C, 50-60% RH, 16:8 h L: D regime). Larvae hatched on the same day were transferred to a new Petri dish as above and allowed to pupate. Adults that emerged on the same day were used in trials.

93 Chapter IV: Effects of cultivar and predatory mites

4.2.1.3 Predatory mites

Predatory mites [T. montdorensis, N. cucumeris and H. miles] used in the study were sourced from commercial suppliers (Biological Services, SA; Chilman IPM Services, WA; and Beneficial Bug Company, NSW). Mites were provided in plastic buckets containing vermiculite. Trials were conducted immediately upon receipt of mites.

4.2.2 Experiment: effect of single versus multiple species releases of mites combined with spinosad on WFT

To determine the effectiveness of predatory mites with or without spinosad, an experiment with a split-plot design was conducted in a glasshouse at UWA. One hundred and sixty potted strawberry plants, 2-3 weeks old with 2-3 leaves (excess leaves were pruned), were divided into two groups (80 per group), and sprayed with either spinosad at the recommended rate (80 mL/100 L rate, 0.096 g a.i./L) or water with a hand-held atomiser (Hills Sprayers, BH220063) until run-off (after van Driesche et al. (2006)). Plants were then covered with a modified thrips cage (45 x 35 cm, open both ends) made from thrips-proof mesh (105 µ), supported by quadrate steel-rod stands. The bottom end of the cage was taped to the pot. The top end of the cage was closed with a rubber band.

Fifteen previously collected 15 WFT adults (2 d old) were released onto each plant 24 h after spraying. Each group of 80 plants (spinosad or untreated control) were further divided into eight treatments (mite release) with 10 plants per treatment. All possible combinations (single and multiple species) of mite were included:

(i) no mites (ii) T. montdorensis (iii) N. cucumeris (iv) H. miles (v) T. montdorensis and N. cucumeris (vi) T. montdorensis and H. miles (vii) N. cucumeris and H. miles (viii) T. montdorensis, N. cucumeris and H. miles.

Predatory mites were released six days after spraying (Khan and Morse 2006). The numbers of predatory mites released per plant were (i) six (ii) three + three and (iii) two

94 Chapter IV: Effects of cultivar and predatory mites

+two + two for single, double and triple species combinations respectively. WFT (adults and larvae) were counted every third day for three weeks (from the release of WFT to the end of the third week). Thrips were counted between 0600 to 0800 h, when they were less active. Each plant was checked with a battery-powered magnifying glass [50mm (2") illuminated round 2x power with 4x bifocal magnifier].

To assess the numbers of predatory mites, at the end of the trial, plants were removed from pots and preserved in a container with 80% ethyl alcohol. The plant was washed onto a double-layer sieve (made from 105µ mesh) and checked under a stereomicroscope, and the numbers of T. montdorensis and N. cucumeris per plant were recorded. To determine the numbers of H. miles (soil-dwelling), plant parts and top soil (2-3 cm) from pots [H. miles released pots] were collected and preserved as above. The plant materials and soils were washed onto a double-layer sieve as above and checked under a stereomicroscope. However, no H. miles were recovered.

The trial pots were fitted with sprinklers (watering every third day) in such a way that water did not reach the leaf and upper parts of the plant to avoid washing out WFT or mites. As an extra precaution, the watering program was set for the afternoon (1900 h).

4.2.3 Data analysis

The effectiveness of mite treatments (single-, double- or triple species-releases) with or without spinosad on WFT numbers over time, were analysed with repeated measures ANOVAs (Proc Mixed Procedure) with split-plot design. WFT adults and larvae were separately analysed (independent fixed variables: mites treatments, spray treatment and time; random variable: plant numbers; response variables: adults and larvae). Because there was a significant three-way interaction between the main effects of spray, predatory mites and time (days), additional ANOVAs (repeated measures) were performed for each spray treatment (Quinn and Keough 2002). Due to the number of multiple tests, an adjustment to the significance level was made [α = 0.025 (0.05/2)]. If significant differences among means were detected, the means were separated using least square means with the adjusted significance level (SAS 2002-2003).

The difference in predatory mite numbers for single species releases (T. montdorensis and N. cucumeris) and treatment (spinosad, water) was analysed with two-way ANOVA (Proc Mixed Procedure; independent fixed variables: spray, mite species; response variable: numbers of mites). Similarly, two-way ANOVAs were used to determine the

95 Chapter IV: Effects of cultivar and predatory mites difference in numbers of T. montdorensis and N. cucumeris in double-species (T. montdorensis and N. cucumeris) and triple-species (T. montdorensis, N. cucumeris and H. miles) combinations. A two-way ANOVA was used to determine the difference of T. montdorensis and N. cucumeris when released in double-species combination with H. miles (‘T. montdorensis and H. miles’ and ‘N. cucumeris and H. miles’). Two separate two-way ANOVAs were used to determine the difference in numbers of T. montdorensis in double species-releases (combinations: ‘T. montdorensis and N. cucumeris’ and ‘T. montdorensis and H. miles). Similarly, two separate two-way ANOVAs were used to evaluate N. cucumeris numbers in double species releases (combinations: T. montdorensis and N. cucumeris’ and ‘N. cucumeris and H. miles).

Data were subjected to square root transformations when appropriate, to meet the assumption of homogeneity of variances (Zar 1999) before the data were subjected to statistical analysis. Data were reverse transformed for presentation in figures. All statistical analyses were computed using the SAS 9.1 Statistical Package (SAS 2002- 2003), while figures were constructed using Graphpad Prism 5.0 (GraphPad Software Inc 2007).

4.3 Results

4.3.1 Western flower thrips

4.3.1.1 Adults

Across all mite combination treatments, spinosad-treated plants had significantly fewer

WFT adults than water-treated plants (Figure 4.1, F 7,144 = 14.06, P < 0.0001; Appendix

4.1). Since there was a significant interaction (F 35, 720 = 2.52, P < 0.0001) between spray, predatory mite treatments and time (days), further repeated measures ANOVAs were carried out separately for each spray treatment (Appendix 4.1).

96 Chapter IV: Effects of cultivar and predatory mites

Spinosad Water 40 SE)  30

20

10

Number of WFT adults (Mean adults WFT of Number 0 No mites Tm Nc Hm Tm+Nc Tm+HmNc+Hm Tm+Nc+Hm

Figure 4.1 Comparison of mean number of WFT adults per plant sprayed with either spinosad or water and in the presence of no mites or different mite combinations. Within each group, means were significantly different (α = 0.05). Tm = T. montdorensis, Nc = N. cucumeris, Hm = H. miles.

When plants were sprayed with spinosad, the numbers of WFT on plants with different combinations of predators varied over time (F 35, 360 = 23.94, P < 0.0001) (Appendix 4.1, Figure 4.2A). Mites appeared to take some time to establish. Six and nine days after WFT release, the numbers of WFT adults per plant were not different between mite treatments (Figure 4.2A). From days 12 to 21, the mean numbers of WFT adults were lowest on plants that received the ‘T. montdorensis and H. miles’ two-species combination only. From days 15 to 21, the numbers of WFT adults did not differ between plants with ‘T. montdorensis and H. miles’ and ‘T. montdorensis, N. cucumeris and H. miles’ combinations. From days 12 to 21, the numbers of WFT adults were highest on plants that received no mites.

When plants were sprayed with water only (control), the number of WFT adults on plants with different combinations of predatory mites varied significantly over time (F

35, 360 = 37.21, P < 0.0001) (Appendix 4.1, Figure 4.2B). As with the spinosad treatment, it took mites a few days to establish on plants. On day six, the mean numbers of WFT adults were not different among the mite treatments. Differences in numbers of WFT adults between mite treatments began to appear on day nine. From days 9 to 21, plants with the two-species combinations of T. montdorensis and H. miles had the lowest numbers of WFT adults, similar to the spinosad treatment. Plants that did not receive any mites had the highest numbers of WFT.

97 Chapter IV: Effects of cultivar and predatory mites

No mites Tm Nc Hm Tm+Nc Tm+Hm Nc+Hm Tm+Nc+Hm 60 A

SE) 40 

20

0 60 B

40

20 Numberof WFTadults (Mean

0 6D 9D 12D 15D 18D 21D

Figure 4.2 Effects of predatory mites on mean number of WFT adults per plant sprayed with (A) spinosad or (B) water. X-axis represents days after initial WFT release. Tm = T. montdorensis, Nc = N. cucumeris, Hm = H. miles.

40 Spinosad Water SE)  30

20

10

Number of WFT larvae (Mean larvae WFT of Number 0 No mites Tm Nc Hm Tm+Nc Tm+HmNc+Hm Tm+Nc+Hm

Figure 4.3 Comparison of mean number of WFT larvae per plant sprayed with either spinosad or water and in the presence of no mites or different mite combinations. Within each group, means were significantly different (α = 0.05). Tm = T. montdorensis, Nc = N. cucumeris, Hm = H. miles.

4.3.1.2 Larvae

Similar to WFT adults, there were fewer WFT larvae (F 7, 144 = 46.62, P < 0.0001) on plants sprayed with spinosad compared to the water (control) Figure 4.3). There was a

98 Chapter IV: Effects of cultivar and predatory mites

significant three-way interaction (F 35, 720 = 2.52, P < 0.0001) between spray treatment, predatory mite treatment and time (days) that influenced the numbers of WFT larvae. Further ANOVAS for each spray (spinosad, water) were carried out (Appendix 4.1).

In plants sprayed with spinosad, the number of WFT larvae on plants with different combinations of predatory mites varied over time (Figure 4.4A; F 35, 360 = 6.64, P < 0.0001). On days six and nine, the least numbers of WFT larvae were found on plants that received ‘T. montdorensis and N. cucumeris’. From days 12 to 21, the two-species mite combination of ‘T. montdorensis and H. miles’ had the lowest numbers of WFT larvae. From days 15 to 21, WFT larvae numbers did not differ between plants treated with the two-species combination of ‘T. montdorensis and H. miles’ or the three-species combination. WFT larvae were generally highest on the plants that were not treated with mites, except on day six, when no difference was found between plants that were not treated with mites and plants treated with H. miles.

No mites Tm Nc Hm Tm+Nc Tm+Hm Nc+Hm Tm+Nc+Hm 50

A 40

30 SE)  20

10

0 50 B

40

Number of WFT larvae (Mean ofWFT larvae Number 30

20

10

0 6D 9D 12D 15D 18D 21D

Figure 4.4 Effects of predatory mites on the mean number of WFT larvae per plant sprayed with (A) spinosad or (B) water. X-axis represents days after initial WFT release. Tm = T. montdorensis, Nc = N. cucumeris, Hm = H. miles.

99 Chapter IV: Effects of cultivar and predatory mites

On plants sprayed with water, the number of WFT larvae on plants with different combinations of predatory mites tended to increase over time (Figure 4.4B; F 35, 360 = 5.21, P <0.0001). Mites, in any combination, reduced the numbers of WFT larvae per plant (Figure 4.4B). On day six, plants with ‘T. montdorensis, N. cucumeris and H. miles’ had the lowest numbers of WFT larvae but there was little difference in the numbers of thrips in any of the other mite treatments. The numbers of WFT larvae were low on plants with all three predatory mite species throughout the rest of the trial, though these did not differ between plants treated with ‘T. montdorensis and H. miles’ or ‘T. montdorensis and N. cucumeris’. By 21 days, plants that received ‘T. montdorensis and H. miles’ had the lowest numbers of WFT larvae. However, the number of WFT larvae did not differ between plants receiving ‘T. montdorensis and H. miles’ or ‘T. montdorensis, N. cucumeris and H. miles’.

4.3.2 Predatory mites

When T. montdorensis and N. cucumeris were applied singly, there was no significant interaction between the spray treatment and mite species (F 1, 36 = 0.12, P = 0.73). Overall (spinosad and water), the mean numbers of T. montdorensis (20.18 ± 0.86) and

N. cucumeris (19.75 ± 0.88) did not differ (F 1, 36 = 3.65, P = 0.06). When released as a single-species, the overall mean numbers of predatory mites (T. montdorensis and N. cucumeris) between spinosad (19.55 ± 1.22) and water (20.38 ± 0.88) treatments were not different (F 1, 36 = 00.85, P = 0.36; Figure 4.5A). Similar to single-species releases, in double-species releases (T. montdorensis and N. cucumeris), there was no interaction between spray and mite treatments (F 1, 36 = 0.49, P = 0.49). However, there were significantly more T. montdorensis (10.50 ± 0.49) than N. cucumeris (8.55 ± 0.55) (F (1,

36) = 15.29, P = 0.0004; Figure 4.5B). Overall, mean numbers of predatory mites (T. montdorensis and N. cucumeris) per plant between spinosad (9.10 ± 0.55 mites/plant) and water (9.95 ± 11.35 mites/plant) treatments did not differ (F 1, 36 = 2.95, P = 0.10). When either T. montdorensis or N. cucumeris were released with H. miles, there was no significant interaction of spray and mite treatments on mite numbers per plant (F 1, 36 =

0.01, P = 0.10). There was also no significant difference (F 1, 36 = 0.24, P = 0.63 Figure 4.5C) between the mean numbers of T. montdorensis (12.10 ± 0.91) and N. cucumeris (12.35 ± 0.52). Similarly, overall mean numbers of predatory mites (T. montdorensis and N. cucumeris) did not differ (F 1, 36 = 0.77, P = 0.39) between spinosad (12.00 ± 11.51 mites/plant) and water (12.45 ± 1.32 mites/plant) treatments. For the three-species combination (T. montdorensis, N. cucumeris and H. miles), there was no interaction (F 1,

36 = 0.43, P = 0.52) of spray and mite treatments that affected predatory mite numbers

100 Chapter IV: Effects of cultivar and predatory mites per plant. Though more T. montdorensis were found per plant (8.35 ± 1.42) than N. cucumeris (7.50 ± 1.22), the difference (F 1, 36 = 1.22, P = 0.198) was not significant (Figure 4.5). Similarly, overall mean numbers of predatory mites did not differ between spinosad (7.95 ± 0.99) and water (8.95 ± 1.31) treatments.

25 A B

SE) 20  15 b a 10 5 0 25 C D 20 15 10 5 Number of predatory mites per plant (Mean plant per mites predatory of Number 0 Tm Nc Tm Nc

Figure 4.5 Comparison of mean number of T. montdorensis and N. cucumeris per plant applied with (A) single-species releases of T. montdorensis and N. cucumeris, (B) double-species releases of T. montdorensis and N. cucumeris, (C) double species releases of T. montdorensis and H. miles and N. cucumeris and H. miles, and (D) triple- species release. Means with different letters differed significantly (α = 0.05). Tm = T. montdorensis, N. cucumeris, Hm = H. miles.

When T. montdorensis were released with N. cucumeris and H. miles in two separate combinations, spray and mite combination had no significant influence (F 1 , 36 = 0.73, P = 0.398) on T. montdorensis numbers per plant. It also appeared that spray had no significant (F 1, 36= 2.92, P = 0.096) influence on the overall numbers of T. montdorensis per plant (10.90 ± 0.46 and 11.72 ± 0.55 for spinosad and water treatment respectively). The mean numbers of T. montdorensis per plant were higher when combined with H. miles than N. cucumeris (F 1, 36= 11.73, P = 0.002; Figure 4.6). When N. cucumeris was released with T. montdorensis and H. miles in double-species combinations, spray and mites combination had no significant interaction (F 1, 36 = 0.34, P = 0.566). The mean numbers of N. cucumeris per plant were higher when combined with H. miles than T. montdorensis and the difference (F 1, 36 = 49.51, P < 0.0001) was significant (Figure 4.6). The overall mean numbers of N. cucumeris did not differ (F 1, 36

101 Chapter IV: Effects of cultivar and predatory mites

= 0.86, P = 0.361) between spinosad (10.19 ± 0.59) and water (10.70 ± 0.48) treatments.

T. montdorensis 15 b a 10 SE)  5

0 Tm+Nc Tm+Hm Spinosad Water N. cucumeris 15 b

10 a

5 Number of predatory mites (Mean mites predatory of Number

0 Tm+Nc Nc+Hm Spinosad Water

Figure 4.6 Mean numbers of (A) T. montdorensis and (B) N. cucumeris per plant in double-species combinations. Tm = T. montdorensis, Nc = N. cucumeris, Hm = H. miles.

4.4 Discussion

The results of the present study suggest that mites can reduce the WFT population. However, single mite species may not be able to keep WFT populations under the economic threshold level. According to Steiner and Goodwin (2005), 45% of flowers with five or more WFT adults is the economic threshold in strawberry. In this study the plants were kept flowerless therefore it is not possible to make a direct assessment of the impact of predatory mites on lowering the thrips below an economic threshold. Nevertheless, the most effective treatment combination in terms of lowering the thrips population was an application of spinosad followed by multiple predatory mite releases (T. montdorensis, N. cucumeris and H. miles). The two-species combination of T. montdorensis and H. miles, or the three-species combination provided better management of WFT than single-species releases or other two-species combinations. Predatory mite releases combined with a spinosad application also provided a higher suppression of WFT adults and larvae than either spinosad or mites alone. This confirms the findings of the previous study (Chapter 3), that integration of T. montdorensis, N. cucumeris or H. miles with spinosad is more successful in reducing WFT in glasshouse-grown strawberry than applying either spinosad or releasing predatory mites alone.

102 Chapter IV: Effects of cultivar and predatory mites

Present study demonstrates that either of the predatory mites and their combination performed better against WFT when released after spinosad spray. While, in water treatment, either mite species was able to reduce WFT population to some extent, which might not be enough to reduce WFT population effectively. Previous studies have also shown that WFT can be controlled with an insecticide application followed by the release of beneficials (Ludwig and Oetting 2001, Ludwig 2002, Thoeming and Poehling 2006). Thoeming and Poehling (2006) reported that an application of neem (Botanical, 17% azadiractin) combined with the release of a combination of two predatory mite species increased efficacy to 99%, without causing any significant harm to predatory mites. The previous findings of Ludwing and Oetting (2001) and Ludwig (2002) suggest that applications of spinosad and the predatory bug, Orius insidiosus Say (Hemiptera: Anthocoridae) to glasshouse- potted chrysanthemums and marigold significantly reduced WFT compared to the control (without spinosad or Orius). Ludwing and Oetting (2001), Ludwig (2002) and Funderburk et al. (2000) demonstrate that spinosad had no or little effect on O. insidiosus. However, the laboratory studies of Elzen et al. (1998) and Pietrantonia and Benedict (1999) showed that spinosad has low toxicity to O. insidiosus. Similarly, Kongchuensin and Takafuji (2006) reported that fresh spinosad residues (up to 48 h old) have a significant, negative effect on eggs and the immature stage of the predatory mite Neoseiulus longispinosus (Evans) (Acari: Phytoseiidae). However, spinosad was not harmful to N. longispinosus seven days after application. This study and the previous study (Chapter 3) suggest that spinosad poses no detrimental effect to T. montdorensis, N. cucumeris and H. miles if these mites are released six days after spraying.

Multiple-species release of predatory mites can be more effective in reducing WFT populations than single-species releases. However, release of multiple species of predator may result in intraguild predation (IGP) which can have a significant effect on the prey suppression (Losey and Denno 1998). Although IGP is common in many predators (Vance-Chalcraft et al. 2007) including several predatory mites (Schausberger and Walzer 2001, Walzer and Schausberger 2005), it is not known whether any intraguild predation can occur among T. montdorensis, N. cucumeris and H. miles. Brodeur et al. (2002) recognised that the release of multiple predator species is an effective strategy that would suppress pest populations in a manner that is more economically viable than the use of single predator species. Wiethoff et al. (2004) reported that cucumber plants had fewer WFT when N. cucumeris and H. miles were applied together than either N. cucumeris or H miles alone. Similarly, combined releases of Phytoseiulus persimilis (Athias-Henriot) and Neoseiulus californicus (McGregor) (Acari: Phytoseiidae) can provide long-term suppression of carmine spider mite (Tetranychus cinnabarinus Boisduval (Tetranychidae)) in potted gerbera plants, compared to individual-species releases (Schausberger and Walzer 2001). Premachandra et al. (2005) reported that the emergence of

103 Chapter IV: Effects of cultivar and predatory mites

WFT adults was significantly lower if entomopathogens were released with the predatory mite, Hypoaspis aculeifer (Berlese) (Acari: Laelapidae), than applications of either entomopathogens or H. aculeifer alone. Premachandra et al. (2005) also showed that when foliage-inhabiting (T. montdorensis) and soil-dwelling (H. miles) mite species were applied together, they provided the highest suppression of WFT. Synergistic effects can be expected if a plant-dwelling predator can evoke the escape behaviour of the prey, making the prey available for ground-dwelling predators (Losey and Denno 1999).

In the present study, two foliage foraging predatory mites combined together provided better results in suppressing WFT than their single-species release. However, this combination was less effective than when combined with release of the soil-dwelling H. miles. This can be explained by the resource competition between two predators, which occurs if two predators compete for a shared prey (Janssen et al. 1998). As a result, one species might outcompete the other. When T. montdorensis and N. cucumeris are released together, there were more T. montdorensis than N. cucumeris, which is not the case when they are combined with H. miles separately. When all three species were released, no interaction seemed to occur between T. montdorensis and N. cucumeris. One possibility is that as in the two-species combination, plants had higher numbers of T. montdorensis and N. cucumeris, which might increase the interspecific competition. In the triple-species combinations, plants had less T. montdorensis and N. cucumeris, and perhaps less competition for WFT. Thus for the successful implementation of multiple releases of predatory mites in a pest management program, further studies need to be carried out to determine if cannibalism, intra-guild predation and preference between primary prey (pest population) and secondary prey (predator population) occurs, especially when two or more predators utilise the same resource. Furthermore, it is also important to determine the optimal release rate of predatory mites that can effectively reduce the pest population, without having any negative impact on each other.

In conclusion, the integration of spinosad and multiple-species releases of predatory mites could be a sustainable management strategy for WFT management in strawberry. When single-species release is the only option, T. montdorensis in conjunction with spinosad appears to be the best combination for the management of WFT in strawberry. WFT management could be further improved by reducing the interval between spray application and predatory mite release. However, the residual toxicity of spinosad to predatory mites would first need to be determined.

4.5 Literature cited

Anonymous. 2006. Typhlodromus occidentalis. Biological Services, Loxton, Australia.

104 Chapter IV: Effects of cultivar and predatory mites

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Gillespie, D. R., and D. J. M. Quiring. 1992. Competition between Orius tristicolor (White) (Hemiptera: Anthocoridae) and Amblyseius cucumeris (Oudemans) (Acari: Phytoseiidae) on Frankliniella occidentalis (Pergande) (Thysanoptera: Thripidae). The Canadian Entomologist 124: 1123-1128. GraphPad Software Inc 2007. GraphPad Prism computer program, version 5.0. By GraphPad Software Inc. Herron, G. A., and D. F. Cook. 2002. Initial verification of the resistance management strategy for Frankliniella occidentalis (Pergande) (Thysanoptera: Thripidae) in Australia. Australian Journal of Entomology 41: 187-191. Hoddle, M. S. 2004. Biological control of whiteflies on ornamental crops, pp. 149-170. In K. M. Heinz, R. G. van Driesche and M. P. Parrella [eds.], Biocontrol in protected culture. Ball Publishing, Batavia, Illinois. Janssen, A., A. Palini, M. Venzon, and M. W. Sabelis. 1998. Behaviour and indirect interactions in food webs of plant-inhabiting arthropods. Experimental and Applied Acarology 22: 497-521. Jensen, S. E. 1998. Acetylcholinesterase activity associated with methiocarb resistance in a strain of western flower thrips, Frankliniella occidentalis (Pergande). Pesticide Biochemistry and Physiology 61: 191-200. Jensen, S. E. 2000. Insecticide resistance in the western flower thrips, Frankliniella occidentalis. Integrated Pest Management Reviews 5: 131-146. Jones, T., C. Scott-Dupree, R. Harris, and B. Harris. 2002. Spinosad: an effective biocide for inclusion in integrated pest management programs for Frankliniella occidentalis Pergande (Thysanoptera: Thripidae) on greenhouse cucumbers. IOBC/WPRS Bulletin 25: 119-122. Jones, T., C. Scott-Dupree, R. Harris, L. Shipp, and B. Harris. 2005. The efficacy of spinosad against the western flower thrips, Frankliniella occidentalis, and its impact on associated biological control agents on greenhouse cucumbers in southern Ontario. Pest Management Science 61: 179-185. Khan, I., and J. G. Morse. 2006. Impact of citrus thrips chemical treatments on the predatory mite Euseius tularensis. Journal of Applied Entomology 130: 386-392. Kirk, W. D. J., and L. I. Terry. 2003. The spread of the western flower thrips Frankliniella occidentalis (Pergande). Agricultural and Forest Entomology 5: 301-310. Kongchuensin, M., and A. Takafuji. 2006. Effects of Some Pesticides on the Predatory Mite, Neoseiulus longispinosus (Evans) (Gamasina: Phytoseiidae). Journal of Acarology Society of Japan 15: 17-27. Lewis, T. 1998. Pest thrips in perspective, pp. 385-390, Proceedings The 1998 Brighton Conference- Pest and Diseases. British Crop Protection Council, Brighton, UK. Losey, J. E., and R. F. Denno. 1998. Positive predator-predator interactions: enhanced predation rates and synergistic suppression of aphid populations. Ecology 79: 2143- 2152. Losey, J. E., and R. F. Denno. 1999. Factors facilitating synergistic predation: the central role of synchrony. Ecological Application 9: 378-386. Ludwig, S. W. 2002. Impact of spinosad on Orius insidiosus populations on greenhouse Marigolds, pp. 3, First floriculture industry research and scholarship trust. Texas A & M Agricultural Research and Extension Centre Overton, Tx and Kelli Hoover, Department of Entomology, The Pennsylvania State University, Pennsylvania. Ludwig, S. W., and R. D. Oetting. 2001. Effect of spinosad on Orius insidiosus (Hemiptera: Anthocoridae) when used for Frankliniella occidentalis (Thysanoptera: Thripidae) control on greenhouse pot chrysanthemums. Florida Entomologist 84: 311-313. Magalhăes, S., C. Tudorache, M. Montserrat, R. van Maanen, M. W. Sabelis, and A. Janssen. 2004. Diet of intraguild predators affects antipredator behaviour in intraguild prey. Behavioral Ecology 16: 364-370. Malipatil, M. B., A. C. Postle, J. A. Osmelak, M. Hill, and J. Moran. 1993. First record of Frankliniella occidentalis (Pergande) (Thysanoptera: Thripidae) in Australia. Australian Journal of Entomology 32: 378.

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McMurtry, J. A., and B. A. Croft. 1997. Life-styles of phytoseiid mites and their roles in biological control. Annual Review of Entomology 42: 291-321. Pietrantonia, P. V., and J. H. Benedict. 1999. Effect of new cotton insecticide chemistries, tebufenozide, spinosad and chlorfenapyr, on Orius insidiosus and two Cotesia species. Southwestern Entomologist 24: 21-29. Premachandra, D. W. T. S., C. Borgemeister, and H.-M. Poehling. 2005. Effects of Neem and Spinosad on Ceratothripoides claratris (Thysanoptera: Thripidae), an Important Vegetable Pest in Thailand, Under Laboratory and Greenhouse Conditions. Journal of Economic Entomology 98: 438-448. Premachandra, W. T. S. D., C. Borgemeister, O. Berndt, and R.-U. Ehilers. 2003. Combined release of entomopathogenic nematodes and the predatory mite Hypoaspis aculeifer to control soil-dwelling stages of western flower thrips Frankliniella. Biocontrol 48: 529-541. Quinn, G. P., and M. J. Keough. 2002. Experimental design and data analysis for biologists. University Press, Cambridge. Sanderson, J. P., H. F. Brødsgaard, and A. Enkegaard. 2005. Preference assessment of two Orius spp for Neoseiulus cucumeris vs Frankliniella occidentalis, pp. 221-224, I.B.O.C./W.P.R.S. Bulletin. SAS 2002-2003. SAS 9.1 computer program, version 9.1. By SAS, Cary, NC, USA. Schausberger, P., and A. Walzer. 2001. Combined versus Single Species Release of Predaceous Mites: Predator-Predator Interactions and Pest Suppression. Biological Control 20: 269-278. Shipp, J. L., and P. M. J. Ramakers. 2004. Biological control thrips on vegetable crops, pp. 265-276. In K. M. Heinz, R. G. van Driesche and M. P. Parrella [eds.], Biocontrol in protected culture. Ball Publishing, Batavia, Illinois. Skirvin, D. J., L. Kravar-Garde, K. Reynolds, J. Jones, J. J. Reynolds, and M. E. de Courcy Williams. 2006. The influence of pollen on combining predators to control Frankliniella occidentalis in ornamental chrysanthemum crops. Biocontrol Science and Technology 16: 99-105. Sparks, T. C., G. D. Thomson, H. A. Kirst, M. B. Hertlein, L. L. Larson, T. V. Worden, and S. T. Thibault. 1998. Biological activity of the spinosyns, new fermentation derived insect control agents, on tobacco budworm (Lepidoptera: Noctuidae) larvae. Journal of Economic Entomology 91: 1277-1283. Steiner, M., and S. Goodwin. 2000. Development and evaluation of usage protocols for newly developed western flower thrips biocontrol agents, pp. 15-25. In D. Cook [ed.], National strategy for the management of western flower thrips and tomato spotted wilt virus. Department of Agriculture Western Australia, South Perth. Steiner, M. Y., and S. Goodwin. 2001. Development and marketing of an IPM package for western flower thrips, Frankliniella occidentalis. Phase III: Development and evaluation of usage protocols for newly developed western flower thrips biocontrol agents. HRDC report. NSW Agriculture, Gosford, Australia. Steiner, M. Y., and S. Goodwin. 2005. Management of thrips (Thysanoptera: Thripidae) in Australian strawberry crops: within-plant distribution characteristics and action thresholds. Australian Journal of Entomology 44: 175-185. Thoeming, G., and H.-M. Poehling. 2006. Integrating soil-applied azadiractin with Amblyseius cucumeris (Acari: Phytoseiidae) and Hypoaspis miles (Acari: Laelapidae) for the management of Frankliniella occidentalis (Thysanoptera: Thripidae). Environmental Entomology 35: 746-756. Ullio, L. 2002. Australia's national strategy for the management of western flower thrips (WFT), Frankliniella occidentalis (Pergande), pp. 687-689. In T. Hietaranta, M.-M. Linna, P. Palonen and P. Parikka [eds.], Proceedings of the fourth International Strawberry Symposium. Acta Horticulturae, MTT Agrifood Research, Finland. Ullman, D. E., J. L. Sherwood, and T. L. German. 1997. Thrips as vectors of plant pathogens, pp. 539-565. In T. Lewis [ed.], Thrips as Crop Pests. CAB International, Wallingford, U. K.

107 Chapter IV: Effects of cultivar and predatory mites van Dijken, F. R., M. T. A. Dik, B. Gebala, J. de Jong, and C. Mollema. 1994. Western flower thrips (Thysanoptera: Thripidae) effect on chrysanthemum cultivars: plants growth and leaf scarring in nonflowering plants. Journal of Economic Entomology 87: 1312-1317. van Driesche, R. G., S. Lyon, and C. Nunn. 2006. Compatibility of spinosad with predacious mites (Acari: Phytoseiidae) used to control western flower thrips (Thysanoptera: Thripidae) in greenhouse crops. Florida Entomologist 89: 396-401. van Lenteren, J. C., and J. Woets. 1988. Biological and Integrated Pest control in Greenhouses. Annual Review of Entomology 33: 239-269. Vance-Chalcraft, H. D., J. A. Rosenheim, J. R. Vonesh, and C. W. Osenberg. 2007. The influence of intraguld predaation on prey suppression and prey release: A meta-analysis. Ecology 88: 2689-2696. Villanueva, R. T., and J. F. Walgenbach. 2005. Development, oviposition, and mortality of Neoseiulus fallacis (Acari: Phytoseiidae) in response to reduced-risk Iisecticides. Journal of Economic Entomology 98: 2114-2120. Walzer, A., and P. Schausberger. 2005. Are two better than one? Combined effects of the predatory mites Phytoseiulus persimilis and Neoseiulus californicus (Acari: Phytoseiidae) on spider mite control. IOBC/WPRS Bulletin 28: 309-312. Wiethoff, J., H.-M. Poehling, and R. Meyhöfer. 2004. Combining plant- and soil-dwelling predatory mites to optimise biological control of thrips. Experimental and Applied Acarology 34: 239-261. Wijkamp, I., R. Goldbach, and D. Peters. 1996. Propagation of tomato spotted wilt virus in Frankliniella occidentalis does neither result in pathological effects nor in transovarial passage of the virus. Entomologia Experimentalis et Applicata 81: 285-292. Williams, T., J. Valle, and E. Vinuela. 2003. Is the naturally derived insecticide Spinosad compatible with insect natural enemies? Biocontrol Science and Technology 13: 459- 475. Wittmann, E. J., and S. R. Leather. 1997. Compatibility of Orius laevigatus Fieber (Hemiptera: Anthocoridae) with Neoseiulus (Amblyseius) cucumeris Oudemans (Acari: Phytoseiidae) and Iphiseius (Amblyseius) degenerans Berlese (Acari: Phytoseiidae) in the biocontrol of Frankliniella occidentalis Pergande (Thysanoptera: Thripidae). Experimental and Applied Acarology 21: 523-538. Zar, J. H. 1999. Biostatistical Analysis. Prentice Hall International, Upper Saddle River, New Jersey.

108 CHAPTER V

Use of spinosad and predatory mites (Acari) for the management of western flower thrips, Frankliniella occidentalis (Pergande) (Thysanoptera: Thripidae) in strawberry [Fragaria x ananassa Duchesne (Rosaceae)]: a field study

Key words: Frankliniella occidentalis, Typhlodromips montdorensis, Neoseiulus cucumeris, Hypoaspis miles, spinosad, strawberry, low tunnel, integration, multiple release

Abstract

The efficacy of single- and multiple- species releases of predatory mites (Acari), Typhlodromips montdorensis Schicha (Phytoseiidae), Neoseiulus cucumeris Oudemans (Phytoseiidae) and Hypoaspis miles Berlese (Laelapidae) and their compatibility with spinosad (Success™, Dow AgroSciences, Australia) for the control of western flower thrips, Frankliniella occidentalis (Pergande) (Thysanoptera: Thripidae) was evaluated in commercial strawberry. The trial was carried out in a commercial strawberry [Fragaria x ananassa Duchesne (Rosaceae)] farm at Bullsbrook, Western Australia, from September to November 2007 (spring). Naturally occurring F. occidentalis infestations on low tunnel-grown strawberry were sprayed with water (control), ‘spinosad (80 mL/100 L rate, 0.096 g a.i./L) then mites’ or ‘mites then spinosad (80 mL/100 L rate, 0.096 g a.i./L)’ applications. Predatory mites (Acari), Typhlodromips montdorensis Schicha (Phytoseiidae), Neoseiulus cucumeris Oudemans (Phytoseiidae) and Hypoaspis miles Berlese (Laelapidae) were released as single-, two-, and three- species combinations. Predatory mites reduced the number of F. occidentalis on strawberry plants sprayed with either water or spinosad, compared to the no mite treatment. Frankliniella occidentalis numbers were lower on spinosad-treated plants that received predatory mites than on the plants sprayed with water and received predatory mites. Spinosad posed no negative effect to predatory mites, as mite numbers on plants sprayed with spinosad did not differ from the water treated plants. However, predatory mites were most effective in reducing thrips when released after spinosad was applied (‘spinosad then mite’ treatment). The three species combination of predatory mites appeared to perform better in reducing thrips numbers compared to their individual release. The two species combination of T. montdorensis (foliage inhabiting) and H. miles (soil dwelling) appeared to be the most effective in suppressing F. occidentalis. The next most effective combination was a triple-species release (T. montdorensis, N. cucumeris and H. miles). The double-species combination of T. montdorensis and N. cucumeris was the least effective, and may interact with each other. Although spinosad and predatory mites were able to reduce thrips numbers, F. occidentalis numbers increased five weeks after treatment. Chapter V: Compatibility of predatory mites and spinosad

This suggests that a further application of predatory mites, spinosad or both is required. Single- and multiple-species release of predatory mites combined with spinosad for F. occidentalis management in low tunnel-grown strawberry are discussed.

5.1 Introduction

Western flower thrips (WFT), Frankliniella occidentalis (Pergande) (Thysanoptera: Thripidae), is considered one of the most devastating pest thrips in a range of horticultural crops, including strawberry (Lewis 1973, Mound 1997). In Australia, the strawberry [Fragaria ananassa Duchesne (Rosaceae)] industry is worth AUD$308 million annually (Anonymous 2009), and is an intensively managed crop cultivated for its fresh, aromatic, red berries. However, strawberry production is often hampered by direct damage caused by WFT (Ullio 2002). WFT damage contributes to the ‘seediness’ of strawberry fruit (Medhurst and Steiner 2001), and is responsible for uneven ripening and yield loss (Houlding and Woods 1995). WFT feeding on strawberry blossoms may also cause stigmas and anthers to turn brown and wither prematurely (Zalom et al. 2001), and reduce flower receptacle size (Coll et al. 2007).

Because of its cryptic behaviour, minute size, and high reproductive rate, WFT is difficult to control using pesticides. Moreover, it has developed resistance against major classes of insecticides that are currently in use in many parts of the world (Helyer and Brobyn 1992, Brødsgaard 1994), including Australia (Herron and James 2005). Because of the inadequacy of chemical control, there is a need to develop integrated control methods. One particular challenge is to integrate biological and chemical control, as many pesticides have lethal and sublethal effects on biological control agents. Spinosad is a novel pesticide, derived from the fermentation of the actinomycete Saccharopolyspora spinosa Mertz and Yao (Sparks et al. 1998), and is classified as an environmentally and toxicologically reduced-risk chemical (Sparks et al. 1998, Cleveland et al. 2002, Thompson et al. 2002). Since its discovery, spinosad has been used in over 180 different crops to control a wide range of pests worldwide (Bret et al. 1997, Thompson et al. 1997, Zhao et al. 2002). In Australia, spinosad has been used for the control of Lepidopteran and Thysanopteran pests (Downard 2001), and is highly effective against WFT (Funderburk et al. 2000). Spinosad is regarded to have no or reduced toxicity to natural enemies (Brunner et al. 2001, Elzen 2001, Villanueva and Walgenbach 2005a). However, the selectivity of spinosad on predatory insects is under review (Pietrantonio and Benedict 1999, Williams et al. 2003). Spinosad is regarded to have low to moderate toxicity to predatory mites and the toxicity can vary from species to species (Williams et al. 2003, Cote et al. 2004, Jones et al. 2005). van Driesche et al. (2006) reported that fresh residues of spinosad applied at the recommended rate to control WFT on glasshouse flower

110 Chapter V: Compatibility of predatory mites and spinosad crops, had no toxic effect on the predatory mite Neoseiulus cucumeris (Oudemans) (Acari: Phytoseiidae), but lowered the survival of Iphiseius degenerans (Berlese) (Acari: Phytoseiidae). Spinosad is reported to be harmless to Phytoseiulus persimilis Athias-Henriot (Acari: Phytoseiidae), widely used for the control of two-spotted spider mites (Holt et al. 2006), but is highly toxic to Neoseiulus fallacis (Garman) (Acari: Phytoseiidae), which is used in North Carolina apple orchards to control European red mite (Panonychus ulmi Koch) and two-spotted mite (Tetranychus urticae Koch) (Villanueva and Walgenbach 2005b). Despite its detrimental effect on some species, spinosad can be integrated with biological control agents for WFT management (Funderburk et al. 2000), if a period of time is maintained between pesticide application and release of natural enemies (Jones et al. 2005, Khan and Morse 2006).

Several species of natural enemies are reported to attack above- and below-ground stages of WFT (van Driesche et al. 1998). Several predatory mite species (Acari) prey on either the larval or pupal stages, and are currently used to control WFT in protected crops (e.g. glasshouses) with some success (Riduavets 1995, Sabelis and Van Rijn 1997). Most attention has focused on three species: Amblyeius barkeri (Hughes), N. cucumeris and I. degenerans (Macgill 1927, Rodrìguez-Reina et al. 1992). The efficacy of N. cucumeris is limited because the adults only feed on thrips first instar larvae (Gillespie and Ramey 1988); it is the most commonly used biocontrol agent for thrips in protected cropping. Neoseiulus cucumeris is particularly successful in glasshouse capsicum (Ramakers 1988) in the Netherlands, but to my knowledge, little is known about the use of phytoseiid predators in semi-open or open fields.

Recently, several strawberry growers in Western Australia have begun releasing P. persimilis, a predatory mite for the control of the two-spotted mite, T. urticae, in the glasshouse and field, and N. cucumeris for control of WFT in glasshouses. The use of predatory mites in semiprotected crops such as low tunnels (floating covers) is limited. In Western Australia, field populations of WFT are low during winter (June-August), and increase during late September as temperature increases (L. Chilman, pers. comm. 2006). Therefore, the release of predatory mites before the spring population increase might be a useful approach for managing WFT in low tunnel-grown strawberry. Four species of predatory mite are commercially available in Australia (Neoseiulus cucumeris, Typhlodromips montdorensis, Hypoaspis miles and Hypoaspis aculeifer) (Biological Services 2009). Neoseiulus cucumeris and T. montdorensis are plant- dwelling species that predate on first instar WFT larva (Steiner and Goodwin 2002). Hypoaspis miles is a soil-dwelling species that predates on WFT pupae (Glockemann 1992). However, recent studies suggest that H. miles also predates on second instar WFT larva (Berndt 2002, Berndt 2003). The difference in their use on different parts of the plant, and predation on different WFT life stages raises the question of their compatibility in single versus combined

111 Chapter V: Compatibility of predatory mites and spinosad releases. The overall objective of the present study was to evaluate the efficacy of T. montdorensis, N. cucumeris, and H. miles as single-species and multiple-species releases, and their compatibility with spinosad for WFT control in low tunnel-grown strawberry. Specifically, this study had three main aims: i. Determine if spinosad and mites can be used effectively in combination to reduce WFT. It is expected that the combined application of T. montdorensis, N. cucumeris, and H. miles with spinosad would provide better suppression of WFT. ii. Determine if spinosad affects predatory mites. iii. Determine if a single-species release is more, or less effective than multiple-species releases in reducing WFT numbers. A combination of plant and soil dwelling mites could provide better control of WFT than plant dwelling mite alone.

5.2 Materials and methods

5.2.1 Study site

The trial was carried out on a commercial strawberry farm at Bullsbrook (S 31°39.294’, E 115º58.589), 54 km north of Perth, Western Australia, from 20 September to 26 November 2007. Strawberry (cv. Camarosa) was grown on silver plastic mulch under floating row covers. The grower made available four tunnels for experimentation. Each tunnel was 1.5 m wide and 50 m long, with a total of 668 (4 x 167) strawberry plants per tunnel. Strawberry runners were transplanted on 28 April 2007, with 30 cm intervals between runners within and between rows. After transplanting, the tunnels were covered with clear plastic sheets to keep the tunnel hot during the winter. Plants received no pesticide sprays three weeks before commencement of the experiment.

A data logger (Hobo Pro Series, HO8-031-08, Onset, USA) was installed inside each tunnel to record air temperature. Before installation, the data logger was programmed using Boxcar® Pro 4.3 software to log ambient air temperature at one-hour intervals during the experimental period. At the end of the experiment, temperature data was extracted from the data logger and averaged to obtain daily mean temperature (⁰C). Maximum, minimum and daily average air temperatures were collected from the Bureau of Meteorology, WA. The closest weather station was the RAAF Air Base, Pearce, approximately 22 km north-west of Bullsbrook (Figure 5.1).

112 Chapter V: Compatibility of predatory mites and spinosad

Max air temp Min air temp Ave temp Ave Temp inside tunnel 40

30 C) 

20 Temperature ( Temperature 10

0 5-Oct 4-Nov 9-Nov 10-Oct 15-Oct 20-Oct 25-Oct 30-Oct 30-Sep 14-Nov 19-Nov

Figure 5.1 Maximum, minimum and average daily air temperature (⁰C), and average daily temperature (⁰C) inside low tunnel (25 September to 20 November 2008). Maximum and minimum air temperature collected from RAAF, Pearce (22 km north-west of Bullsbrook). Temperature inside the tunnel was recorded using a data logger.

5.2.2 Predatory mites

Predatory mites [Typhlodromips montdorensis, Neoseiulus cucumeris and Hypoaspis miles] used in the study were sourced from commercial Australian suppliers (Biological Services, SA; Chilman IPM Services, WA; and the Beneficial Bug Company, NSW). Mites were provided in plastic buckets or plastic bag containing vermiculite. Trials were conducted immediately upon receipt of predatory mites.

5.2.3 Treatments

The treatments were applied in a two-factor, split-plot design (three spray treatments x eight mite treatments). Each of the four tunnels was divided into three plots, 1.2 m wide and 16.5 m long, with 220 (4 x 55) plants. Within each tunnel, spray treatments were randomly assigned to a plot and plants were sprayed with either water (control) or spinosad (Success™, Dow AgroSciences Australia) (Table 5.1). Spinosad treatments were applied either before the mites were released (‘spinosad then mites’), or after the mites were released (‘mites then spinosad’). The lapse of time between a spinosad application and predatory mite release was six days. Water or spinosad was applied with a Knapsack Sprayer (12 L, Rapid Spray™; Tank Management Ltd, Australia) until run-off. Because the experiment was carried out in low tunnel

113 Chapter V: Compatibility of predatory mites and spinosad where the environmental conditions somewhat similar to glasshouse, plants were sprayed until run-off). Spinosad (0.096 g a.i./L) was applied at the recommended rate of 80 mL/100 L.

Each spray plot within each tunnel was further divided into eight ‘mite release’ sub-plots, 1.2 m wide and 1.9 m long consisting of 24 (4 x 6) plants. Mite release treatments were:

(i) No mites (control) (ii) T. montdorensis (iii) N. cucumeris (iv) H. miles (v) T. montdorensis and N. cucumeris (vi) T. montdorensis and H. miles (vii) N. cucumeris and H. miles (viii) T. montdorensis, N. cucumeris and H. miles.

All mite treatments within each spray plot were randomly assigned. Within each tunnel, spray plots were separated by a row of plants. Similarly, a row of plants was kept as a buffer zone between each sub-plot, which was treated with neither spinosad nor mites. On either side of the buffer row, a mesh net (105 µ, Sefar Filter Specialists Pty Ltd., Malaga, WA) supported by a wooden frame created a barrier to prevent movement of WFT and predatory mites between sub- plots.

Table 5.1 Schedules of treatment applications and sampling.

Weeks Treatment 1-3 3 4 4-8 Water then Water Mites mites sprayed released Spinosad then Pre-treatment Spinosad Mites Post-treatment mites sampling sprayed released sampling Mites then Mites Spinosad spinosad released sprayed

5.2.3.1 Pre-treatment sampling

Before treatment application, plants in each plot (within each tunnel) were sampled for three weeks at weekly intervals to determine if there were any differences in WFT populations within or between tunnels. At each sample, three plants were randomly selected from a plot within each tunnel. Three flowers and three fruits were selected from each plant and removed with a pair of sharp scissors. Each flower or fruit was placed into a separate glass container (with 80%

114 Chapter V: Compatibility of predatory mites and spinosad ethyl alcohol) and labelled. In the laboratory, the numbers of WFT larvae and adults in each flower and fruit were counted under a binocular stereomicroscope. Across plots, the numbers of WFT larvae and adults were averaged per flower or fruit.

5.2.3.2 Post-treatment sampling

After the third sample was collected, the spray treatment areas were treated with water or spinosad, or sprinkled with predatory mites (Table 5.1). After six days and after the fourth sample had been collected, predatory mites were released onto the water- or spinosad-treated plants. Plots that had previously received predatory mites (week three) were sprayed with spinosad. Approximately 300 mites per m-2 were sprinkled over plants in the single-species treatment (de Courcy Williams 2001). In the two-species combination, the same release rate was used, but each species made up 50% of the total. Similarly, for the three-species combination, the above release rate was used, with each species comprising a third of the total.

Plots were sampled for four more weeks at weekly intervals as described above. In the laboratory, collected samples were checked under a binocular stereomicroscope and the numbers of WFT larvae and adults, T. montdorensis and N. cucumeris in each flower and fruit were recorded. Across plants, WFT larvae and adults, T. montdorensis and N. cucumeris numbers were averaged per flower or fruit. Since the H. miles count was not successful in previous experiments (chapters 3 and 4), no attempt was made to count H. miles.

5.2.4 Data analysis

To determine if counts of WFT differed within and between tunnels, before mite-release treatments, the numbers of WFT larvae and adults on the flowers and fruits were analysed with repeated measures ANOVA, with a split design (Proc Mixed Procedure). Independent fixed factors were sub-plot (within factor) and time (sampling week, repeated factor); random factor: plant number, block factor: tunnel (between); response variables: WFT adults and larvae. There were no significant differences in WFT adult and larval numbers within and between tunnels. The influence of spray application and predatory mite treatments on WFT adults and larvae over time from week four to eight (post-treatment) was subjected to repeated measures ANOVA with split-plot design (Proc Mixed Procedure). Independent fixed factors were spray treatment and mite treatments, time (repeated factor); random factors were plant numbers; block factor: tunnel; and response variables were WFT adults and larvae on flower and fruits. However, because there was a significant interaction of spray treatment, mite treatments and time, a series of repeated measures ANOVA with a split-plot design (Proc Mixed Procedure) was conducted

115 Chapter V: Compatibility of predatory mites and spinosad for each spray treatment (Quinn and Keough 2002). Since multiple comparisons were made, an adjustment to the significance level was required [α = 0.01667 (0.05/3]. If ANOVAs indicated a significant difference, means were subjected to pair wise comparison.

Because of the different release rates of predatory mites (T. montdorensis and N. cucumeris) in different mite combinations, several repeated measures ANOVAs with split-plot design (Proc Mixed Procedure) were used. Since mites were released in either week four (‘spinosad then mite’) or week three (‘mites then spinosad’ treatment), data collected from weeks five to eight were used to analyse differences in mite numbers. When released as single species, the effect of spray treatment and time were subjected to repeated measures ANOVA with a split-plot design [Proc Mixed Procedure; Fixed independent variable: spray, mite species, time (repeated factor); random factor: tunnel; response variables: flower, fruit]. Influence of spray treatment and time on predatory mites numbers (T. montdorensis and N. cucumeris) for double-species and triple- species releases were evaluated with a series of repeated measures ANOVA with a split-plot design [fixed independent variable: spray, mite species/combinations, times (repeated factor); random factor: tunnel; response variables: flower, fruit]. Additionally, the influence of spray treatment and species combination on T. montdorensis numbers in releases of T. montdorensis with the other mite species, and N. cucumeris with the other mite species, were analysed with repeated measures ANOVA with split-plot design [Proc Mixed Procedure; Fixed independent variable: spray, mite combinations, times (repeated factor); random factor: tunnel; response variables: T. montdorensis and N. cucumeris on flower and fruit]. If ANOVAs were significant, means were separated using least square means difference (α = 0.05).

The data were transformed with square root before analysis, to meet the assumption of homogeneity of variances (Zar 1999). However, actual means are presented in figures and tables. All statistical analyses were performed with SAS 9.1 Statistical Package (SAS 2002-2003). Figures were constructed using GraphPad Prism 5.0 (GraphPad, 2007).

5.3 Results

There were no pre-treatment differences in the mean number of WFT adults and larvae on flowers or fruit within and between tunnels (Appendix 5.1). An average of 7.37 ± 0.55 WFT adults were collected from flowers and 4.67 ± 0.67 from fruits. An average of 7.09 ± 0.34 WFT larvae was collected from flowers and 4.64 ± 0.22 from fruits. The number of WFT adults in flowers was above the economic threshold established for strawberry, which is 45% of flowers with five or more adult WFT (Steiner and Goodwin 2005).

116 Chapter V: Compatibility of predatory mites and spinosad

5.3.1 Impact of the spray and predatory mite species combinations on WFT adults

5.3.1.1 Flower

When applied in different combinations, predatory mites and spray treatments significantly affected (F14, 63 = 10.63, P < 0.0001) the number of WFT adults (Appendix 5.2, Figure 5.2). Across mite treatments, there were fewer WFT adults on flowers on ‘spinosad then mites’ treated plants and most on water-treated plants. However, there were significant interactions between spray treatment, predatory mite species combinations and time (F56, 288 = 18.93, P < 0.0001), and the effect of mite species combinations on WFT adults was evaluated with a series of ANOVAs by spray treatment (Appendix 5.2).

Water Spinosad then mites Mites then spinosad 10 SE)

 b c b 8 c b c b c a a 6 b b a a a a a a a a a b b a 4 a

2

0

Number of WFT flower per adult (Mean No mites Tm Nc Hm Tm*Nc Tm*Hm Nc*Hm Tm*Nc*Hm

Figure 5.2 Effect of spray treatment and predatory mite species releases on the number of WFT adults/flower in low tunnel strawberry. Tm = T. montdorensis, Nc = N. cucumeris, Hm = H. miles. Means with different letters within each group differed significantly (LS means, α = 0.05).*indicates mite species combinations.

The mean number of WFT adults on plants treated with water (F28, 96 = 276.74, P < 0.0001),

‘spinosad then mites’ (F 28, 96 = 86.57, P < 0.0001) or ‘mites then spinosad’ (F 28, 96 = 157.72, P < 0.0001), were affected by predatory mite species combinations (Figure 5.3, Appendix 5.2). The lowest number of WFT adults was recorded from plants where T. montdorensis and H. miles had been released in combination, or where all three mite species had been released (Figure 5.3). In the water and ‘spinosad then mites’ treatments, there was no significant difference in the number of WFT adults at week four (Figure 5.3). Across time, the number of WFT adults per flower was highest on plants that did not receive any mites (Figure 5.3).

117 Chapter V: Compatibility of predatory mites and spinosad

No mites Tm Nc Hm Tm*Nc Tm*Hm Nc*Hm Tm*Nc*Hm 12 A

10

8

6

4

2 SE)

 12 B 10

8

6

4

2 12 Number of WFT adults per flower (Mean flower per adults of WFT Number C 10

8

6

4

2 PoT1 PoT2 PoT3 PoT4 PoT5

Figure 5.3 Influence of predatory mite species combinations on WFT adults per flower over time (X-axis) in (A) water, (B) ‘spinosad then mites’ and (C) ‘mites then spinosad’. WFT adults’ counts were commenced at weekly interval. PoT = Post-spinosad spray/mites release. Within each week, means were separated by LS means (α = 0.017). Tm = T. montdorensis, Nc = N. cucumeris, Hm = H. miles. *indicates predatory mite species combination.

5.3.1.2 Fruit

Spray treatments and predatory mite species combinations affected the number of WFT adults per fruit (F14, 63 = 9.63, P < 0.0001; Figure 5.4, Appendix 5.2). WFT adult numbers were

118 Chapter V: Compatibility of predatory mites and spinosad highest on water-treated plants and on plants that did not receive any mites (> 4 WFT adults/fruit), and lowest, on ‘spinosad then mites’ treated plants (<3.75 WFT/fruit; Figure 5.4). There was a significant interaction between spray treatment, predatory mite species and time, (F

56, 288 = 3.78, P < 0.0001). The influence of mite species combinations on WFT adults/fruit over time was therefore evaluated by separate ANOVAs for each spray treatment (Appendix 5.2).

Water Spinosad then mites Mites then spinosad 8 SE) 

c 6 b b a c c c b c b

b a 4 b a b a a a b a a a a a 2 a

Number of WFT adults per fruit (Mean fruit per adults WFT of Number 0 No mites Tm Nc Hm Tm*Nc Tm*Hm Nc*Hm Tm*Nc*Hm

Figure 5.4 Effect of spray treatment and predatory mite species combinations (X-axis) on the number of WFT adults per fruit (Y-axis). Tm = T. montdorensis, Nc = N. cucumeris, Hm = H. miles. Means with different letters within each group differed significantly (LS means, α = 0.05). *indicates mite species combination.

Predatory mite species combinations and time affected the number of WFT adults per fruit in control (water) (F 28, 96 = 3.22, P < 0.0001), ‘spinosad then mite’ (F 28, 96 = 32.59, P < 0.0001) and ‘mites then spinosad’ (F 28, 96 = 47.34, P < 0.0001) treatments (Appendix 5.2, Figure 5.5). In all spray treatments, the lowest numbers of WFT adults per fruit were on plants where T. montdorensis and H. miles had been released in combination. However, for the water (control) treatment, WFT adults were lowest on plants with the triple-species combination, five weeks after treatment. There was no difference in the number of WFT adults /fruit on plants in water (control) and ‘mite then spinosad’ treatments on which ‘T. montdorensis and H. miles’ and ‘T. montdorensis, N. cucumeris and H. miles’ combinations were released. In addition, in the control (water) and ‘spinosad then mites’ treatments, WFT adults per fruit on week four (post- treatment sampling, PoT1) among mite treatments were not different. For all spray treatments across time, WFT adults per fruit were highest on plants with no mites.

119 Chapter V: Compatibility of predatory mites and spinosad

No mites Tm Nc Hm Tm*Nc Tm*Hm Nc*Hm Tm*Nc*Hm 7 A

5

3

1 SE)

 7 B

5

3

1

Number of WFT adults per fruit (Mean fruit per adults of WFT Number 7 C

5

3

1 PoT1 PoT2 PoT3 PoT4 PoT5

Figure 5.5 Influence of predatory mites on WFT adults per fruit over time (X-axis) in (A) water, (B) ‘spinosad then mites’ and (C) ‘mites then spinosad’. WFT adults’ counts were commenced at weekly interval. PoT = Post-spinosad spray/mites release. Tm = T. montdorensis, Nc = N. cucumeris, Hm = H. miles. Within each week, means were separated by LS means (α = 0.017). *indicates predatory mite species combination.

5.3.2 Impact of the spray and predatory mite species combinations on WFT larvae

5.3.2.1 Flower

Spray treatments and predatory mite release treatments affected the number of WFT larvae per flower (F14, 63 = 20.77, P < 0.0001; Figure 5.6). The lowest numbers of WFT larvae per flower

120 Chapter V: Compatibility of predatory mites and spinosad were on ‘spinosad then mites’-treated plants (3.72 ± 0.34 larvae/flower), and highest on water- treated plants (6.75 ± 0.27 larvae/flower). However, because there was a significant interaction of spray, mite release treatment and time (F56, 288 = 37.11, P < 0.0001), the effect of mite species combinations over time were evaluated with separate ANOVAs for each spray treatment (Appendix 5.3).

Water Spinosad then mites Mites then spinosad 10 SE)

 b

8 c c c c c a a c c 6 b

b b a b b a b b a 4 a a a a

2

0 Numberof WFT flower per larvae (Mean No mites Tm Nc Hm Tm*Nc Tm*Hm Nc*Hm Tm*Nc*Hm

Figure 5.6 Effect of spray treatment and predatory mite species combinations (X-axis) on the number of WFT larvae per flower (Y-axis). Tm = T. montdorensis, Nc = N. cucumeris, Hm = H. miles. Means with different letters within each group differed significantly (LS means, α = 0.05).*indicates mite species combination.

The highest number of WFT larvae was on plants with no mite releases (Figure 5.7). Predatory mite species combinations had a significant effect on the number of WFT larvae per flower (Appendix 5.3, Figure 5.7). For all spray treatments over the five weeks, the lowest numbers of WFT larvae were recorded from plants where T. montdorensis and H. miles had been released in combination (Figure 5.7). There were no differences in the number of WFT larvae on plants treated with either T. montdorensis or H. miles, or all three species, except at week five post- treatment.

121 Chapter V: Compatibility of predatory mites and spinosad

No mites Tm Nc Hm Tm*Nc Tm*Hm Nc*Hm Tm*Nc*Hm 12 A

10

8

6

4 SE)

 9 B 7

5

3

1 10 Number of WFT larvae (Mean flower per WFT of Number C

8

6

4

2 PoT1 PoT2 PoT3 PoT4 PoT5

Figure 5.7 Influence of predatory mites on WFT larvae per flower over time (X-axis) in (A) water, (B) ‘spinosad then mites’ and (C) ‘mites then spinosad’. WFT larvae counts commenced at weekly interval. PoT = Post-spinosad spray/mites release. Within each week, means were separated by LS means (α = 0.017). Tm = T. montdorensis, Nc = N. cucumeris, Hm = H. miles. *indicates predatory mite species combination.

5.3.2.2 Fruit

Predatory mite species combinations and spray treatment had a significant effect on the number of WFT larvae per fruit (F14, 63 = 25.43, P < 0.000; Appendix 5.3, Figure 5.8). The lowest number of larvae was on plants treated with ‘spinosad then mites’, and the highest on water- treated plants. There was a significant interaction of spray, predatory mite species combinations

122 Chapter V: Compatibility of predatory mites and spinosad

and time (week) (F56, 288 = 9.15, P < 0.0001). Separate ANOVAs were used to evaluate the effect of predatory mite species combinations on spray treatment (Appendix 5.3).

Water Spinosad then mites Mites then spinosad 8 SE) 

c 6 c b c c c c c b a 4 a b b a b b b b a a a a a a 2

0

Number of WFT larvae per fruit (Mean larvae fruit per of WFT Number No mites Tm Nc Hm Tm*Nc Tm*Hm Nc*Hm Tm*Nc*Hm

Figure 5.8 Effect of spray treatment and predatory mite species combinations (X-axis) on the number of WFT larvae per fruit. Tm = T. montdorensis, Nc = N. cucumeris, Hm = H. miles. Means with different letters within each group differed significantly (LS means, α = 0.05). *indicates mite species combination.

Any combination of predatory mite species appeared to have a significant effect on the number of larvae per fruit over the five weeks trial period, regardless of spray treatment (Appendix 5.3, Figure 5.9). The mean number of WFT larvae per fruit was lowest on plants receiving ‘T. montdorensis and H. miles’ and highest on plants with no mites. The mean number of WFT larvae per fruit did not differ across predatory mite treatments one week after treatment. There was no difference in mean number of larvae on plants that received either T. montdorensis or H. miles.

123 Chapter V: Compatibility of predatory mites and spinosad

No mites Tm Nc Hm Tm*Hm Nc*Hm Tm*Nc Tm*Nc*Hm 7 A

6

5

4

3 SE)

 6 B 5

4

3

2

1

Number of WFT larvae per plant (Mean plant per larvae WFT of Number 6 C

5

4

3

2

1 PoT1 PoT2 PoT3 PoT4 PoT5

Figure 5.9 Influence of predatory mites on WFT larvae per fruit over time (X-axis) in (A) control (water), (B) ‘spinosad then mites’ and (C) ‘mites then spinosad’. PoT = Post-spinosad spray/mites release. Within each week, means were separated by LS means (α = 0.017). Tm = T. montdorensis, Nc = N. cucumeris, Hm = H. miles. *indicates predatory mite species combination.

5.3.3 Impact of the spray and mite species combinations on predatory mites

5.3.3.1 Single species

There was no interaction of spray treatment and time (F 6, 63 = 0.97, P = 0.46; Appendix 5.4) on predatory mite numbers (T. montdorensis and N. cucumeris) per flower. The mean number of T. montdorensis and N. cucumeris per flower was significantly different (P = 0.26; Appendix 5.4,

124 Chapter V: Compatibility of predatory mites and spinosad

Figure 5.10). However, the mean number of predatory mites per flower differed significantly (F

2, 6 = 38.63, P = 0.0004) between spray treatments (Appendix 5.4, Figure 5.10). There was no difference in the number of predatory mites per flower in the control or ‘spinosad then mites’ treatments (Figure 5.10).

When mites were applied as single-species treatments, there was no significant interaction between spray treatment and time (P = 0.85; Appendix 5.4). There were significantly more T. montdorensis per fruit than N. cucumeris (F 1, 11 = 29.39, P = 0.0002; Figure 5.10). The overall mean numbers of predatory mites differed significantly amongst spray treatments (F 2, 6 = 29.97, P = 0.0008; Appendix 5.4, Figure 5.10). Overall, the highest numbers of predatory mites were found in the control (water) treatment and lowest on the ‘mites then spinosad’ treatment. However, there was no difference in the number of predatory mite numbers in the control and ‘spinosad then mites’ treatments.

6 Flower b b a 4 SE)  2

0 6 Fruit

b 4 b b a a

Number of predatory mites (Mean 2

0 Tm Nc Water S-M M-S Mite species Spray

Figure 5.10 Comparison of the mean number of T. montdorensis (Tm) and N. cucumeris (Nc) per flower or fruit, when applied as single species. Left side of the figure compares mean numbers of T. montdorensis and N. cucumeris. Right side compares the combined mean numbers of mites (T. montdorensis and N. cucumeris) between spray treatments. Means with different letters within group were significantly different (LS means α = 0.05). S-M = ‘Spinosad then mites’, M-S = ‘Mites then spinosad’.

125 Chapter V: Compatibility of predatory mites and spinosad

5.3.3.2 Two-species combinations

When T. montdorensis and N. cucumeris were applied in a double-species combination, spray treatment and time had no significant effect on predatory mite numbers (flower: F 6, 63 = 0.22, P

= 0.9680; fruit: F 6, 63 = 0.07, P = 0.9986; Appendix 5.4). However, there were significantly more T. montdorensis than N. cucumeris (flower: F 1, 11 = 10.53, P = 0.0092; fruit: F 1, 11 = 9.99,

P = 0.0081; Figure 5.11). The predatory mite numbers per flower (F 2, 6 = 11.95, P = 0.0081) or fruit (F 2, 6 = 13.95, P = 0.0062) differed between spray treatments (Figure 5.11). In both flowers and fruits, most predatory mites were found in the control treatment and least in ‘mites then spinosad’ treatment (Figure 5.11). The mean numbers of predatory mites in the control and ‘spinosad then mites’ treated plants did not differ.

For two-species combinations with H. miles, there was no interaction between spray treatment and time on predatory mite numbers (flowers: F 6, 63 = 0.50, P =0.81; fruit: F 6, 63 = 0.90, P = 0.50; Appendix 5.4). The mean number of predatory mites differed between spray treatments

(flower: F 2, 6 = 30.70, P =0.0007; fruit: F 2, 6 = 21.41, P = 0.0019; Figure 5.11), and were highest in the control and lowest in the ‘mites then spinosad’ treatment (Figure 5.11). The predatory mite numbers on control plants and ‘spinosad then mites’ treatments did not differ (Figure 5.11).

126 Chapter V: Compatibility of predatory mites and spinosad

4 Flower b b b A 3 a a

2

1

0 4 Fruit

3 b b b SE) a a  2

1

0 4 Flower b b B 3 a

2

Number of predatory mites (Mean 1

0 4 Fruit

3 b b a 2

1

0 Tm Nc Water S-M M-S Mite species Spray

Figure 5.11 Comparison of the mean numbers T. montdorensis (Tm) and N. cucumeris (Nc) applied in double-species combination as (A) ‘T. montdorensis and N. cucumeris’, and (B) T. montdorensis in ‘T. montdorensis and H. miles’ and N. cucumeris in ‘N. cucumeris and H. miles’. Left side of the figure compares mean numbers of T. montdorensis and N. cucumeris. Right side compares combined mean numbers of predatory mites (T. montdorensis and N. cucumeris) between spray treatments. Means with different letters within a group were significantly different (LS means, α = 0.05). S-M = ‘Spinosad then mites’, M-S = ‘Mites then spinosad’.

127 Chapter V: Compatibility of predatory mites and spinosad

5.3.3.3 Three-species release

Spray treatment and time had no significant effect on the overall mean numbers predatory mites

(T. montdorensis and N. cucumeris) (flower: F 6, 63 = 0.08, P = 0.9982; fruit: F 6, 63 = 0.09, P = 0.9974; Appendix 5.4). Similarly, the mean number of T. montdorensis and N. cucumeris were not different on flowers (F 1, 11 = 0.15, P = 0.7070) or fruits (F 1, 11 = 0.15, P =0.7078) (Figure 5.12). However, overall mean numbers of predatory mites differed significantly among spray treatments (flowers: F 2, 6 = 8.04, P = 0.0201; fruit: F 2, 6 = 8.03, P = 0.0201; Figure 5.12). The highest and lowest numbers of predatory mites were in the water and ‘mites then spinosad’ treatments respectively (Figure 5.12). However, the mean number of predatory mites per flower or fruit did not differ between control and ‘spinosad then mites’ treatments.

Flower 3 b b

2 a SE)  1

0 3 Fruit

b b 2 a

Number of predatory mites (Mean 1

0 Tm Nc Water S-M M-S Mite species Spray

Figure 5.12 Comparison of mean numbers of T. montdorensis (Tm) and N. cucumeris (Nc) when applied in a three-species combination (T. montdorensis, N. cucumeris and H. miles). Left side of the figure compares mean numbers of T. montdorensis and N. cucumeris per flower/fruit. Right side compares overall mean numbers of predatory mites between spray treatments. Means within each group with different letters were significantly different (LS means α = 0.05). S-M = ‘Spinosad then mites’, M-S = ‘Mites then spinosad’.

128 Chapter V: Compatibility of predatory mites and spinosad

5.3.3.4 Species interactions

When T. montdorensis was applied with either N. cucumeris or H. miles, mite species combinations and time had no effect on the number of T. montdorensis (flower: F 6, 54 = 0.60, P

= 0.73; fruit: F 6, 54 = 0.08, P = 0.10; Appendix 5.5). More T. montdorensis were found when applied with H. miles, and least when applied with N. cucumeris on flowers (F 1, 9 = 48.66, P <

0.0001) and fruit (F 1, 9 = 11.46, P = 0.0081) (Figure 5.13). When N. cucumeris was applied as a double-species combination with T. montdorensis or H. miles, mite species combinations and time had no significant effect on the mean numbers of N. cucumeris (flowers: F 6, 54 = 0.31, P =

0.93; fruit: F 6, 54 = 0.50, P = 0.81; Appendix 5.5). However, N. cucumeris numbers were highest on flowers (F 1, 9 = 20.43, P = 0.001) and on fruit (F 1, 9 = 14.42, P = 0.002), when applied with H. miles and lowest with T. montdorensis (Figure 5.13).

4 Fruit Flower b A a 3 b a 2 SE) 

1

0 Tm*Nc Tm*Hm Tm*Nc Tm*Hm 4 B Flower b Fruit 3 a b a Number of predatory mites (Mean 2

1

0 Tm*Nc Nc*Hm Tm*Nc Nc*Hm

Figure 5.13. Comparison of mean number of (A) T. montdorensis and (B) N. cucumeris when applied in double-species combination (X-axis). Tm = T. montdorensis, Nc = N. cucumeris, Hm = H. miles. Means within each group (flower/fruit) with different letters were significantly different (LS means, α = 0.05). *indicates mites species combinations.

129 Chapter V: Compatibility of predatory mites and spinosad

5.4 Discussion

The integration of reduced-risk chemicals and biological control for arthropod pest management may provide more comprehensive management than either approach alone. Unfortunately, insecticides are often found to be detrimental to natural enemies (Li et al. 2006, van Driesche et al. 2006). The negative impact of pesticides has been reported on several phytoseiid species used for WFT management (Hassan et al. 1987, Hassan et al. 1988, Kim and Paik 1996, Kim and Seo 2001, Amano et al. 2004, Li et al. 2006, van Driesche et al. 2006). Therefore, before recommending the integrated used of chemicals and biological control agents for commercial use, the effectiveness of the strategy needs to be evaluated. However, to my knowledge, there are limited numbers of studies that test the effectiveness of the combined use of chemical and biological control agents in Australia. In addition, no research has focused on the integration of insecticide and predatory mites for the management of WFT. The aim of this study was to investigate the compatibility of a reduced-risk chemical (spinosad) and predatory mites in single- versus multiple-species releases, and their effectiveness against WFT in low tunnel- grown strawberry in spring. As an integrated pest management approach, this study also evaluated whether predatory mites should be released before (‘mites then spinosad’) or after (‘spinosad then mites’) spinosad is applied.

The results indicate that T. montdorensis, N. cucumeris and H. miles can be used in low tunnel- grown strawberry for WFT management in spring. During the study period, the air temperature varied between 10-29⁰C (17⁰C ± 0.43), however, the temperature inside the tunnel was higher (range 15 to 35⁰C, 21.18⁰C ± 0.48) since it was heated by the sun. Gillespie and Ramey (1988) reported that N. cucumeris can survive at a constant temperature of 9⁰C for months, and oviposited within three days when returned to room temperature (20-22⁰C). Steiner et al. (2003) found that T. montdorensis did not diapause when reared under 10⁰C under long day conditions, and was able to survive at 8⁰C (Steiner and Goodwin 2002). This suggests that, based on temperature, T. montdorensis and N. cucumeris can survive in low tunnels during spring. However, the lower spring temperature may affect their reproduction, as there was no significant increase in mite numbers over time. Hypoaspis miles was not counted in this study, and its ability to survive in low tunnels during spring is not known. However, there were fewer WFT where plants had been treated with H. miles, suggesting that H. miles was able to survive and effectively reduce WFT.

The present study suggests that releases of T. montdorensis, N. cucumeris or H. miles combined with spinosad is more effective in managing WFT in low tunnel-grown strawberry than either spinosad applications or mite releases alone. Glasshouse studies also indicated that the

130 Chapter V: Compatibility of predatory mites and spinosad integration of T. montdorensis, N. cucumeris, or H. miles with spinosad is more successful in controlling WFT than spinosad or predatory mites alone (Chapters 3 and 4). Thoeming and Poehling (2006) reported that the application of azadiractin (Botanical, Neem Azal-U, 17% azadiractin) with N. cucumeris and H. aculeifer increased efficacy to 99% to control WFT. Ludwig and Oetting (2001) and Ludwig (2002) report that when the predatory bug Orius insidiosus Say (Heteroptera: Anthocoridae) was released after spinosad was applied to potted chrysanthemum and marigold, better control of WFT was achieved than with either approach alone. This combined approach will only be effective if the pesticide has little or no effect on the natural enemy, and is efficacious against the pest. Spinosad poses no detrimental effects to O. insidiosus, but is highly efficacious against WFT (Funderburk et al. 2000, Ludwig and Oetting 2001, Ludwig 2002). Chapman et al. (2009) state that an environmentally sound approach to managing Ostrinia nubilalis (Hübner) (Lepidoptera: Crambidae) is possible by releasing a parasitic wasp (Trichogramma ostriniae Pang et Chen (Hymenoptera: Trichogrammatidae)) and applying biorational insecticides (spinosad, indoxacarb and methoxyfenozide) to bell peppers. The combined use of reduced-risk insecticides (pyrethrin, insecticidal soap and mineral oil) and the ladybird Harmonia axyridis (Pallas) (Coleoptera: Coccinellidae), provides better management of the soybean aphid, Aphis glycine Matsumura (Hemiptera: Aphididae) on North American soybean (Kraiss and Cullen 2008). In the present study, spinosad initially reduced WFT numbers on strawberry plants, but WFT reached the economic threshold one week after application (>5 WFT adults per flower, (Steiner and Goodwin 2005) This also occurred when predatory mites were applied alone (without spinosad).

Effective management of WFT in low tunnel-grown strawberry was achieved by integrating spinosad and predatory mite releases. In this strategy, the WFT population is initially reduced by spinosad. Mites were released six days after spinosad was applied, and spinosad did not appear to impact negatively on them. When a lapse of time is maintained between pesticide application and mite release, spinosad has likely been degraded by photolysis (Viktorov et al. 2002) by the time that the mites are released. However, apart from direct mortality, spinosad could affect survival, reproduction or prey handling efficiency (Li et al. 2006, van Driesche et al. 2006). Laboratory and semi-field studies suggest that the direct application of pesticides and even aged residues can be harmful to many natural enemies (Li et al. 2006, van Driesche et al. 2006), including predatory mites (Hassan et al. 1987, Hassan et al. 1988, Kim and Seo 2001, Amano et al. 2004). Although spinosad is considered harmless to predatory mites, direct application and fresh residues are toxic to some species. Kongchuensin and Takafuji (2006) demonstrated that fresh spinosad residues up to 48 h old (12% suspensible concentrate) significantly reduced the number of eggs and the immature produced by Neoseiulus longispinosus (Evans) (Phytoseiidae). However, there was no effect on N. longispinosus if

131 Chapter V: Compatibility of predatory mites and spinosad exposed to residues after seven days. van Driesche et al. (2006) reported that fresh spinosad residues had no effect on the survival of N. cucumeris or Iphiseius degenerans (Berlese) (Phytoseiidae), but it lowered their oviposition rate. Khan and Morse (2006) tested the impact of four pesticides on the predatory mite Euseius tularensis Congdon and found a significant effect if mites were released five - six days after a spinosad application, but no effect if released after seven days. Since the toxicity of a given pesticide varies from species to species, a bioassay is needed to evaluate the toxicity of the recommended rate of spinosad for WFT management on T. montdorensis, N. cucumeris, and H. miles. This will allow mites to be released to avoid any toxicity posed by spinosad, while providing more effective control of WFT.

The efficiency of natural enemies in the a pest management program often varies from species to species (Chyzik et al. 1996, Berndt et al. 2004a, Berndt et al. 2004b, Wiethoff et al. 2004). The present study demonstrates that T. montdorensis, N. cucumeris, or H. miles were effective against WFT in low tunnel-grown strawberry. However, the effectiveness of these predatory mites against WFT appeared to differ. When released as a single species, T. montdorensis appeared to be the most effective predator, resulting in fewer WFT, followed by N. cucumeris and H. miles. This validates the previous findings (see Chapters 3 and 4) that T. montdorensis performed better in reducing WFT over the other two mite species. The success of natural enemies in a pest management program depends on several factors such as predation on prey stage, predation rate, within-plant distribution of predators and prey, and availability of supplemental food sources. Neoseiulus cucumeris is reported to predate on first instar larvae (Bakker and Sabelis 1989), which is a potentially limiting characteristic. Hypoaspis miles preys on WFT pupae (Glockemann 1992), though recent studies found that H. miles may also prey upon second instar larvae (Berndt 2003). The predation rate also varies from species to species. Berndt et al. (2004b) reported that efficiency of the predatory mites, Hypoaspis aculeifer Canestrini (Laelapidae) and H. miles against WFT were different, mainly because H. aculeifer ate more WFT compared to H. miles. Similarly, Brødsgaard (1989) and van Houten et al. (1995) reported that N. cucumeris consumed more thrips compared to H. aculeifer and Neoseiulus barkeri (Hughes) (Phytoseiidae). Rhodes and Liburd (2006) report variation in the performance of predatory mites against two-spotted spider mites in strawberry. In field-grown strawberry, Phytoseiulus persimilis Athias-Henriot takes a longer time to bring two-spotted spider mites under control, compared to Neoseiulus californicus McGregor (Rhodes and Liburd 2006). It was reported that T. montdorensis can prey 7-14 first instar WFT larvae per day (Steiner et al. 2003). While, on an average, N. cucumeris and H miles can prey six and two first instar WFT larvae per day respectively (Berndt et al. 2004b, Zilahl-Balogh et al. 2007). The distribution of predatory mites can also influence their effectiveness in pest management programs. Typhlodromips montdorensis is a generalist predator and has the ability to distribute rapidly on

132 Chapter V: Compatibility of predatory mites and spinosad different parts of the plant (Steiner and Goodwin 1998, 2001), which may give it an advantage over the other two species. Steiner and Goodwin (1998, 2001) also reported that the when release, T. montdorensis rapidly distribute over the whole plants, whilst the within-plant distribution of N. cucumeris is uneven i.e. distribute to certain part of the plant (Messelink et al. 2006). Neoseiulus cucumeris prefers the lower part of the of plant, while WFT prefer to remain on the upper part of the plants (Messelink et al. 2006). Hypoaspis miles is a soil-dwelling predator, which limits its prey to thrips pupae.

Variation in the reproduction and development of predatory mites often plays an important role in their success (Messelink et al. 2006). The present field trial indicates that in single-species release, N. cucumeris numbers were fewer than T. montdorensis. However, it is not known why there were more T. montdorensis than N. cucumeris, as the same numbers of both species were released. The efficacy of predatory mites may be influenced by the presence of supplemental food such as pollen. In chrysanthemum, for example, the presence of pollen as supplemental food reduced the predation efficiency of N. cucumeris on WFT by up to 55% (Skirvin et al. 2007). Similarly, van Rijn and Tanigoshi (1999) and van Rijn et al. (2002) reported that Iphiseius degenrans Berlese (Acari) also feeds on pollen, effecting the predation of WFT on cucumber. The presence of pollen as a supplement food has no influence on T. montdorensis (Steiner and Goodwin 2002). Foraging efficiency of commercially supplied predators varies from species to species which may also affect their effectiveness in reducing pest population (Steiner and BjØrnson 1996).

Although the single-species release of predatory mites in all treatments appears to be effective for WFT management in low-tunnel grown strawberry, the release of a foliage inhabiting (T. montdorensis or N. cucumeris) and a soil-dwelling predator (H. miles) was more effective. There is a growing trend to use two or more species of natural enemies to manage insect populations effectively (Premachandra et al. 2003, Avilla et al. 2004, Blümel 2004, Brødsgaard 2004, Chow and Heinz 2004, Hoddle 2004, Shipp and Ramakers 2004, Thoeming and Poehling 2006, Chow et al. 2008) might be partly due to the complementary effects on each other. Brodeur et al. (2002) recognised that the release of multiple-predator species as an effective strategy that would ideally suppress pest populations in a manner that is more economically viable than the use of single-predator species. Wiethoff et al. (2004) reported cucumber plants had WFT when N. cucumeris and H. miles were applied together than either N. cucumeris or H miles released alone. The combined release of P. persimilis and N. californicus also provided long-term effective suppression of carmine spider mite Tetranychus cinnabarinus (Boisduval) (Tetranychidae) in potted gerbera plants, compared to their individual release (Schausberger and Walzer 2001). Premachandra et al. (2005) reported that the emergence of WFT adults was

133 Chapter V: Compatibility of predatory mites and spinosad significantly lower in the combined application of entomopathogenic pathogens and the predatory mite, H. aculeifer, than applications of either alone. The synergistic effects can be expected if a plant-dwelling predator evokes escape behaviour of the prey and makes the prey available for ground-dwelling predators (Losey and Denno 1999). However, the release of T. montdorensis and N. cucumeris may not be effective since they may compete for the same prey (Janssen et al. 1998). As a result, one species might outcompete the other. Although the present study did not address this, it appears that when T. montdorensis and N. cucumeris are released in combination, T. montdorensis numbers were higher than N. cucumeris. However, when applied in triple-species combinations, mite performance was only second to the double-species combination of T. montdorensis and H. miles. When all species were released together there appeared to be no interaction between T. montdorensis and N. cucumeris. When these two predatory mites were applied in a double-species combination, 150 mites of each species were released per m-2, but only 100 individuals per m-2 of each species in the triple-species combination. The higher ratio used in the double species released may have increased the chance of interspecific competition. Therefore, for the successful implementation of multiple releases of predatory mites in a pest management program, further studies need to be carried out to determine if intraguild predation occurs and preference between primary prey (pest), secondary prey (predator) and other interactions. Furthermore, it is important to determine the optimal release rate of predatory mites that can effectively reduce the pest population, whilst having no negative impact on each other.

In conclusion, the integration of spinosad and multiple combinations of predatory mites (T. montdorensis, N. cucumeris, and H. miles) could be a sustainable strategy for WFT management in low tunnel-grown strawberry during spring. Although both spinosad treatments (‘spinosad then mites’ or ‘mites then spinosad’) appeared to be effective against WFT, applying spinosad before releasing the mites seems to be the more effective strategy, which has several advantages. Firstly, the initial WFT population is reduced by spinosad. It is known that biological control of arthropod pests by predatory mites can fail if the initial pest population is very high (Malezieux et al. 1992). Secondly, direct toxicity of spinosad is harmful to T. montdorensis, N. cucumeris and H. miles. Spraying first eliminates the problem of direct exposure. Thirdly, this strategy enhances the chance to eliminate resistant populations. If the lapse of times between spinosad application and mite release could be further shortened, control could be further improved. Studies based on residual toxicity of spinosad to predatory mites will allow the precise residual threshold to be determined. In addition, the present study commenced during spring. The residual toxicity of spinosad to predatory mites may vary from season to season. van de Veire et al. (2002) reported that abamectin (bioinsecticide) is more persistent in

134 Chapter V: Compatibility of predatory mites and spinosad spring than summer. Therefore, seasonal variation also needs to be considered and the supplementary release of predatory mites after five weeks, may also be needed.

5.5 Literature cited

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Medhurst, A., and M. Y. Steiner. 2001. Western Flower Thrips and Strawberries. National Strategy for the Management of WFT & TSWV, East Melbourne, Victoria, Australia. Messelink, G. J., S. E. F. van Steenpaal, and P. M. J. Ramakers. 2006. Evaluation of phytoseiid predators for control of western flower thrips on greenhouse cucumber. Biocontrol 51: 753-768. Mound, L. A. 1997. Biological diversity, pp. 197-216. In T. Lewis [ed.], Thrips as Crop Pests. CAB International, Wallingford, UK. Pietrantonio, P. V., and J. H. Benedict. 1999. Effect of new cotton insecticide chemistries, tebufenozide, spinosad and chlorfenapyr on Orius insidiosus and two Cotesia species. Southwestern Entomologist 24: 21-29. Premachandra, D. W. T. S., C. Borgemeister, and H.-M. Poehling. 2005. Effects of Neem and Spinosad on Ceratothripoides claratris (Thysanoptera: Thripidae), an Important Vegetable Pest in Thailand, Under Laboratory and Greenhouse Conditions. Journal of Economic Entomology 98: 438-448. Premachandra, W. T. S. D., C. Borgemeister, O. Berndt, and R.-U. Ehilers. 2003. Combined release of entomopathogenic nematodes and the predatory mite Hypoaspis aculeifer to control soil-dwelling stages of western flower thrips Frankliniella. Biocontrol 48: 529-541. Quinn, G. P., and M. J. Keough. 2002. Experimental design and data analysis for biologists. University Press, Cambridge. Ramakers, P. M. J. 1988. Population dynamics of the thrips predators Amblyseius mckenziei and Amblyseius cucumeris (Acarina: Phytoseiidae) on sweet pepper. Netherland Journal of Agricultural Science 36: 247-252. Rhodes, E. M., and O. E. Liburd. 2006. Evaluation of predatory mites and acramite for control of two-spotted spider mites in strawberries in North Central Florida. Journal of Economic Entomology 99: 1291-1298. Riduavets, J. 1995. Predators of Frankliniella occidentalis (Perg.) and Thrips tabaci Lind.: a review. Wageningen Agricultural University Papers 95: 43-87. Rodrìguez-Reina, J. M., F. Garcìa-Marì, and F. Ferragut. 1992. Actividad depredatoria de varios acarosfioseidos sobre distintos estados de desarrollo del trips de las flores Frankliniella occidentalis (Pergande). Bol. San. Veg. Palagas 18: 253-263. Sabelis, M. W., and P. C. J. Van Rijn. 1997. Predation by Insects and Mites, pp. 259-354. In T. Lewis [ed.], Thrips as Crop Pests. CAB International, Wallingford, UK. SAS 2002-2003. SAS 9.1 computer program, version 9.1. By SAS, Cary, NC, USA. Schausberger, P., and A. Walzer. 2001. Combined versus Single Species Release of Predaceous Mites: Predator-Predator Interactions and Pest Suppression. Biological Control 20: 269-278. Shipp, J. L., and P. M. J. Ramakers. 2004. Biological control thrips on vegetable crops, pp. 265-276. In K. M. Heinz, R. G. van Driesche and M. P. Parrella [eds.], Biocontrol in protected culture. Ball Publishing, Batavia, Illinois. Skirvin, D. J., L. Kravar-Garde, K. Reynolds, J. Jones, A. Mead, and J. Fenlon. 2007. Supplemental food affects thrips predation and movement of Orius laevigatus (Hemiptera: Anthocoridae) and Neoseiulus cucumeris (Acari: Phytoseiidae). Bulletin of Entomological Research 97: 309-315. Sparks, T. C., G. D. Thomson, H. A. Kirst, M. B. Hertlein, L. L. Larson, T. V. Worden, and S. T. Thibault. 1998. Biological activity of the spinosyn, new fermentation derived insect control agents, on tobacco budworm (Lepidoptera: Noctuidae) larvae. Journal of Economic Entomology 91: 1277-1283. Steiner, M. Y., and S. BjØrnson. 1996. Performance of Phytoseilus persimilis and other biological control agents - on what are basing our standards? I.B.O.C./W.P.R.S. Bulletin 5: 65-72. Steiner, M. Y., and S. Goodwin. 1998. Development and marketing of an IPM package for western flower thrips, Frankliniella occidentalis (Pergande). Phase II: Evaluation and producing the natural enemies. HRDC/HSNA Report. NSW Agriculture, Gosford, Australia.

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Steiner, M. Y., and S. Goodwin. 2001. Development and marketing of an IPM package for western flower thrips, Frankliniella occidentalis. Phase III: Development and evaluation of usage protocols for newly developed western flower thrips biocontrol agents. HRDC report. NSW Agriculture, Gosford, Australia. Steiner, M. Y., and S. Goodwin. 2002. Development of a new thrips predator, Typhlodromips montdorensis (Schicha) (Acari: Phytoseiidae) indigenous to Australia. Bulletin of IOBC/WPRS 25: 245-247. Steiner, M. Y., and S. Goodwin. 2005. Management of thrips (Thysanoptera: Thripidae) in Australian strawberry crops: within-plant distribution characteristics and action thresholds. Australian Journal of Entomology 44: 175-185. Steiner, M. Y., S. Goodwin, T. M. Wellham, I. M. Barchia, and L. J. Spohr. 2003. Biological studies of the Australian predatory mite Typhlodromips montdorensis (Schicha) (Acari: Phytoseiidae), a potential biocontrol agent for western flower thrips, Frankliniella occidentalis (Pergande) (Thysanoptera: Thripidae). Australian Journal of Entomology 42: 124-130. Thoeming, G., and H.-M. Poehling. 2006. Integrating soil-applied azadiractin with Amblyseius cucumeris (Acari: Phytoseiidae) and Hypoaspis miles (Acari: Laelapidae) for the management of Frankliniella occidentalis (Thysanoptera: Thripidae). Environmental Entomology 35: 746-756. Thompson, D. G., B. J. Harris, L. J. Lanteigne, T. M. Buscarini, and D. T. Chartrand. 2002. Fate of spinosad in litter and soils of a mixed conifer stand in the Acacian forest region of New Brunswick. Journal of Agricultural and Food Chemistry 50: 790-795. Thompson, G. D., K. H. Michel, R. C. Yao, J. S. Mynderse, C. T. Mosburg, T. V. Warden, E. H. Chio, T. C. Sparks, and S. H. Hutchins. 1997. The discovery of Saccharopolyspora spinosa and a new class of insect control products. Down to Earth 52. Ullio, L. 2002. Australia's national strategy for the management of western flower thrips (WFT), Frankliniella occidentalis (Pergande), pp. 687-689. In T. Hietaranta, M.-M. Linna, P. Palonen and P. Parikka [eds.], Proceedings of the fourth International Strawberry Symposium. Acta Horticulturae, MTT Agrifood Research, Finland. van de Veire, M., M. Klein, and L. Tirry. 2002. Residual activity of abamectin and spinosad against the predatory bug Orius laevigatus. Phytoparasitica 30: 525-528. van Driesche, R. G., S. Lyon, and C. Nunn. 2006. Compatibility of spinosad with predacious mites (Acari: Phytoseiidae) used to control western flower thrips (Thysanoptera: Thripidae) in greenhouse crops. Florida Entomologist 89: 396-401. van Driesche, R. G., K. M. Heinz, J. C. Van Lenteren, A. J. M. Loomans, R. Wick, T. Smith, P. Lopes, J. P. Sanderson, M. Daughtrey, and M. Brownbridge. 1998. Western flower thrips: a review of its biological control and other methods. Floral Facts, pp. 30. Ed. Univ. of Massachusetts, Amherst, MA. van Houten, Y. M., P. van Stratum, J. Bruin, and A. Veerman. 1995. Selection for non- diapause in Amblyseius cucumeris and Amblyseius barkeri and exploration of the effectiveness of selected strains for thrips control. Entomologia Experimentalis et Applicata 77: 289-295. van Rijn, P. C. J., and L. K. Tanigoshi. 1999. Pollen as food for the predatory mites Iphiseius degenerans and Neoseiulus cucumeris (Acari: Phytoseiidae): dietary range and life history. Experimental and Applied Acarology 23: 785-802. van Rijn, P. C. J., Y. M. van Houten, and M. W. Sabelis. 2002. How plants benefit from providing food to predators even when it is also edible to herbivores. Ecology 83: 2644- 2679. Viktorov, A. V., E. N. Preshokov, and V. A. Driniaev. 2002. Comparative analysis of degradation of native avermectin and spinosyn by UB-HPLC. Antibiot Khimioter 47: 6- 10. Villanueva, R. T., and J. F. Walgenbach. 2005a. Development, Oviposition, and Mortality of Neoseiulus fallacis (Acari: Phytoseiidae) in Response to Reduced-Risk Insecticides. Journal of Economic Entomology 98: 2114-2120.

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Villanueva, R. T., and J. F. Walgenbach. 2005b. Development, oviposition, and mortality of Neoseiulus fallacis (Acari: Phytoseiidae) in response to reduced-risk Insecticides. Journal of Economic Entomology 98: 2114-2120. Wiethoff, J., H.-M. Poehling, and R. Meyhöfer. 2004. Combining plant- and soil-dwelling predatory mites to optimise biological control of thrips. Experimental and Applied Acarology 34: 239-261. Williams, T., J. Valle, and E. Vinuela. 2003. Is the naturally derived insecticide Spinosad compatible with insect natural enemies? Biocontrol Science and Technology 13: 459- 475. Zalom, F. G., P. A. Phillips, N. C. Toscano, and M. Bolda. 2001. Strawberry, Western flower thrips. UC IPM Pest Management Guidelines: Strawberry. UC ANR Publication 3468, Insects and Mites. Davis, California, USA. http://www.ipm.ucdavis.r734301211.html. Zar, J. H. 1999. Biostatistical Analysis. Prentice Hall International, Upper Saddle River, New Jersey. Zhao, J. Z., Y. X. Li, H. L. Collins, L. Gusukuma-Minuto, R. F. L. Mau, G. D. Thompson, and A. M. Shelton. 2002. Monitoring and characterization of diamondback moth (Lepidoptera: Plutellidae) resistance to spinosad. Journal of Economic Entomology 95: 430-436. Zilahl-Balogh, G. M. G., J. L. Shipp, C. Cloutier, and J. Brodeur. 2007. Predation by Neoseiulus cucumeris on western flower thrips, and its oviposition on greenhouse cucumber under winter vs. summer conditions in a temperate climate. Biological Control 40: 160-167.

140 CHAPTER VI

Compatibility of spinosad with predaceous mites (Acari) used to control western flower thrips, Frankliniella occidentalis (Pergande) (Thysanoptera: Thripidae)

Keywords: Spinosad, Frankliniella occidentalis, Typhlodromips montdorensis, Neoseiulus cucumeris, Hypoaspis miles, direct toxicity, residual toxicity, preference, LT25

Abstract

Spinosad™ (Dow AgroSciences, USA) is a biopesticide widely used for control of western flower thrips, Frankliniella occidentalis (Pergande) (Thysanoptera: Thripidae). Spinosad is reported to be non-toxic to several predatory mite species used for the biological control of thrips, and is recommended for use in integrated pest management programs for this reason. Predatory mites (Acari) have recently become available to Australian growers for the control of thrips. This includes Typhlodromips montdorensis (Schicha), Neoseiulus cucumeris (Oudemans) and Hypoaspis miles (Berlese), which feed on thrips larvae or pupae. This study investigated the impact of direct and residual toxicity (contact) of spinosad (recommended rate: 80 mL/100 L, 0.096 g a.i./L) on F. occidentalis and direct and residual (contact, indirect via consumption of intoxicated thrips larvae and simultaneous exposure via contact and consumption of intoxicated thrips larvae) toxicity on predatory mites. This study also investigated the repellency of spinosad residues to mites. Direct contact with spinosad effectively reduced the numbers of F. occidentalis adults and larvae, causing >96% mortality. Two to 96 h old spinosad residues were also toxic to F. occidentalis. Direct exposure to spinosad resulted in >90% mortality of all three mite species. Thresholds for the residual toxicity (contact) of spinosad LT25 (lethal time for 25% mortality) were estimated as 4.2, 3.2 and 5.8 days for T. montdorensis, N. cucumeris and H. miles respectively. When mites were simultaneously exposed to spinosad residues and fed spinosad-intoxicated thrips larvae, toxicity increased. Residual thresholds were re-estimated as 5.4, 3.9 and 6.1 days for T. montdorensis, N. cucumeris, and H. miles respectively. Spinosad residues were also repellent to mites. Residues aged two to 48 h repelled T. montdorensis and H. miles, and residues aged two to 24 h repelled N. cucumeris. These data suggest that mites could be safely released six days after spinosad is applied for the management of F. occidentalis.

6.1 Introduction

Spinosad (Success™; Dow AgroSciences Australia) is a novel pesticide derived from fermentation of the actinomycete, Saccharopolyspora spinosa Mertz and Yao (Sparks et al. Chapter VI: Bioassay

1998). It is active against Lepidoptera, Diptera, and Thysanoptera (Cloyd and Sadof 2001). Spinosad is classified as an environmentally and toxicologically reduced-risk chemical (Cleveland et al. 2002, Thompson et al. 2002). Spinosad was first registered in Australia in 1998 and the USA in 1997 for use in cotton (Thompson and Hutchins 1999), and is now used in more than 180 crops worldwide (Zhao et al. 2002). The marketing of spinosad has focused on its favourable environmental profile, emphasising its potential for use in the integrated pest management (IPM) systems (Thompson and Hutchins 1999, Thompson et al. 2000). Several studies suggest that spinosad is less toxic to natural enemies including predatory mites, than their prey (Miles and Dutton 2000, Thompson et al. 2000, Medina et al. 2001, Holt et al. 2006, Kim et al. 2006, Arthurs et al. 2007). However, it has been reported that spinosad toxicity to natural enemies is variable (Cote et al. 2004).

Spinosad is the primary insecticide used to control western flower thrips (WFT), Frankliniella occidentalis (Pergande) (Thysanoptera: Thripidae) in strawberry in Australia. However, there are concerns about WFT evolution of resistance to spinosad (Herron and James 2005, Bielza et al. 2007). Biological control has been successfully incorporated with pesticide use to manage WFT in commercial glasshouses in North America and Europe (Chambers and Sites 1989, Gillespie 1989, Gilkeson 1990, Brødsgaard 2004, Shipp and Ramakers 2004). The most widely employed natural enemies are predatory phytoseiid mites (Acari) that feed on thrips larvae (Riudavets 1995, Sabelis and Van Rijn 1997). In Australia, predatory mites Typhlodromips montdorensis (Schicha) (Phytoseiidae), Neoseiulus cucumeris (Oudemans) (Phytoseiidae) and Hypoaspis miles (Berlese) (Laelapidae), have shown some potential for controlling WFT (Steiner and Goodwin 2000). Nevertheless, it is not known what effect spinosad has on these species of predatory mites that could be used in an IPM program in Australian strawberry production. To determine whether spinosad could be integrated with mites into existing Australian strawberry pest management, a series of laboratory bioassays were carried out to evaluate the toxicity of spinosad to the mites and to determine if predator-prey interactions would be affected. Specifically, the objectives of this study were to evaluate:

(i) Toxicity of direct contact with the recommended rate of spinosad to WFT and predatory mites. (ii) Residual toxicity of spinosad to WFT. (iii) Residual toxicity (contact, indirect via consumption of intoxicated WFT and, simultaneous exposure to spinosad via direct contact and consumption of treated prey) and residual thresholds of spinosad for T. montdorensis, N. cucumeris and H. miles. (iv) Repellency of spinosad residues to mites.

142 Chapter VI: Bioassay

6.2 Materials and methods

Trials were conducted in a controlled temperature (CT) room (25±1⁰C, 50-60% RH, 16:8 h L:D regime) from June to September 2008 at the University of Western Australia (UWA).

6.2.1 Source of cultures

6.2.1.1 Strawberry plants

Strawberry runners [Fragaria ananassa Duchesne (Rosaceae)] cultivar ‘Camino Real’ were planted into pots (32.5l x 32.5w x 40.5h cm) containing potting mix (Baileys Fertilisers, Rockingham, WA) in glasshouses at the Department of Agriculture and Food WA (DAFWA) and UWA. All pots were fitted with sprinklers with automatic timers. The plants were watered every third day. A liquid fertiliser (Thrive ®, Yates, Australia NPK: 12.4: 3: 6.2; rate: 5mL/2 L water) was applied once a month.

6.2.1.2 Western flower thrips (WFT)

A glasshouse colony of WFT was established from individuals initially collected from calendula flowers, Calendula officianalis L. (Asterales: Asteraceae) in a glasshouse at DAFWA. From July 2006, WFT colonies were maintained on potted calendula at UWA. Calendulas were grown from seeds collected from calendula plants maintained at DAFWA. Calendula seeds were sown in plastic pots (50 x 100 mm) containing potting mix, and kept in insect-proof Perspex cages (500 mm high, 420 mm deep and 400 mm wide). The cages were fitted with thrips proof mesh net (105 µ; Sefar Filter Specialists Pty Ltd., Malaga, WA), and the entire cage was fitted on top of a Nylex tote box (320 x 420 mm; Blyth Enterprises Pty Ltd, Australia). When calendula plants were flowering, WFT adults were released at the base of the plant to maintain the colony.

6.2.1.3 Predatory mites

Predatory mites [Typhlodromips montdorensis, Neoseiulus cucumeris, and Hypoaspis miles] were sourced from commercial Australian suppliers (Biological Services, SA; Manchil IPM Services, WA; and Beneficial Bug Company, NSW). Mites were provided in plastic buckets or plastic bag containing vermiculite. Trials were conducted immediately upon receipt of predatory mites.

143 Chapter VI: Bioassay

6.2.2 Experiment 1: Direct toxicity of spinosad to WFT and predatory mites

6.2.2.1 Western flower thrips

The experiment was conducted in the controlled temperature (CT) room. Twenty cold- anaesthetised WFT adults (2-3 d old) collected from the colony were placed on a paper towel and lightly sprayed with either 5mL of diluted spinosad (treatment) or water (control) with a hand-held atomiser (Hills Sprayers, BH220063). Spinosad (Success™, 120 g/L emulsifiable concentrate, Dow AgroSciences Australia Ltd) was applied at the recommended rate of 80 mL/100 L rate (0.096 g a.i./L). After spraying excess spinosad or water (if any) was gently removed with a soft tissue. Thrips (n = 20) were then transferred to a glass Petri dish (150 x 15 mm) containing an excised strawberry leaf placed adaxial side up on a piece of moistened filter paper. The filter paper was glued to the bottom of the Petri dish to ensure that thrips could not hide between the Petri dish and filter paper. The leaf petiole was covered with cotton wool soaked in 10% sugar solution to extend leaf life and the edge of the strawberry leaf was glued to the filter paper. The Petri dish was covered with a screen (mesh net 105 µ), and the side of the Petri dish was sealed with paraffin film (Parafilm M®, Micro Analytix Pty Ltd) to prevent thrips escaping (Figure 6.1). This procedure was repeated 20 times to produce 20 Petri dishes each with 20 adult thrips. The procedure was repeated a second time instead using first instar thrips larvae to produce 20 Petri dishes each with 20 first instar larvae.

Figure 6.1 Diagrammatic representation of the testing arena used for toxicity test.

Petri dishes were then placed (randomly) on a laboratory bench and thrips were monitored for mortality. Spinosad is slow acting (Bret et al. 1997) and cumulative mortality of a test organism usually plateaus at 2 days (48 h) to 6 days (144 h) after exposure (Viñuela et al. 2001, Cisneros et al. 2002). Consequently, Petri dishes with WFT adults were examined at 6, 24, 48, 72, and 96 h post-release exposure periods under a stereomicroscope. The WFT larvae were checked at 6, 24, 48 and 72 h post-release exposure periods (after 72 h post-release exposure periods, all larvae had pupated). WFT adults or larvae were recorded as dead if they did not respond to probing with a fine paintbrush.

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6.2.2.2 Predatory mites

To assess the effect of direct exposure to spinosad on the three species of predatory mites, I used the same bioassay method as described above except for the following differences. A thin barrier of Tac-Gel (Stickem™, The Olive Centre, Australia) was applied to the edge of the leaf to prevent the escape of predatory mites. In addition, first or second instar WFT larvae were added to Petri dishes to provide food for the mites. In each Petri dish, there were 200 first instar thrips larvae for T. montdorensis [average 10 larvae per mite (Steiner et al. 2003)], 100 first instar larvae for N. cucumeris [average six larvae per mite (Zilahl-Balogh et al. 2007)] and 40 second instar for H. miles [two larvae per mite (Berndt et al. 2004)]. During the trial period, additional thrips larvae were added to the Petri dishes as required. Mortality of the mites was monitored at 6, 24, 48 and 72 h post-release exposure periods (as no mite mortality occurred after 72 h post-release exposure period). Mites were recorded as dead if they did not respond to probing with a fine paintbrush. There were 20 individuals per Petri dish and 20 Petri dishes for each species.

6.2.3 Experiment 2: Residual toxicity of spinosad to WFT and predatory mites

6.2.3.1 Mortality of WFT and mites to spinosad residues (contact) over time

In this experiment, WFT and mites were placed on strawberry leaves with different levels of spinosad residue. Prior to spraying, 3-4 weeks old potted strawberry plants (CA Camino Real) were moved to the CT room and split into groups (2, 12, 24, 48, 72, 96, 120 and 144 h old residues and control). Leaves were marked with a permanent marker at the base of the petiole, to enable leaves with spinosad residues to be tracked. Spinosad (at the recommended rate of 80mL/100 L, 0.096 g a.i./L) or distilled water (control) was applied to both sides of the leaves with a hand-held atomiser until run-off (method devised after Eger et al. (1998)). In order to get residues with different ages at the same times, first those plants in the 144 h old residue treatment were sprayed once, followed by the spray on subsequent group of plants. Strawberry plants in the control group were sprayed with distilled water 24 h prior to the experiment. Separate atomisers were used for the spinosad and the water. After spraying, all plants were covered with a modified thrips-cage (45 x 35 cm) made from mesh (105 µ) supported by a quadrate steel-rod stand to exclude thrips from the plants. The bottom end of the cage was taped to the pot with tape. The top end of the cage was closed with a rubber band. Plants covered with mesh cage were then returned to the glasshouse and kept until use.

145 Chapter VI: Bioassay

Spinosad was also applied to the Petri dishes (150 x 15 mm; testing arena) at the same time in a similar fashion mentioned above in order to obtain strawberry leaves and dishes of the same exposure age. Petri dishes were held with the open end sidewise on a stand and sprayed the inner side with spinosad solution using an atomiser until run-off. Both parts of a Petri dish were sprayed with spinosad. After spraying, once the excess liquid (if any) run-off, the Petri dishes were kept on a tray and allowed to dry in a glasshouse for one and half hours. Dried Petri dishes were stored in separate plastic trays according to residue treatments (2, 12, 24, 48, 72, 96, 120, and 144 h old residues and the control) and kept in the CT room until use.

At the onset of the experiment, strawberry plants sprayed with spinosad or water were brought to the CT room. Strawberry leaves were removed from the sprayed plants and placed adaxial side up at the bottom of a glass Petri dish (150 x 15 mm) with the same residue ages. Each leaf was glued to the bottom of a Petri dish to prevent arthropods from hiding underneath. In addition, a thin barrier of Tac Gel was applied to the edge of the leaf to prevent escape; however, WFT adults can fly. A spinosad-treated Petri dish was used in order to keep the test subject in contact with spinosad residues, if any test individual somehow escaped the treated leaf surface. The petiole was covered with cotton soaked in a 10% sugar solution to extend leaf life. Twenty WFT adults (2-3d old), 20 WFT larvae (1d old), or 20 adult mites (each species) were released onto the leaf surface. A screen cover (mesh net 105 µ) was placed over the Petri dish as previously described. The strawberry leaf and Petri dish was sprayed with distilled water to be used as a control. Petri dishes with WFT and predatory mites were then randomly arranged on a laboratory bench in the CT room. For each of the eight residue times and the control (water) there were 20 replicates of each mite species and WFT adults and larvae. For the mite bioassays, WFT larvae were added (at a rate described above) as food each day for 2-3 hrs, and then removed to ensure that mites were not consuming spinosad-intoxicated larvae.

Mortality was checked under a stereomicroscope. WFT (larvae and adults) and predatory mites (T. montdorensis, N. cucumeris and H. miles) were recorded as dead if they did not respond to probing with a fine paintbrush. WFT adults were checked at 6, 24, 48, 72, and 96 h post-release exposure periods. WFT Larvae were checked at 6, 24, 48, and 72 h post-release exposures (after 72 h all the larvae had pupated). Predatory mite mortality was checked at 24, 48 and 72 h post- release exposure periods (no mortality occurred after 72 h post-release exposure).

6.2.3.2 Indirect exposure of spinosad to predatory mites via consumption of intoxicated WFT larvae

Twenty-four hours prior to the trial, newly emerged first instar WFT larvae were collected from the stock colony. Larvae were transferred to a glass Petri dish (150 x 15 mm) containing a

146 Chapter VI: Bioassay cotton wool ball soaked in a 10% sugar solution, then stored in the CT room for 12 h (to ensure feeding on spinosad- or water-treated leaf at a later stage). Larvae were then released onto excised strawberry leaves previously treated with spinosad (2, 12, 24, 48, 72, 96, 120, or 144 h old residues) or control (water) in a Petri dish and allowed to feed for 12 hours. A thin barrier of Tac Gel was applied to the edge of the leaf to prevent larvae from escaping the leaf. After 12 h, WFT larvae were transferred onto a fresh, untreated strawberry leaf in a Petri dish (150 x 15 mm), containing 20 adult mites of one species of mite. The leaf petiole was covered with cotton soaked in a 10% sugar solution to keep the leaf fresh during the trial period. A thin barrier of Tac Gel was applied to prevent either WFT larvae or predatory mites escaping the leaf surface. In each Petri dish, 200, 120 and 40 first or second instar of WFT larvae that had not been exposed to spinosad were released for T. montdorensis, N. cucumeris and H. miles respectively. During the trial period, additional thrips larvae were added to the Petri dishes as required. All Petri dishes with WFT larvae and predatory mites were covered with mesh net (105 µ) and sealed with paraffin film. All Petri dishes with predatory mites were kept on a laboratory bench (randomly arranged) and checked under stereomicroscope at 24, 48 and 72 h post-release exposure periods. Predatory mites were recorded as dead if they did not respond to probing with a fine paintbrush. Each treatment (residue age) and control (water) was replicated 20 times (20 x 20 = 400 individuals) for each species of predatory mites.

6.2.3.3 Toxicity of spinosad to predatory mites via consumption of intoxicated WFT larvae and direct exposure to spinosad residues of different ages

This bioassay evaluated mortality of predatory mites (T. montdorensis, N. cucumeris and H. miles) via two exposure routes: (i) via consumption of intoxicated WFT larvae, and (ii) simultaneous direct exposure to spinosad residues on strawberry leaves. WFT larvae were collected and exposed to spinosad as described in 6.2.3.2; strawberry leaves were prepared as described in 6.2.3.1. Testing arenas were prepared using the same age residue of leaf, Petri dish, and WFT larvae. A thin layer of Tac Gel was applied at the edge of the leaf to prevent escape of WFT larvae or predatory mites from the testing arena. Intoxicated first instar WFT larvae were released onto the leaf surface in a Petri dish at a rate described above. Twenty mite adults of one species were then released onto the leaf surface and the Petri dish was covered and sealed as described above. A Petri dish and strawberry leaf were sprayed with distilled water to be used as a control. All Petri dishes were arranged randomly on the laboratory bench in the CT room and mortality was checked under a stereomicroscope at 24, 48, and72 h post-release exposure periods. The above procedure was repeated 20 times (20 x 20 = 400 individuals) with spinosad residues aged 2, 12, 24, 48, 72, 96, 120 and 144 h for each species of predatory mite.

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6.2.4 Experiment 3: Repellency of spinosad to predatory mites (choice test)

To evaluate the possible repellency of different aged residues of spinosad (2, 12, 24, 48, 72, and 96 h old) to T. montdorensis, N. cucumeris and H. miles, a choice test was conducted in the CT room using a method modified from van Driesche et al. (2006). A test arena was constructed by cutting spinosad-treated (leaves with spinosad treated residues 2, 12, 24 48, 72 and 96 h old) or water-treated (control) strawberry leaves in half along the mid-vein. Leaf halves (one half consisting of a spinosad treated leaf, the other the control leaf) were then taped onto filter paper to form a full leaf, with a 2 mm channel between the halves for mite releases. A thin barrier of Tac Gel was applied to the edges of the leaf and filter paper. Additionally, at the centre of each leaf half, a rounded arena was created using Tac-Gel where WFT larvae were released as an attractant. The Tac-Gel barrier prevented WFT larvae moving from the spinosad treated leaf to water treated leaf half. In a pilot experiment, it was found that most of the adults of predatory mite (T. montdorensis, N. cucumeris, or H. miles) did not move from its release point if no WFT larvae were provided as an attractant. Before predatory mites were released into the test arena, 15 WFT larvae (first instar for T. montdorensis and N. cucumeris and second instar for H. miles) were released onto each leaf half. Ten adult mites of one species were then placed on the filter paper channel in each Petri dish. The test arena was observed three times during a 60 min period at three 20 min intervals (three observations) and the number of predatory mites on the spinosad- and the water-treated leaf surfaces was counted at each observation. With each species of predatory mite, the experiment was repeated 20 times (20 x 10 = 200 individuals of each species) for each residue (aged 2, 12, 24, 48, 72 and 96 h). A new testing arena was used for each test.

6.2.5 Data analysis

For each experiment in which mortality was assessed, the number of individuals that died at each observation was counted and expressed as a percentage of the total number of individuals in the arena and corrected using Abbott’s formula (Abbott 1925):

Abbott’s formula takes into account the proportion of control thrips or mites dying in the trial that have not been exposed to the spinosad, and amongst those that have been exposed to spinosad, some may die of natural causes. In these trials, control mortality of either thrips or mites never exceeded 5%.

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The cumulative mortality of predatory mites due to direct toxicity of spinosad was analysed with one-way ANOVA (Proc Mixed Procedure) to differentiate mortality amongst species. In addition, if mortality differed between post release exposure periods, mortality data of each species of predatory mite (T. montdorensis, N. cucumeris, and H. miles) was subjected to separate one-way ANOVAs. To determine if residues of different ages affected the mortality of WFT, the mortality of adults and larvae was analysed with a one-way ANOVA for each post- release exposure period. Cumulative mortality of WFT adult and larvae was also analysed with separate one-way ANOVAs. Similarly, the influence of residual age of spinosad on mortality of each species of predatory mites was analysed with separate one-way ANOVAs for each post release exposure period. Least square means significant difference tests at the 5% probability level were used to test for treatment differences.

Based on the mortality of predatory mites, results were classified into four categories following the IOBC (International Organization of Biological Control) guidelines and ranked as: 1 = harmless (<25% mortality) 2 = slightly harmful (25-50% mortality) 3 = moderately harmful (51- 75% mortality) 4 = harmful (>75% mortality) (Sterk et al. 1999).

The persistence of spinosad for each species of predatory mites was also classified according to the time taken to lose toxicity (<30% mortality): A = short lived (<5 days) B = slightly persistent (5-15 days) (Hassan et al. 1994, Sterk et al. 1999).

To determine the contact or contact and indirect toxicity (via consumption of intoxicated WFT larvae and simultaneous exposure to residues), the residual threshold of spinosad for T. montdorensis, N. cucumeris and H. miles was estimated with Probit analyses (Finney 1971).

The LT25 (lethal time of 25% mortality) was used, which is considered an acceptable level (Shipp et al. 2000).

The repellency of spinosad residues (2, 12, 24, 48, 72 and 96 h old) to predatory mites was analysed by paired t -test (Proc ttest Procedure).

Prior to statistical analyses data were transformed using √(x + 0.5) (Healy and Taylor 1962); however, untransformed means were shown in tables and figures. All statistical analyses were computed with SAS 9.1 statistical package, Carry, NC, USA (SAS 2002-2003). Figures were drawn with GraphPad Prism 5.0 software (GraphPad Software Inc 2007).

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6.3 Results

6.3.1 Experiment 1: Direct contact toxicity of spinosad to WFT and predatory mites

When exposed directly to spinosad, WFT adults experienced 98% mortality (at 96 h post- release exposure) and larvae experienced 96% mortality (at 72 h post-release exposure).

All species of predatory mites had greater than 90% mortality when exposed directly to spinosad, but mortality differed significantly amongst mite species (F 2,57 = 8.58, P = 0.0006), with the highest mortality recorded for H. miles (95.27% ± 2.71) and the lowest for N. cucumeris (90.25% ± 2.71). However, there was no significant difference (p > 0.05) in mortality between T. montdorensis (91.31% ± 2.04) and N. cucumeris. Spinosad toxicity declined over time. Mortality of mites was highest at 24 h post-release exposure period (Table 6.1). Mortality was lowest at 96 h post-release exposure period for T. montdorensis and H. miles and 72 h post-release exposure period for N. cucumeris. No mortality of N. cucumeris was recorded at 96 h post-release exposure period.

Table 6.1 Mean (±SE) corrected mortality (%) at different post-release exposure periods (h) to predatory mites after direct exposure to spinosad (recommended rate, 80 mL/100 L).

Post-release Corrected mortality (%) of predatory mite species (Mean ± SE) exposures period(h) T. montdorensis* N. cucumeris* H. miles* 6 6.00 ± 0.78b 6.55 ± 0.93b 9.23 ± 1.46c 24 60.35 ± 3.19d 62.33 ± 1.95d 68.32 ± 2.00d 48 15.25 ± 1.72c 14.90 ± 1.05c 11.41 ± 0.89c 72 6.46 ± 1.56b 6.47 ± 1.59b 5.56 ± 1.35b 96 3.25 ± 0.91a 0.0† 0.75 ± 0.41a F 106.09 204.28 256.40 df 4 , 95 3, 76 4 , 95 P <0.0001 <0.0001 <0.0001 * Within column values followed by the same letter do not differ significantly at α = 0.05. †No mortality of N. cucumeris at 96 h post-release exposure period was detected, and was therefore not included in analysis.

6.3.2 Experiment 2: Residual toxicity of spinosad to WFT and predatory mites

6.3.2.1 Residual (contact) toxicity of spinosad to WFT and predatory mites

Spinosad residues aged 2, 12, 24, 48, 72 and 96 h old were toxic to WFT adults (46 to 93% mortality), causing death until 96 h post-release exposure period (Table 6.2). The highest and

150 Chapter VI: Bioassay lowest mortality occurred when WFT adults were exposed to 2 h and 96 h old spinosad residues respectively (Table 6.2). Adult mortality was high and not significantly different (p > 0.05) among 2, 12, and 24 h old spinosad residues (Table 6.2). Similar to WFT adults, spinosad residues declined in toxicity to WFT larvae over time. The mean percentage of cumulative mortality to WFT larvae was highest and lowest when exposed to 2 h and 96 h old spinosad residues respectively (Table 6.2).

Table 6.2 Mean (± SE) corrected mortality (%) of WFT adults and larvae when exposed to spinosad residues aged 2, 12, 24, 48, 72, 96, and 120 h at different post-release exposure periods.

Corrected mortality (%) (Mean ± SE) Residue Post-release exposure periods (h) Cumulative age (h) mortality* 6h* 24h* 48h* 72h* 96h* Adults 2h 6.6±0.77c 55.8±2.9c 25.8±1.9a 3.8±0.95a 0.0±0.00a 92.9±1.58c 12h 4.8±0.7bc 52.2±2.9c 25.9±1.9a 4.5±1.14a 0.0±0.00a 89.6±1.72c 24h 2.8±0.75b 50.1±2.7c 27.5±2.1a 5.8±1.11a 0.5±0.34a 89.0±1.83c 48h 1.3±0.5a 31.0±1.6b 28.7±1.9a 7.5±1.18a 1.8±0.75b 71.3±3.32b 72h 0.0±0.0a 21.2±2.1a 24.4±1.9a 7.0±0.92a 0.8±0.4ab 54.2±3.06a 96h† 0.0±0.0a 18.4±1.9a 22.2±1.8a 5.5±0.88a 0.0±0.00a 46.8±2.49a ------F 25.17 47.93 1.45 1.91 3.36 66.88 df 5 & 114 5 & 114 5 & 114 5 & 114 5 & 114 5 & 114 P < 0.0001 < 0.0001 0.2144 0.0987 0.0072 < 0.0001 Larvae 2h 8.1±0.9c 63.6±3.0d 17.5±1.9b 4.8±0.9a - 94.9±2.7e 12h 7.1±0.9c 59.9±2.6d 22.0±2.1bc 4.5±1.1a - 94.7±1.1d 24h 4.1±0.7b 36.5±1.9c 29.7±2.1c 5.5±0.9ab - 79.6±2.6d 48h 1.5±0.6a 22.4±2.2b 27.9±2.1c 9.8±1.2b - 62.0±2.8c 72h 0.5±0.3a 11.7±1.1a 15.9±1.9ab 6.5±0.9ab - 35.2±2.3b 96h 0.0±0.0a 8.8±1.1a 9.8±1.6a 4.0±0.9a - 22.6±1.8a 120h† ------F 37.94 129.82 16.89 3.92 172.75 df 5 & 114 5 & 114 5 & 114 5 & 114 5 & 114 P < 0.0001 < 0.0001 < 0.0001 0.0026 < 0.0001 * Within column for each WFT stage, values followed by the same letter do not significantly differ at α = 0.05. †No mortality of adults and larvae was found for 120 h old spinosad residue.

Spinosad residues aged 2, 12, 24, 48, 72, 96 and 120 h old were toxic to mites, causing death until 72 h post-release exposure period (Table 6.3). Mortality due to residual contact toxicity of spinosad differed with residue age. Mortality was the highest and lowest for all three species when mites were exposed to 2 h and 144 h old residues respectively. There was no significant difference in mortality when T. montdorensis or N. cucumeris were exposed to 2 h and 12 h old residues. For H. miles, mortality did not differ between 2 h, 12 h and 24 h old residues.

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According to the IOBC toxicity classification, contact toxicity of spinosad ranged from harmless (<25% mortality) to harmful (>75% mortality) (Table 6.3). Spinosad residues aged 2 h to 96 h were slightly to moderately harmful to T. montdorensis at 72 h post-release exposure

Table 6.3 Residual toxicity (contact) of spinosad to predatory mites at 24, 48, and 72 h post release exposure periods. The IOBC classification: 1 = harmless (<25% mortality), 2 = slightly harmful (25-50% mortality), 3 = moderately harmful (51- 75% mortality) and 4 = harmful (>75% mortality). Persistence class: A = short-lived (<5 d), B = slightly persistent (5-15 d).

Residue Corrected mortality (%) (mean ± SE) at Toxicity class Persistence age (h) post release exposure period† class 24h* 48h* 72h* 24h 48h 72h T. montdorensis 2h 48.5±4.1g 57.34.6±5.1f 70.6±5.1f 2 3 3 A 12h 44.7±3.7fg 57.01±4.8ef 63.01±4.9ef 2 3 3 24h 39.9±2.6ef 56.0±4.5e 62.7±4.9e 2 3 3 48h 33.3±2.2e 49.7±4.6e 59.4±5.1e 2 2 3 72h 21.0±2.4d 38.0±3.8d 45.4±3.8d 1 2 2 96h 13.9±2.2c 31.1±3.3cd 33.7±3.6c 1 2 2 120h 4.6±0.5b 8.4±0.7b 9.2±0.9b 1 1 1 144h 0.0±0.0a 0.0±0.0a 0.8±0.4a 1 1 1 F 91.62 110.43 105.95 df 7, 152 7, 152 7, 152 P < 0.0001 < 0.0001 < 0.0001 N. cucumeris 2 44.4±3.2e 71.7±5.1g 76.0±4.9f 2 3 4 A 12 42.9±3.1e 70.2±4.9g 74.5±4.9f 2 3 3 24 29.8±1.9d 53.1±4.3f 58.2±4.4e 1 3 3 48 23.5±1.8c 41.1±2.8e 45.7±2.9d 1 2 2 72 18.2±1.82c 32.1±3.2d 36.2±3.2d 1 2 2 96 6.6±0.6b 13.5±0.9c 14.8±1.1c 1 1 1 120 0.8±0.4a 4.6±0.5b 4.6±0.5b 1 1 1 144 0.0±0.0a 0.5±0.3a 0.5±0.3a 1 1 1 F 174.95 128.75 145.63 df 7, 152 7, 152 7, 152 P < 0.0001 < 0.0001 < 0.0001 H. miles 2 55.5±3.6f 78.9±2.5f 82.34±2.5f 3 4 4 B 12 54.0±3.6f 75.3±3.0f 80.6±3.3f 3 4 4 24 52.34±1.9f 69.4±2.2f 76.45±2.6ef 3 3 4 48 46.7±3.9e 63.9±4.4e 73.49±4.6de 2 3 3 72 37.0±1.7d 55.2±2.4d 62.9±2.3d 2 3 3 96 17.2±1.7c 42.5±2.5c 44.6±2.8c 1 2 2 120 9.9±1.5b 17.3±1.6b 27.4±2.1b 1 1 2 144 2.4±0.7a 8.5±1.3a 10.5±1.7a 1 1 1 F 183.76 139.47 121.37 df 7, 152 7, 152 7, 152 P < 0.0001 < 0.0001 < 0.0001 †no mortality was recorded after 72 h post-release exposure period. *within column, for each species of predatory mites, means with different letters differed significantly (α = 0.05).

152 Chapter VI: Bioassay period. Meanwhile, spinosad residues aged 2 h to 72 h were slightly harmful to harmful to N. cucumeris at 72 h post-release exposure period. In case of H. miles, spinosad residues aged from 2 h to 120 h were slightly harmful to harmful at 72 h post-release exposure period. According to the IBOC guidelines, spinosad toxicity for T. montdorensis and N. cucumeris was short-lived, while slightly persistent for H. miles.

The relationships between mite mortality and spinosad residues, with mortality as a function of residue age (log hrs) are presented in Figure 6.2. The LT25 (25% mortality of non-target organisms caused by residual toxicity) indicates that the release of each species of predatory mite after a spinosad application would cause only 25% mortality, allowing 75% survival of the predatory mites. LT25 of spinosad differed for each species and were estimated as 4.2 days 2.01 1.89 2.143 (101.63 h; Antilog10 ), 3.2 days (77.72 h; Antilog10 ) and 5.8 days (138.83 h; Antilog10 ) for T. montdorensis, N. cucumeris and H. miles respectively (Table 6.4; Figure 6.2).

Table 6.4 Probit analysis (Abbott 1925) of the mortality of adult predatory mites exposed to spinosad residues of different ages.

LT 95% CL Spp Slope ± SE 25 χ2 df P (hrs) Lower Upper T. montdorensis -2.09 ± 0.078 101.63 91.34 114.39 16.11 6 0.013 N. cucumeris -2.11 ± 0.091 77.72 70.27 86.719 13.53 6 0.035 H. miles -3.18 ± 0.131 138.83 127.79 152.40 21.02 6 0.002

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1.0 A 0.8

0.6

0.4

0.2

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0.6

0.4

Probit mortality Probit 0.2

0.0 1.0 C 0.8

0.6

0.4

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0.0 0 1 2 3 4

Log10 (hrs)

Figure 6.2 Probit mortality of (A) T. montdorensis, (B) N. cucumeris, and (C) H. miles recorded against spinosad residues of different ages (log10 hrs).

6.3.3.2 Indirect exposure of spinosad to predatory mites via consumption of intoxicated WFT larvae

This bioassay evaluated mortality of predatory mites (T. montdorensis, N. cucumeris and H. miles) via consumption of intoxicated WFT larvae. WFT larvae were allowed to feed for 12 hours on strawberry leaves previously treated with spinosad (2, 12, 24, 48, 72, 96, 120 or 144 h old residues). The consumption of intoxicated WFT larvae killed predatory mites until 72 h post-release exposure periods as shown in Table 6.5. Mortality varied among residue ages and mortality was highest when mites fed on intoxicated WFT larvae that had fed on leaves with 2 h old residue. Meanwhile, mortality was lowest when T. montdorensis and N. cucumeris fed on intoxicated WFT larvae that had fed on 120 h old residue. No mortality of T. montdorensis or N.

154 Chapter VI: Bioassay cucumeris was recorded when mites fed on WFT larvae that had fed on 144 h old residue. In case of H. miles, mortality was lowest when mites fed on WFT larvae that had fed on 144 h old residue.

According to the IOBC classification, spinosad via indirect exposure was harmless to moderately harmful to predatory mites (Table 6.5). Spinosad residues aged 2 h to 24 h were slightly to moderately harmful to T. montdorensis at 24 h and 48 h post-release exposure periods, and moderately harmful at 72 h post-release exposure period. Meanwhile, 72 h to 120 h old residues of spinosad via indirect exposure were harmless to T. montdorensis. For N. cucumeris, indirect toxicity of 2 h and 12 h old residues was slightly harmful when examined at 24 h and 48 h post-release exposure periods and moderately harmful at 72 h post-release exposure period. Residues 24 h and 48 h old (at 72 h post-release exposure period) were only slightly toxic to N. cucumeris. For H. miles, 2 h, 12 h and 24 h old residues were moderately harmful, though 12 h and 24 h old residues were classified as slightly harmful when examined 24 h post-release exposure period. Forty-eight hour and 72 h old residues were slightly harmful for H. miles at 72 h post-release exposure period.

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Table 6.5 Mortality of predatory mites after feeding on spinosad intoxicated WFT larvae at 24 h, 48 h, and 72 h post-release exposure periods. The IOBC classification: 1 = harmless (<25% mortality), 2 = slightly harmful (25-50% mortality), 3 = moderately harmful (51- 75% mortality) and 4 = harmful (>75% mortality).

Corrected mortality (%) (mean ± SE) at post-release Residue Toxicity class exposure period† age (h) 24h* 48h* 72h* 24h 48h 72 T. montdorensis 2h 43.25±2.06f 45.23±2.72e 56.32±2.80e 2 2 3 12h 40.25±3.05f 42.24±3.39de 52.32±3.52e 2 2 3 24h 32.75±2.40e 40.35±3.35de 51.23±3.58e 2 2 3 48h 22.75±2.09d 37.38±2.13d 40.36±2.11d 1 2 2 72h 11.75±1.27c 22.75±2.07c 24.75±2.03c 1 1 1 96h 6.5±0.97b 10.75±1.27b 10.75±11.27b 1 1 1 120h 0.75±0.41a 3.75±0.71a 3.75±0.71a 1 1 1 144h†† ------F 115.73 140.98 1156.59 df 6, 133 6, 133 6, 133 P < 0.0001 < 0.0001 < 0.0001 N. cucumeris 2 44.27±2.41f 42.36±3.92e 51.23±3.64f 2 2 3 12 37.93±3.01f 40.36±3.91de 50.78.36±3.80f 2 2 3 24 30.07±1.92e 37.89±3.99de 40.93±3.99e 2 2 2 48 17.68±1.72d 30.20±2.67d 32.80±2.68d 1 1 2 72 11.91±1.73c 20.85±3.12c 22.74±3.36c 1 1 1 96 1.56±0.53b 8.16±0.83b 8.16±0.95b 1 1 1 120 0.71±0.34a 1.11±0.41a 1.11±0.41a 1 1 1 144†† ------F 164.90 167.00 183.16 df 6, 133 6, 133 6, 133 P < 0.0001 < 0.0001 < 0.0001 H. miles 2 50.14±3.23g 53.35±2.44e 62.34±2.11f 3 3 3 12 46.56±3.45fg 50.01±2.93e 59.99±3.28f 2 3 3 24 40.34±2.25ef 44.35±2.74de 55.45±2.91f 2 3 3 48 35.78±3.89e 37.25±4.42d 40.56±2.40e 2 2 2 72 20.76±2.60d 22.13±3.37c 34.20±3.67d 1 1 2 96 11.65±1.39c 20.13±2.05c 23.64±2.05c 1 1 1 120 5.06±0.62b 9.72±1.79b 9.72±1.79b 1 1 1 144 0.00±0.00a 0.95±0.39a 0.95±0.39a 1 1 1 F 218.32 192.998 201.76 df 6, 152 6, 152 6, 152 P < 0.0001 < 0.0001 < 0.0001 †no mortality was recorded after 72 h post-release exposure. ††no mortality of T. montdorensis and N. cucumeris was recorded for 144 h old spinosad residue. *Within column, for each species of predatory mites, means with different letters differed significantly (LS means, α = 0.05).

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6.3.5 Residual toxicity to predatory mites via consumption of spinosad-intoxicated WFT larvae and direct exposure to spinosad residues

Toxicity of spinosad residues via consumption of intoxicated WFT larvae and simultaneous exposures to residues is shown in Table 6.6. The mean mortality of T. montdorensis and N. cucumeris was highest when exposed to 2 h old residue and lowest when exposed to 144 h old residue. There was no difference in T. montdorensis or N. cucumeris mortality between 2 h and 12 h old residues. The mortality of H. miles was similarly highest when exposed to 2 h old residue and lowest when exposed to 144 h old residue. Hypoaspis miles mortality was high and not significantly different among 2 h, 12 h and 24 h old residues.

Indirect and contact toxicity of spinosad residues were classified as harmless to harmful (Table 6.6). Spinosad residues aged 2 h, 12 h and 24 h were harmful to T. montdorensis at 72 h post- release exposure period. Meanwhile, 48 h to 96 h old residues were slightly harmful to moderately harmful to T. montdorensis at 72 h post-release exposure period. For N. cucumeris, 2 h and 12 h old residues were harmful at 48 h and 72 h post-release exposure periods. Meanwhile, 24 h and 48 h old residues appeared to be moderately harmful for N. cucumeris at 72 h post-release exposure period. Spinosad residue aged 72 h was slightly harmful at 48 h and 72 post-release exposure periods only. Spinosad residues aged 96 h, 120 h and 144 h were classified harmless to N. cucumeris. For H. miles, 2 h, 12 h, and 24 h old residues were moderately harmful at 24 h post-release exposure period and became harmful when examined at 48 h and 72 h post-release exposure periods. Spinosad residues aged 48 and 72 h were moderately harmful to harmful for H. miles at 72 h post-release exposure period. Residues 96 h and 120 h old were classified as slightly harmful to H. miles.

As no harmful effects were found when T. montdorensis and N. cucumeris were exposed to residues aged 120 h, spinosad was categorised as short-lived (persistence <5 d) (Table 6.6). However, 120 h old spinosad residue posed some harmful effect to H. miles and was ranked as slightly persistent.

The relationships between mortality of the predatory mites and spinosad residues were analysed by probit procedure (Table 6.7; Figure 6.3) and mortalities as a function of residue ages (log hrs) are presented in Figure 6.3. The residual thresholds (LT25) were estimated as 5.4 d (129.67 2.11285 1.97816 2.16638 h, Antilog10 ), 3.9 d (95.09 h, Antilog10 ) and 6.1 d (146.68 h, Antilog10 ) for T. montdorensis, N. cucumeris and H. miles respectively.

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Table 6.6 Residual toxicity of spinosad to predatory mites at 24 h, 48 h, and 72 h post release exposure periods. Mites were fed spinosad-intoxicated WFT larvae and simultaneously exposed to residue. The IOBC classification: 1 = harmless (<25% mortality), 2 = slightly harmful (25- 50% mortality), 3 = moderately harmful (51- 75% mortality) and 4 = harmful (>75% mortality). Persistence class: A = short lived (<5 d), B = slightly persistence (5-15 d).

Corr. mortality (%) (mean ± SE) at post- Persist. Residue † Toxicity class treatment periods class age (h) 24h* 48h* 72h* 24h 48h 72h T. montdorensis 2h 51.39±2.04f 81.47±2.61f 87.56±2.84f 3 4 4 12h 50.63±2.47f 81.47±2.66f 86.55±2.73f 3 4 4 24h 42.82±2.93e 73.09±3.77ef 76.90±3.94f 2 3 4 48h 35.77±1.59d 70.05±2.68e 73.86±2.56e 2 3 3 72h 31.74±1.81d 48.98±3.11d 55.33±3.58d 2 2 3 A 96h 15.62±0.89c 28.68±1.07c 31.73±1.54c 1 2 2 120h 5.79±0.90v 9.13±1.09b 9.14±1.09b 1 1 1 144h 0.00±0.00a 3.27±0.40a 3.27±0.40a 1 1 1 F 240.55 288.40 287.23 df 6, 152 6, 152 6, 152 P < 0.0001 < 0.0001 < 0.0001 N. cucumeris 2 53.02±1.80g 80.97±2.65f 84.52±2.81f 3 4 4 12 45.23±2.91fg 76.14±3.81ef 80.46±3.70ef 2 4 4 24 39.70±1.45ef 67.01±2.69e 72.08±2.51e 1 3 3 48 31.16±1.63d 48.73±2.44d 53.30±2.57d 1 2 3 72 22.11±1.29c 43.15±1.86d 46.45±2.40d 1 2 2 A 96 10.55±0.86b 19.29±1.47c 20.56±1.57c 1 1 1 120 1.28±0.40a 5.33±0.66b 5.33±0.66b 1 1 1 144 0.00±0.00a 1.21±0.43a 1.21±0.43a 1 1 1 F 325.02 313.48 322.23 df 6, 152 6, 152 6, 152 P < 0.0001 < 0.0001 < 0.0001 H. miles 2 71.14±3.23f 87.98±2.44f 94.63±2.36f 3 4 4 12 70.13±2.11f 86.19±2.55f 91.05±2.84f 3 4 4 24 65.06±2.20f 83.89±1.97f 89.26±2.12ef 3 4 4 48 50.64±2.70e 70.84±3.19e 78.26±2.91e 3 3 4 72 38.48±1.53d 57.03±2.61d 63.68±2.72d 2 3 3 B 96 17.72±1.15c 43.73±1.44c 48.59±2.27c 1 2 2 120 10.88±1.13b 32.23±1.85b 32.99±1.99b 1 2 2 144 4.56±0.41a 11.76±0.90a 11.76±0.90a 1 1 1 F 257.59 200.28 211.91 df 6, 152 6, 152 6, 152 P < 0.0001 < 0.0001 < 0.0001 †no mortality was recorded after 72 h post-treatment in either of the species. *within column, for each species, means with different letters differed significantly (α = 0.05).

158 Chapter VI: Bioassay

Table 6.7 Probit analysis (Abbott 1925) of the mortality of adult predatory mites simultaneously exposed to spinosad via consumption of intoxicated WFT larvae and contact with spinosad residues of different ages.

95% CL LT Species Slope ± SE 25 χ2 df P (hrs) Lower Upper T. montdorensis -1.46±0.12 129.67 118.52 143.42 14.36 6 0.0258 N. cucumeris -1.43±0.10 95.09 87.17 104.61 12.35 6 0.0546 H. miles -1.81±0.15 146.68 135.91 159.89 15.68 6 0.0155

1.0

A 0.8

0.6

0.4

0.2

0.0 1.0 B 0.8

0.6

0.4

Probit mortality Probit 0.2

0.0 1.0 C 0.8

0.6

0.4

0.2

0.0 0 1 2 3 4

Log10 (hrs)

Figure 6.3 Probit mortality of (A) T. montdorensis, (B) N. cucumeris, and (C) H. miles recorded against spinosad residues of different ages (log10 hrs).

6.3.6 Experiment 3: Repellency of spinosad to predatory mites (choice test)

Results showed that T. montdorensis, N. cucumeris and H. miles were repelled by spinosad residues, though this repellence decreased with residue age (Table 6.8). There were more T.

159 Chapter VI: Bioassay montdorensis and H. miles on the water-treated strawberry leaves than leaves with 2 h, 12 h, 24 h and 48 h old spinosad residues. However, 72 h and 96 h old residues did not repel T. montdorensis or H. miles (Table 6.8). While, N. cucumeris were repelled by residues aged 2 h, 12 h and 24 h, but not by residues aged 48-96 h.

Table 6.8. Mean (±SE) numbers of predatory mites on spinosad- and water-treated strawberry leaf in a choice test (t-test, df = 19).

Residual Mean ±SE Species t-value P-value age (h) Control (Water) Spinosad 2 7.17 ± 0.16 1.87 ± 0.19 15.66 < 0.0001 12 7.13 ± 0.14 1.85 ± 0.16 18.72 < 0.0001 24 7.07 ± 0.16 1.97 ± 0.17 16.22 < 0.0001 T. montdorensis 48 5.97 ± 0.21 2.58 ± 0.22 8.83 < 0.0001 72 4.65 ± 0.09 4.40 ± 0.11 1.48 0.1556 96 4.62 ± 0.11 4.67 ± 0.15 -0.23 0.8242 2 6.98 ± 0.23 1.17 ± 0.16 15.74 < 0.0001 12 7.02 ± 0.18 1.13 ± 0.12 24.67 < 0.0001 24 6.45 ± 0.17 2.22 ± 0.18 14.90 < 0.0001 N. cucumeris 48 4.65 ± 0.09 4.48 ± 0.12 1.01 0.3248 72 4.72 ± 0.09 4.53 ± 0.11 1.21 0.2423 96 4.83 ± 0.08 4.75 ± 0.11 0.58 0.5663 2 7.40 ± 0.19 1.03 ± 0.13 24.78 < 0.0001 12 7.47 ± 0.19 0.95 ± 0.13 25.19 < 0.0001 24 7.52 ± 0.19 1.07 ± 0.15 25.17 < 0.0001 H. miles 48 5.73 ± 0.22 3.68 ± 0.22 4.98 < 0.0001 72 4.82 ± 0.17 4.48 ± 0.18 1.03 0.3171 96 4.46 ± 0.14 4.73 ± 0.16 -0.98 0.3394

6.4 Discussion

This laboratory bioassay confirms that spinosad is efficacious against WFT, causing 98% mortality of adults and 96% mortality of larvae when applied at the recommended rate (80 mL/100 L spinosad). Morishita (2001) similarly found that spinosad (25% WP) killed 100% WFT larvae in Petri dish bioassays. Apart from direct contact, spinosad residues also killed WFT adults and larvae, with 2 h old residue causing 100% mortality (Jones et al. 2005, van Driesche et al. 2006). The present study demonstrated that spinosad residues were highly toxic for longer: 6-48 hours under laboratory conditions. After 72 h, spinosad was either less or no longer toxic to WFT. Whilst it is difficult to extrapolate results from the laboratory to the field, there could also be a residual effect in the field.

Spinosad also affected T. montdorensis, N. cucumeris, and H. miles both directly and indirectly and choice tests indicate that spinosad residues are repellent to mites. Residues aged 2 h to 48 h

160 Chapter VI: Bioassay repelled T. montdorensis and H. miles, and residues aged 2 h to 24 h repelled N. cucumeris. Villanueva and Walgenbach (2006) reported that spinosad residues repelled two-spotted spider mite Tetranychus urticae Koch, but not European red mite, Panonychus ulmi (Koch). Tetranychus urticae females also congregated in untreated areas where they laid more eggs (Villanueva and Walgenbach 2006). While, van Driesche et al. (2006) found no significant difference in the numbers of N. cucumeris and the predatory mite Iphiseius degenerans (Berlese) on untreated or spinosad treated leaves, although they found relatively more mites on water-treated leaves.

Spinosad was highly toxic to T. montdorensis, N. cucumeris and H. miles when applied at the recommended rate (80 ml/100 L), causing >90% mortality. Elzen et al. (1998) using a bean-pod bioassay, found that spinosad killed >90% adult Orius insidiosus (Say) (Hemiptera: Anthocoridae). Spinosad residues were also toxic to mites and based on the age of the residue, the IOBC toxicity rating ranged from harmless to harmful. Spinosad residues aged 2 h to 72 h old were moderately to slightly harmful to N. cucumeris; residues aged 2 h to 96 h old were moderately to slightly harmful to T. montdorensis. Residues aged 2 h to 120 h old were harmful to slightly harmful to H. miles. This agrees with the data of Kongchuensin and Takafuji (2006) and Villanueva and Walgenbach (2005), who reported that spinosad residues were toxic to Neoseiulus longispinosus (Evans) and Neoseiulus fallacies (Garmen) (Acari: Phytoseiidae). Based on IOBC testing, Kopperts reports that spinosad is persistent to N. cucumeris (1-2 weeks) (Kopperts 2009), while Biobest reports that spinosad is not persistent to N. cucumeris (Biobest 2009). Contrary to the present findings, Jones et al. (2005) and van Driesche et al. (2006) reported that 2 h old spinosad residues were only slightly toxic to N. cucumeris. However, differences in results may be due to differences between N. cucumeris populations, such as previous exposure to spinosad. No spinosad has been applied in the insectaries in which the mites were reared (pers. comm., Chilman, Manchil IPM Services, WA).

The present bioassay also indicates that toxicity increased when mites simultaneously ate WFT larvae that had previously fed on spinosad-treated leaves and contacted to spinosad residues. For example, contact of T. montdorensis with 2 h old spinosad residue killed 70% adults, but mortality increased to 87% when T. montdorensis was also fed intoxicated WFT larvae. Villanueva and Walgenbach (2005) reported that adult Neoseiulus fallacies (Garman), a predator of two-spotted spider mite (T. urticae), died after feeding on T. urticae eggs exposed to spinosad. Thresholds for the residual (contact) toxicity of spinosad LT25 (lethal time for 25% mortality) were estimated as 4.2 days (101.63h), 3.2 days (77.72) and 5.8 days (138.83 h) for T. montdorensis, N. cucumeris and H. miles respectively. When mites fed on intoxicated WFT larvae and were simultaneously exposed to residues, toxicity further increased. Thresholds were

161 Chapter VI: Bioassay re-estimated as 5.4 d (129.67 h), 4 d (95.09 h), and 6.1 d (146.68 h) for T. montdorensis, N. cucumeris, and H. miles respectively. This suggests that T. montdorensis, N. cucumeris and H. miles are likely to survive if released six days after spinosad is applied. This is in general agreement with studies on other predatory mites. Kongchuensin and Takafuji (2006) stated there was no or very little negative influence of spinosad on adults, eggs or immature of N. longispinosus if exposed to spinosad seven days after application. Khan and Morse (2006) found a significant effect if the predatory mite, Euseius tularensis Congdon (Acari: Phytoseiidae) was released within five to six days of a spinosad application, but not if E. tularensis was released seven days after application. Residual thresholds were also higher for all three mite species when indirect exposure and contact toxicity was taken into account. Therefore, the interval between pesticide application and predatory mites release should be based on persistence and harmfulness of indirect and contact toxicity, not contact toxicity alone.

In conclusion, studies of the effects of pesticides on natural enemies often aim to assess the suitability of pesticides for IPM. To minimise any detrimental effects on non-target organisms, selectivity tests are performed with the aim of choosing pesticides with a high degree of lethal toxicity against the target pests and minimal non-target lethal toxicity. Although, this study was conducted under controlled conditions, it still provides valuable information on the likely trends of toxicity of spinosad residues. However, the sublethal effects of spinosad on these mites also need to be examined. Natural enemies subjected to multiple routes of exposure to pesticides may respond in unexpected ways that would be impossible to predict based on single route, laboratory toxicity tests (Kunkel et al. 2001). Side effects can include changes in behaviour, impact on fecundity and immature development (Desneux et al. 2007).

6.5 Literature cited

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Brødsgaard, H. F. 2004. Biological control of thrips-ornamentals, pp. 253-264. In K. M. Heinz, R. G. van Driesche and M. P. Parrella [eds.], Biocontrol in Protected Culture. Ball Publishing, Batavia, IL, USA. Chambers, W. S., and R. W. Sites. 1989. Overwintering thrips fauna in croplands of the Texas South plains. Southwestern Entomologist 14: 325-328. Cisneros, J., D. Goulson, L. C. Derwent, D. I. Penagos, O. Hernandez, and T. Williams. 2002. Toxic effects of spinosad on predatory insects. Biological Control 23: 2002. Cleveland, C. B., G. A. Bormett, D. G. Saunders, F. L. Powers, A. S. McGibbon, G. L. Reeves, L. Rutherford, and J. L. Balcer. 2002. Environmental fate of spinosad. 1. Dissipation and degradation in aqueous systems. Journal of Agricultural and Food Chemistry 50: 3244-3256. Cloyd, R. A., and C. S. Sadof. 2001. Effects of spinosad and acephate on western flower thrips inside and outside a greenhouse. Horticultural Technology 10: 359-362. Cote, K. W., P. B. Schultz, and E. E. Lewis. 2004. Using acaricides in combination with Phytoseiulus persimilis Athias-Henriot to suppress Tetranychus urticae Koch populations. Journal of Entomological Science 39: 267-274. Desneux, N., A. Decourtye, and J.-M. Delpuech. 2007. The sublethal effects of pesticides on beneficial arthropods. Annual Review of Entomology 52: 81-106. Eger Jr., J. E., J. Stavisky, and J. E. Funderburk. 1998. Comparative toxicity of spinosad to Frankliniella spp. (Thysanoptera: Thripidae), with notes on a bioassay technique. Florida Entomologist 81: 547-551. Elzen, G. W., P. J. Elzen, and E. G. King. 1998. Laboratory toxicity of insecticide residues to Orius insidiosus, Geocoris punctipes, Hippodamia convergens and Chrysoperla carnea. Southwestern Entomologist 23: 335-342. Finney, D. J. 1971. Probit Analysis. Cambridge University press, Cambridge. Gilkeson, L. A. 1990. New crops, new pests, new predators. Grower Talks August: 102-104. Gillespie, D. R. 1989. Biological control of thrips (Thysanoptera: Thripidae) on greenhouse cucumber by Amblyseius cucumeris. Entomophaga 34: 185-192. GraphPad Software Inc 2007. GraphPad Prism computer program, version 5.0. By GraphPad Software Inc. Hassan, S. A., F. Bigler, H. Bogenschütz, E. Boller, J. Brun, J. N. M. Calis, J. Coremans- Pelseneer, C. Duso, A. Grove, U. Heimbach, N. Helyer, H. Hokkanen, G. B. Lewis, F. Mansour, L. Moreth, L. Polgar, L. Samsoe-Petersen, B. Sauphanor, A. Staubli, G. Sterk, A. Vainio, M. van de Veire, G. Viggiani, and H. Vogt. 1994. Results of the sixth joint pesticide testing programme of the IOBC/WPRS-working group <>. Entomophaga 39: 107-119. Healy, M. J. R., and L. R. Taylor. 1962. Tables of Power - Law transformations. Biometrika 49: 557-559. Herron, G. A., and T. M. James. 2005. Monitoring insecticide resistance in Australian Frankliniella occidentalis Pergande (Thysanoptera: Thripidae) detects fipronil and spinosad resistance. Australian Journal of Entomology 44: 299-303. Holt, K. M., G. P. Opit, J. R. Nechols, and D. C. Margolies. 2006. Testing for non-target effects of spinosad on two-spotted spider mites and their predator Phytoseiulus persimilis under greenhouse conditions. Experimental and Applied Acarology 38: 141- 149. Jones, T., C. Scott-Dupree, R. Harris, L. Shipp, and B. Harris. 2005. The efficacy of spinosad against the western flower thrips, Frankliniella occidentalis, and its impact on associated biological control agents on greenhouse cucumbers in southern Ontario. Pest Management Science 61: 179-185. Khan, I., and J. G. Morse. 2006. Impact of citrus thrips chemical treatments on the predatory mite Euseius tularensis. Journal of Applied Entomology 130: 386-392. Kim, D. S., D. J. Brooks, and H. Riedl. 2006. Lethal and sublethal effects of abamectin, spinosad, methoxyfenozide and acetamiprid on the predaceous plant bug Deraeocoris brevis in the laboratory. Biocontrol 51: 465-484.

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Kongchuensin, M., and A. Takafuji. 2006. Effects of Some Pesticides on the Predatory Mite, Neoseiulus longispinosus (Evans) (Gamasina: Phytoseiidae). Journal of Acarology Society of Japan 15: 17-27. Kopperts. 2009. Side effects. Kopperts: Biological systems, The Netherlands. http://side- effects.koppert.nl/ download: 27.08.2009. Kunkel, B. A., D. W. Held, and D. A. Potter. 2001. Lethal and sublethal effects of bendiocarb, halofenozide and imidacloprid on Harpalus pennsylvanicus (Coleoptera: Carabidae) following different modes of exposure in turf grass. Journal of Economic Entomology 94: 60-67. Medina, P., F. Buda, L. Tirry, G. Smagghe, and E. Vinuela. 2001. Compatibility of spinosad, tebufenozide and azadirachtin with eggs and pupae of the predator Chrysoperla carnea (Stephens) under laboratory conditions. . Biocontrol Science and Technology 11: 597-610. Miles, M., and R. Dutton. 2000. Spinosad - a naturally derived insect control agent with potential for use in integrated pest management systems in greenhouses, pp. 339-344, Brighton Crop Protection Conference on Pests and Diseases. BCPC, Farnham, Surrey, UK. Morishita, M. 2001. Toxicity of some insecticides to larvae of Frankliniella occidentalis (Pergande) (Thysanoptera: Thripidae) evaluated by the petri dish-spraying tower method. Applied Entomology and Zoology 36: 137-141. Riudavets, J. 1995. Predators of Frankliniella occidentalis (Perg.) and Thrips tabaci Lind.: a review. Wageningen Agricultural University Papers 95: 43-87. Sabelis, M. W., and P. C. J. Van Rijn. 1997. Predation by Insects and Mites, pp. 259-354. In T. Lewis [ed.], Thrips as Crop Pests. CAB International, Wallingford, UK. SAS 2002-2003. SAS 9.1 computer program, version 9.1. By SAS, Cary, NC, USA. Shipp, J. L., and P. M. J. Ramakers. 2004. Biological control thrips on vegetable crops, pp. 265-276. In K. M. Heinz, R. G. van Driesche and M. P. Parrella [eds.], Biocontrol in protected culture. Ball Publishing, Batavia, Illinois. Shipp, J. L., K. Wang, and G. Ferguson. 2000. Residual Toxicity of Avermectin b1 and Pyridaben to Eight Commercially Produced Beneficial Arthropod Species Used for Control of Greenhouse Pests. Biological Control 17: 125-131. Sparks, T. C., G. D. Thomson, H. A. Kirst, M. B. Hertlein, L. L. Larson, T. V. Worden, and S. T. Thibault. 1998. Biological activity of the spinosyn, new fermentation derived insect control agents, on tobacco budworm (Lepidoptera: Noctuidae) larvae. Journal of Economic Entomology 91: 1277-1283. Steiner, M., and S. Goodwin. 2000. Development and evaluation of usage protocols for newly developed western flower thrips biocontrol agents, pp. 15-25. In D. Cook [ed.], National strategy for the management of western flower thrips and tomato spotted wilt virus. Department of Agriculture Western Australia, South Perth. Steiner, M. Y., S. Goodwin, T. M. Wellham, I. M. Barchia, and L. J. Spohr. 2003. Biological studies of the Australian predatory mite Typhlodromips montdorensis (Schicha) (Acari: Phytoseiidae), a potential biocontrol agent for western flower thrips, Frankliniella occidentalis (Pergande) (Thysanoptera: Thripidae). Australian Journal of Entomology 42: 124-130. Sterk, G., S. A. Hassan, M. Baillod, F. Bakker, F. Bigler, S. Blümel, H. Bogenschütz, E. Boller, B. Bromand, J. Brun, J. N. M. Calis, J. Coremans-Pelseneer, C. Duso, A. Garrido, A. Grove, U. Heimbach, H. Hokkanen, J. Jacas, G. Lewis, L. Moreth, L. Polgar, L. Rovestl, L. Samsoe-Petersen, B. Sauphanor, L. Schaub, A. Stăubli, J. J. Tuset, A. Vainio, M. van de Veire, G. Viggiani, E. Viñuela, and H. Vogt. 1999. Results of the seventh joint pesticide testing programme carried out by the IOBC/WPRS working group pesticides and beneficials. Biocontrol 44: 99-117. Thompson, D. G., B. J. Harris, L. J. Lanteigne, T. M. Buscarini, and D. T. Chartrand. 2002. Fate of spinosad in litter and soils of a mixed conifer stand in the Acacian forest region of New Brunswick. Journal of Agricultural and Food Chemistry 50: 790-795. Thompson, G., and S. Hutchins. 1999. Spinosad. Pesticide Outlook 10: 78-81.

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Thompson, G. D., R. Dutton, and T. C. Sparks. 2000. Spinosad-a case study: an example from natural products discovery programme. Pest Management Science 56: 696-702. van Driesche, R. G., S. Lyon, and C. Nunn. 2006. Compatibility of spinosad with predacious mites (Acari: Phytoseiidae) used to control western flower thrips (Thysanoptera: Thripidae) in greenhouse crops. Florida Entomologist 89: 396-401. Villanueva, R. T., and J. F. Walgenbach. 2005. Development, oviposition, and mortality of Neoseiulus fallacis (Acari: Phytoseiidae) in response to reduced-risk Insecticides. Journal of Economic Entomology 98: 2114-2120. Villanueva, R. T., and J. F. Walgenbach. 2006. Acaricidal Properties of Spinosad Against Tetranychus urticae and Panonychus ulmi (Acari: Tetranychidae). Journal of Economic Entomology 99: 843-849. Viñuela, E., M. P. Medina, M. Schneider, M. González, F. Budia, A. Adán, and P. Delestal. 2001. Comparison of side effects of spinosad, tebufenozide, and azadirachtin on the predators Chrysoperla carnea and Podisus maculiventris and the parasitoids Orius concolor and Hyposoter didymator under laboratory conditions. Bulletin IOBC/WPRS 24: 25-34. Zhao, J. Z., Y. X. Li, H. L. Collins, L. Gusukuma-Minuto, R. F. L. Mau, G. D. Thompson, and A. M. Shelton. 2002. Monitoring and characterization of diamondback moth (Lepidoptera: Plutellidae) resistance to spinosad. Journal of Economic Entomology 95: 430-436. Zilahl-Balogh, G. M. G., J. L. Shipp, C. Cloutier, and J. Brodeur. 2007. Predation by Neoseiulus cucumeris on western flower thrips, and its oviposition on greenhouse cucumber under winter vs. summer conditions in a temperate climate. Biological Control 40: 160-167.

165 CHAPTER VII

Spinosad-resistant western flower thrips, Frankliniella occidentalis (Pergande) (Thysanoptera: Thripidae) can be managed using spinosad and predatory mites (Acari)

Keywords: Frankliniella occidentalis, Typhlodromips montdorensis, Neoseiulus cucumeris,

Hypoaspis miles, resistance, spinosad, residual toxicity, LT25

Abstract

Western flower thrips, Frankliniella occidentalis (Pergande) (Thysanoptera: Thripidae) is a serious pest on a wide range of crops and has developed resistance to one or more insecticides. In Australia, F. occidentalis has developed resistance to the biopesticide, spinosad. To control spinosad-resistant F. occidentalis, growers could double the recommended application rate (80 mL/100 L to 160 mL/100 L). However, increasing the rate could have a detrimental effect on predatory mites (Acari) which are used as biological control agents in an integrated pest management (IPM) approach. This study assessed the effects of applying spinosad (Success™, Dow AgroSciences, Australia) at twice the recommended rate to spinosad-resistant F. occidentalis and to the predatory mites, Typhlodromips montdorensis (Schicha) (Phytoseiidae), Neoseiulus cucumeris (Oudemans) (Phytoseiidae) and Hypoaspis miles (Berlese) (Laelapidae). Direct exposure to twice the recommended rate of spinosad killed 100% of all mite species. Spinosad residues aged two, 24, and 48 h were also highly toxic to all three mite species, causing 96-100% mortality. The persistence of spinosad was rated as short-lived for N. cucumeris, and slightly persistent for T. montdorensis and H. miles. Comparative toxicity indicates that spinosad residues aged 48 to 168 h were less toxic to N. cucumeris followed by T. montdorensis and H. miles. The residual thresholds (LT25) of twice the recommended rate of spinosad for T. montdorensis, N. cucumeris, and H. miles were calculated as 6.1, 5.3, and 6.8 days respectively. By maintaining an interval, of 6-7 days between spinosad application at twice the recommended rate and mite release, F. occidentalis can be effectively controlled. Typhlodromips montdorensis appears to be the most successful species in reducing thrips numbers followed by N. cucumeris and H. miles. Information that could be included in resistance management of F. occidentalis is discussed.

7.1 Introduction

During the last few decades, many arthropod species have developed resistance to a number of insecticides and insecticide classes (Georghiou 1990). This presents a serious challenge to food Chapter VII: Resistance management production when chemical control is the primary tactic used to manage pest populations. Development of a new pesticide takes a significant investment and may not be developed quickly enough to benefit growers. Thus, strategies must be adopted to prolong the effectiveness of current insecticides while new technologies are being developed. Western flower thrips (WFT), Frankliniella occidentalis (Pergande) (Thysanoptera: Thripidae) is a worldwide pest of economic importance (Lewis 1998), infesting a wide range of crops including ornamentals, fruit, and vegetable crops (Tommasini and Maini 1995). Chemical control is the principal management strategy for WFT in many parts of the world (Contreras et al. 2001), including Australia (Herron and Gullick 2001, Herron and James 2005, Herron et al. 2007). However, WFT has developed resistance to several insecticides (Brødsgaard 1994, Broadbent and Pree 1997, Jensen 2000, Espinosa et al. 2002, Herron and James 2005), including newer pesticides such as imidacloprid and amitraz (Zhao et al. 1994, Zhao et al. 1995). In Australia, WFT control relies on a limited number of pesticides (abamectin, acephate, chlorpyrifos, dichlorvos, dimethoate, endosulfan, fipronil, malathion, methamidophos methidathion, methiocarb, methomyl, pyrazophos and spinosad) (Herron and James 2005). However, high levels of resistance to pyrethroids (cypermethrin, bifenthin, deltamethrin and fluvalinate) has been detected and pyrethroids are no longer recommended for use against WFT (Herron and Gullick 2001). WFT also developed resistance to spinosad in Australia (Herron and James 2005) and other parts of the world (Bielza et al. 2007). Spinosad is considered to be not toxic or less toxic on beneficials and is often included in pest management programs that incorporate biological control (Miles et al. 2003, Anh et al. 2004, Kim et al. 2005). Although spinosad resistance has been detected in Australian populations, it may be possible to continue to use this pesticide in conjunction with biological control (Jensen 2000).

In Australia, three species of predatory mites, Typhlodromips montdorensis (Schicha) (Phytoseiidae), Neoseiulus cucumeris (Oudemans) (Phytoseiidae) and Hypoaspis miles (Berlese) (Laelapidae) are commercially available for the management of WFT. Susceptibility of these mites to spinosad has not yet been evaluated. However, it is expected that spinosad application will reduce mite populations (Williams et al. 2003). For example, Kongchuensin and Takafuji (2006) reported fresh spinosad residues (up to 48 h old) posed significant detrimental effects on eggs immature stages of the predatory mite Neoseiulus longispinosus (Evans). van Driesche et al. (2006) reported that fresh residues of spinosad applied at the recommended rate for WFT control on glasshouse flower crops can lower the survival of Iphiseius degenerans (Berlese). Spinosad is reported to be highly toxic to Neoseiulus fallacis (Garman) used in North Carolina apple orchards to control pest mites (Panonychus ulmi Koch and Tetranychus urticae Koch) (Villanueva and Walgenbach 2005).

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Given the current high reliance on spinosad for WFT control and the increasing probability that resistance will spread, it is important to explore ways in which spinosad can be used with biological control agents effectively. One approach is to use an initial high dose of an insecticide to reduce the resistant population of the pest, and thereafter release natural enemies to maintain the population below the economic threshold (Tabashnik and Croft 1982). However, the use of a high dose of an insecticide is likely to increase the detrimental effect on natural enemies, particularly if there is any residual activity. Therefore, it is important to evaluate the threshold period of an insecticide for natural enemies when there is a need to increase the dose. This study investigated the possibility to integrate chemical control (high dose) and biological control for the management of a spinosad-resistant WFT population. The objectives of this study were to evaluate: (i) the residual threshold of high dose spinosad for T. montdorensis, N. cucumeris, and H. miles, and (ii) the effectiveness of T. montdorensis, N. cucumeris, and H. miles with a higher rate of spinosad to manage spinosad-resistant WFT.

7.2 Materials and methods

Trials were conducted in a controlled temperature (CT) room (25±1⁰C, 50-60% RH, 16:8 h L: D regime) from October 2008 to January 2009 at the University of Western Australia (UWA).

7.2.1 Source cultures

7.2.1.1 Strawberry plants

Strawberry runners [Fragaria x ananassa Duchesne (Rosaceae)] cultivar Camino Real were planted into pots (32.5 l x 32.5 w x 40.5 h cm) containing potting mix (Baileys Fertilisers, Rockingham, WA) in glasshouses at the Department of Agriculture and Food WA (DAFWA), South Perth and UWA, Crawley. All pots were fitted with sprinklers with automatic timers. During summer, the plants were watered every third day, while in winter and spring plants were watered once a week. A liquid fertiliser (Thrive ® Yates, Australia; NPK: 12.4: 3: 6.2; rate: 5mL/2 L water) was applied once a month.

7.2.1.2 Western flower thrips (WFT)

A glasshouse colony of WFT was established from individuals initially collected from calendula flowers [Calendula officianalis L. (Asterales: Asteraceae)] from glasshouses at DAFWA. WFT colonies were maintained on potted calendula from July 2006 at UWA. Potted calendula was grown from seeds collected from calendula plants.

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Calendula seeds were sown in plastic pots (50x100 mm) with potting mix (Baileys Fertilisers, Rockingham, WA) and plants were kept in insect proof Perspex cage (500 mm high, 420 mm deep and 400 mm wide), fitted with 105 µ mesh net (Sefar Filter Specialists Pty Ltd., Malaga, WA) fitted on a Nylex tote box (320 x 420 mm; Blyth Enterprises Pty Ltd, Australia). To maintain the WFT colony, every fortnight WFT was collected from caged calendula plants and released onto fresh plants.

Spinosad-resistant WFT used in this study were obtained from NSW Agriculture, which was initially collected from hydroponic lettuce in the Sydney basin, NSW (S. Broughton, pers. comm.). The colony was maintained on calendula in insect-proof Perspex cages as described above from July 2007 at UWA.

7.2.1.3 Predatory mites

Predatory mites [T. montdorensis, N. cucumeris and H. miles] were sourced from commercial Australian suppliers (Biological Services, SA; Chilman IPM Services, WA; and Beneficial Bug Company, NSW). Mites were provided in plastic buckets containing vermiculite. Trials were conducted immediately upon receipt of mites.

7.2.2 Experiment 1: Direct toxicity of spinosad

7.2.21 Western flower thrips

Toxicity of spinosad (Success™, 120 g/L EC, Dow AgroSciences Australia Ltd) at (i) the recommended rate (80mL/100 L, 0.096 g a.i./L), (ii) twice the recommended rate and (iii) three times the recommended rate to spinosad-resistant WFT adults and larvae were evaluated. Twenty-four hours prior to the trial, WFT adults (5-6 d old) from the resistant strain were collected from calendula plants with an aspirator, and brought to the CT room. WFT adults were transferred to a glass Petri dish (150 x 15 mm) containing a strawberry leaf. The top of the Petri dish was covered with mesh net (105 µ) and sealed with paraffin film (Parafilm M®, Micro Analytix Pty Ltd)] and kept for 24 h for acclimatisation to the experimental arena. At the onset of the experiment, 20 cold-anaesthetised WFT adults were placed on a paper towel and lightly sprayed once with either 5mL of spinosad solution (diluted in distilled water) at one of the above rates or distilled water only (control) with a hand-held atomiser (Hills Sprayers, BH220063). After spraying, excess liquid (if any) was gently removed with a soft tissue. WFT adults (n = 20) were then transferred to an excised strawberry leaf (adaxial side up) on a moistened filter paper in a single Petri dish (150 x 15 mm). The leaf petiole was covered with

169 Chapter VII: Resistance management cotton was soaked in a 10% sugar solution to extend the leaf life. The leaf edge was glued to the filter paper so that WFT could not go underneath the leaf. The top of the Petri dish was covered with a mesh net (105 µ) sealed with a paraffin film to prevent thrips escaping. There were 20 replicates (20 x 20 = 400 individuals) of each treatment and a control. The experiment was repeated using the same protocol as above but with WFT larvae (first instar) instead of adults.

All Petri dishes were then placed on a laboratory bench in a random block design in the CT room (25±1⁰C, 50-60% RH and 16:8 h light-dark cycle) and thrips were monitored for mortality. Spinosad is a slow-acting insecticide (Bret et al. 1997) and cumulative mortality of a test organism usually plateaus at 2 days (48 h) to 6 days (144 h) after exposure (Viñuela et al. 2001, Cisneros et al. 2002). Consequently, Petri dishes with WFT adults were examined at 6, 24, 48, 72, and 96 h post-release exposure periods under a stereomicroscope. The WFT larvae were checked at 6, 24, 48 and 72 h post treatments (after 72 h post-release exposure period those larvae were lived had pupated). WFT adults or larvae were recorded as dead if they did not respond to probing with a fine paintbrush.

7.2.2.2 Predatory mites

Bioassay methods were the same as for thrips, except that a thin barrier of Tac Gel (Stickem™, The Olive Centre, Australia) was applied to the edge of the leaf to keep mites on the leaf surface. In addition, first or second instar WFT larvae were added to the Petri dishes to provide food for the mites. In each Petri dish, there were 200 larvae for T. montdorensis [average 10 first instar larvae/ mite (Steiner et al. 2003)], 100 larvae for N. cucumeris [average six fist instar larvae/ mite (Zilahl-Balogh et al. 2007) and 40 larvae for H. miles [two second instar larvae/ mite (Berndt et al. 2004)]. During the trial period, additional thrips larvae were added to the Petri dishes as required. Mortality of the mites was monitored at 6 h, 24 h, 48 h, 72 and 96 h post-release exposure periods. Mites were recorded as dead if they did not respond to probing with a fine paintbrush. Each treatment (spray) was replicated 20 times (20 x 20 = 400 individuals) for each species of predatory mite.

7.2.3 Experiment 2: Bioassay of spinosad residual toxicity to predatory mites

This bioassay evaluated the mortality of predatory mites (T. montdorensis, N. cucumeris and H. miles) via consumption of intoxicated WFT larvae and simultaneous exposure (contact) to spinosad residues. Predatory mites were placed on strawberry leaves that had been treated with spinosad solution at twice the recommended rate (80 mL/100 L), and were allowed to feed on intoxicated WFT larvae that had been exposed to the same aged residue. Twenty-four hours

170 Chapter VII: Resistance management prior to the trial, newly emerged first instar WFT larvae were collected from the stock colony. Larvae were transferred to a glass Petri dish (150 x 15 mm) containing a cotton wool ball soaked in a 10% sugar solution, and then stored in the CT room for 12 h (to ensure feeding on spinosad- or water-treated leaf at a later stage). Larvae were then released onto excised strawberry leaves previously treated with spinosad (2, 24, 48, 72, 96, 120, 144 or 168 h old residues) or control (water) in a Petri dish and allowed to feed for 12 hours. A thin barrier of Tac Gel was applied to the edge of the leaf to prevent larvae from escaping the leaf. The dose of twice the recommended rate of spinosad was chosen for testing residual toxicity on predatory mites in this experiment, because, in the previous experiment, this dose appeared effective in controlling 50% or more of the spinosad-resistant WFT population. Prior to spraying, potted strawberry plants Camino Real (3-4 weeks old) were brought to the CT room and split into groups (2, 24, 48, 72, 96, 120, 144 and 168 h old residue). To obtain the required residues available for experimentation at the same time, strawberry plants were sprayed at different times with spinosad solution until run-off with a hand-held atomiser. Those plants in the 168 h old residue treatment were sprayed first, followed by spray of the subsequent group of plants. In a similar manner, a second set of plants was sprayed with spinosad solution by maintaining 12 h interval from the time when the first set of plants was sprayed for each treatment (residue ages). Strawberry plants in the control group were sprayed with distilled water 24 h prior to the experiment. To avoid contamination, separate atomisers were used for the water and spinosad. Leaves were marked with a permanent marker at the base of the petiole, to enable leaves with residues to be accurately selected. After spraying, plants were covered with a modified thrips- cage (45 x 35 cm, open both ends) made from mesh net (105µ) and supported by quadratic steel-rod stands. The bottom end of the cage was taped to the pot. The top end of the cage was closed with a rubber band. All treated plants were then returned to a glasshouse for natural degradation of spinosad and were kept until use.

Spinosad was also applied to the Petri dishes (150 x 15 mm; testing arena) at the same time as it was sprayed on the plants in order to obtain strawberry leaves and dishes of the same exposure age. The Petri dishes were held with the open end sidewise and sprayed inner on the side with spinosad solution using an atomiser until run-off. Both parts of a Petri dish were sprayed with spinosad. Petri dishes were kept on a tray and allowed to dry in a glasshouse for one and half hours. Dried Petri dishes were stored in separate plastic trays and kept in the CT room until use. The Petri dishes were sprayed with distilled water to be used as the untreated control.

At the beginning of the experiment, the first set of spinosad treated plants was brought to the CT room. A strawberry leaf excised from a treated plant was placed adaxial side top at the bottom of a glass Petri dish. A thin barrier of Tac Gel was applied to the edge of the leaf to prevent

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WFT larvae escape from the leaf surface. First instar WFT larvae collected from the colony were released onto the treated leaf in the Petri dish and allowed to feed for approximately 12 hrs. This was done to get the intoxicated WFT larvae. Twelve hours later, a second set of spinosad treated strawberry plants [plants sprayed by maintaining a 12 h interval of each respective residue age group] were brought to the CT room. Testing arenas were prepared using the same age residue of leaf, Petri dish and WFT larvae as above. Intoxicated first instar WFT larvae [the number of WFT release per Petri dish was the same as mentioned in section 7.2.2.2) were released first onto the leaf, followed by the release of predatory mites. During the trial periods, additional intoxicated WFT larvae with same residue age were added to the Petri dish if required. Twenty adult mites of one species were placed on the leaf surface in a Petri dish covered with a mesh net (105µ) and sealed with paraffin film. The leaf petiole was covered with cotton was soaked in a 10% sugar solution to keep the leaf fresh and extend the life. Before releasing WFT larvae or predatory mites, a thin barrier of Tac Gel was applied to the edge of the leaf surface to prevent escape of WFT larvae and predatory mites. There were 20 replicates (20 x 20 individuals) of each treatment (residue ages). The experiment was repeated using the same protocol above but with two other species of predatory mites. The Petri dish and strawberry leaf were sprayed with distilled water to be used as the control (20 x 20 individuals). All Petri dishes with predatory mites were arranged randomly on a laboratory bench and mortality was checked in the same manner as explained previously for WFT. Spinosad is slow-acting (Bret et al. 1997) and cumulative mortality of a test organism usually plateaus at two days (48 h) to six days (144 h) after exposure (Viñuela et al. 2001, Cisneros et al. 2002). Predatory mites were recorded as dead if they did not respond to probing with a paintbrush.

7.2.4 Experiment 3: Efficacy of predatory mites with spinosad against WFT-resistant strain

This experiment evaluated the efficacy of predatory mites when combined with twice the recommended rate of spinosad (80ml/100 L) against spinosad-resistant WFT strain. In order to obtain uniformly aged WFT adults for this trial, ten adults were collected from the resistant colony, then released onto a fresh (not previously exposed to any insects including WFT) calendula plant in insect-proof Perspex cages as described in 7.2.1. WFT adults were allowed to lay eggs for 24 h. After 24 h, all adults were removed from the plants with an aspirator. Plants were checked daily for larval emergence. Adults that emerged on the same day were used in this trial.

Prior to the trial, WFT adults (spinosad-resistant) were collected from the stock colony, brought to the CT room and released onto freshly caged strawberry plants. Strawberry plants with WFT

172 Chapter VII: Resistance management adults were kept in the CT room for 24 h to acclimatise with the experimental environment. At the onset of the trial, 40 strawberry plants (3-4 leaves stage) cultivar Camino Real, were brought to the CT room. Plants were randomly divided into two groups and sprayed with twice the recommended rate of spinosad or distilled water (control) with a hand-held atomiser until run- off. Plants were then covered with a modified thrips cage (45 x 35 cm) made with mesh net (105µ) and supported by a quadrate steel-rod stand. The treated plants were allowed to dry for two hours. Previously acclimatised spinosad-resistant WFT adults of the same age were then released onto the plants. Fifteen WFT adults per plant were released. Plants with WFT were further divided into four groups and received:

(i) No mites (ii) six T. montdorensis (iii) six N. cucumeris (iv) six H. miles.

Based on the residual threshold (estimated from the previous experiment), , N. cucumeris, T. montdorensis or H. miles were released on to plants at six, five or seven days after spinosad application respectively. Pots were randomly arranged on a laboratory bench in the CT room. There were five plants (pot = replicate) per treatment (mite species). Twenty-four hours after WFT release, the plants were checked with a battery-powered, hand-held magnifying glass [50 mm (2") illuminated round 2x power with 4x bifocal magnifier] for live WFT. Thereafter, every fifth day for five weeks, plants were checked and the number of live WFT adults and larvae per plant were recorded. Plants were checked early in the morning (0600 to 0800 h) as WFT was found to be less active at this time. Plants were watered as required.

7.2.5 Data analysis

For each experiment in which mortality was assessed, the number of individuals that died at each observation was counted and expressed as a percentage of the total number of individuals in the arena and corrected using Abbott’s formula (Abbott 1925).

Abbott’s formula takes into account the proportion of control thrips or mites dying in the trial that have not been exposed to the spinosad, and amongst those that have been exposed to spinosad, some may die of natural causes. In these trials, control mortality of either thrips or mites never exceeded 5%.

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The differences in corrected cumulative mortality (mortality after 96 h post-release exposure period) of each residue ages among predatory mites were analysed by one-way ANOVA (Proc Mixed Procedure), except cumulative mortality due to two and 24 h old residues, as these two residues caused 100% mortality of all three species of predatory mites. To determine the difference in toxicity amongst residue ages, corrected mortality of T. montdorensis, N. cucumeris, and H. miles at each period post-release exposure period was subjected to one-way ANOVA separately. Mortality data were transformed using arcsine transformation (Zar 1999) (Healy and Taylor 1962) to normalise before analysis, though actual means are reported. If ANOVA results showed significant differences, mortality means were separated by least square mean differences at 5% probability.

The residual toxicity of spinosad to T. montdorensis, N. cucumeris, and H. miles at each post- release exposure period was classified following International Organization of Biological Control (IOBC) guidelines:

1 = harmless (<25% mortality) 2 = slightly harmful (25-50% mortality) 3 = moderately harmful (51- 75% mortality) 4 = harmful (>75% mortality) (Sterk et al. 1999).

The persistence of spinosad for each species of predatory mites was also classified according to the time taken to lose toxicity (<30% mortality, IOBC persistence class):

A = short-lived (<5 d) B = slightly persistent (5-15 d) (Hassan et al. 1994, Sterk et al. 1999).

The residual toxicity threshold of spinosad for T. montdorensis, N. cucumeris and H. miles was estimated with Probit analyses (Finney 1971) by Proc Probit Procedure. The LT25 (lethal time of 25% mortality) was used, which is considered an acceptable level (Shipp et al. 2000). The effect of spinosad and predatory mite releases on the numbers of the spinosad-resistant WFT strain (adults and larvae) were compared by two-way ANOVA (Proc Mixed Procedure) (independent variables: spray treatment and predatory mite releases; response variables: WFT adults or larvae) separately at each observation, 10 to 35 DAS (days after spray). Since there was a significant interaction between spray (spinosad and water) and predatory mite releases (no mites, T. montdorensis, N. cucumeris and H. miles), additional one-way ANOVAs were performed (Quinn and Keough 2002). WFT numbers (adults or larvae) at each observation period were analysed with two separate one-way ANOVAs (one for each spray treatment). Due

174 Chapter VII: Resistance management to multiple comparisons, an adjustment to the significance level was made, α = 0.025 (0.05/2). Meanwhile, to determine the difference in WFT adults and larvae between groups before mite release, WFT adults and larval data for spinosad and water treatment of one and five DAS were analysed by one-way ANOVAs (Proc Mixed Procedure). If ANOVA results were significant, means were separated with least square mean differences. Data were transformed using √(x + 0.05) (Healy and Taylor 1962) to normalise before analysis. Data were reversed transformed for presentation.

All analyses were computed using SAS 9.1, SAS Institute, 2003, Cary, NC, USA (SAS 2002- 2003). Figures were drawn with GraphPad Prism 5.0 software (GraphPad Software Inc 2007).

7.3 Results

7.3.1 Direct toxicity of spinosad to WFT and predatory mites

Bioassays confirmed that the recommended rate of spinosad was not more toxic to adults or larvae of the spinosad-resistant strain of WFT used in this study, than the distilled water spray (control) (Table 7.1). Doubling the recommended rate killed >70% of WFT adults and larvae (Table 7.1). Meanwhile, direct exposure to triple the recommended rate of spinosad killed 100% of WFT adults and larvae. In the water treatment, 0.5% WFT adults were killed, whereas no mortality of WFT larvae was recorded in the water treatment.

As it was found that double of the recommended rate of spinosad could be effective against spinosad-resistant WFT population, toxicity of double the recommended rate of spinosad was tested against T. montdorensis, N. cucumeris and H. miles. Direct exposure to twice the recommended rate was very toxic to predatory mites T. montdorensis, N. cucumeris, and H. miles, resulting in 100% mortality. In this trial, in the water treatment, mite mortality never exceeded 5% (1.7%, 2.0% and 3.25% of T. montdorensis, N. cucumeris and H. miles were killed respectively).

Table 7.1 Cumulative corrected mortality (%) of spinosad- resistant WFT adults (at 96 h post- release exposure period) and larvae (at 72 h post-release-exposure period) when exposed directly to spinosad spray at different rates.

Corrected mortality (%) of spinosad-resistant Spinosad application rate WFT (Mean ± SE) Adults Larvae Recommended rate (80 ml/100 L) 0.50 ± 0.46 0.00 Double the recommended rate 87.94 ± 2.36 78.57 ± 3.45 Triple the recommended rate 100 100

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7.3.2 Bioassay of spinosad residual toxicity to predatory mites

Spinosad residues of all ages of double the recommended rate were toxic to some degree to T. montdorensis, N. cucumeris and H. miles (Figure 7.1), although toxicity declined as the residual period increased. Spinosad residues aged 2 h and 24 h were very toxic to T. montdorensis, N. cucumeris and H. miles and caused 100% mortality within 24 h of exposure. Cumulative mortality of 48 h old residue at 96 h post-release exposure period was not different (F 2, 57 = 3.01, P = 0.057) among predatory mite species and was close to 100%. However, each mite species experienced different mortality when exposed to 72-168 h old residues (72 h: F 2, 57 =

35.08, P < 0.0001; 96 h: F 2, 57 = 5.64, P = 0.0058; 120 h: F 2, 57 = 9.61, P 0.0002; 144 h: F 2, 57

= 25.41, P < 0.0001; 168 h: F 2, 57 = 35.43, P < 0.0001) old residues at 96 h post-release exposure period (Figure 7.1). For each residue age (72 h to 168 h), H. miles mortality was the highest and N. cucumeris mortality was the lowest (Figure 7.1). However, mortality of T. montdorensis and N. cucumeris was not different when exposed to 72 h old residue. Similarly, when exposed to 96 h old residue, mortality of T. montdorensis and H. miles was not different. In the control treatment, less than 5% (1.5%, 2% and 2.5% mortality of T. montdorensis, N. cucumeris and H. miles respectively) mortality was found in the predatory mite species.

T. montdorensis N. cucumeris H. miles

100 a a a b SE)  80 a a b 60 b a

40 c b c a b c 20 a b a

Corrected mortality (%) (Mean (%) mortality Corrected 0 48h 72h 96h 120h 144h 168h

Figure 7.1 Toxicity of spinosad residues to predatory mites after 96 h post-release exposure period. Within each residue age, means with different letters differed significantly (α = 0.05).

It also appears that the patterns of toxicity of spinosad of double the recommended rate to T. montdorensis, N. cucumeris and H. miles differed during the post-release exposure periods (Table 7.2). For all three mite species there was a significant decline in mortality during each post-release exposure period as residue time increased. When spinosad was applied at twice the

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Table 7.2 Residual toxicity of spinosad (twice the recommended rate) to predatory mites at 24 h, 48 h, 72, h and 96 h post-release exposure periods. Mites were fed spinosad intoxicated WFT larvae and simultaneously exposed to residue. Residual toxicity was classified: 1 = harmless (<25% mortality), 2 = slightly harmful (25-50% mortality), 3 = moderately harmful (51-75% mortality), and 4 = harmful (>75% mortality). Persistence class: A = short lived (<5 d), B = slightly persistent (5-15 d).

Corrected mortality (%) (mean ± SE) at post- Toxicity class Res. release periods Per. Age 24h* 48h* 72h* 96h* 24 48 72 96 Class h h h h T. montdorensis 2h 100e 100f 100f 100f 4 4 4 4 24h 100e 100f 100f 100f 4 4 4 4 48h 91.3±2.3e 96.0±1.7f 100f 100f 4 4 4 4 72h 50.3±1.4d 60.5±1.5e 67.5±1.6e 71.1±1.9e 3 3 3 3 96h 33.8±1.6c 45.8±1.7d 54.1±1.9d 56.2±1.5d 2 2 3 3 B 120h 10.0±0.9b 22.4±1.3c 30.5±1.6 31.0±1.8c 1 1 2 2 144h 9.5±0.8b 17.6±1.3b 24.6±1.7b 24.6±1.7b 1 1 1 1 168h 3.0±0.9a 8.3±0.9a 16.5±1.5a 16.8±1.4a 1 1 1 1 F 709.26 669.08 409.94 372.11 df 7 & 152 7 & 152 7 & 152 7 & 152 P < 0.0001 < 0.0001 < 0.0001 < 0.0001 N. cucumeris 2h 100g 100g 100g 100f 4 4 4 4 24h 100g 100g 100g 100f 4 4 4 4 48h 91.5±2.2f 94.4±1.8f 96.4±1.3f 96.7±1.3f 4 4 4 4 72h 52.5±1.5e 58.5±1.7e 66.1±1.6e 68.9±1.8e 3 3 3 3 96h 29.8±1.4d 38.5±1.9d 45.2±2.0d 48.7±2.1d 2 2 2 2 A 120h 14.8±1.0c 21.0±1.5c 26.8±1.4c 26.8±1.4c 1 1 1 2 144h 3.3±1.1b 10.7±1.2b 16.3±1.2b 16.3±1.2b 1 1 1 1 168h 0.0±0.0a 5.8±0.6a 7.4±0.9a 7.4±0.9a 1 1 1 1 F 881.76 647.50 636.77 621.14 df 7 & 152 7 & 152 7 & 152 7 & 152 P < 0.0001 < 0.0001 < 0.0001 < 0.0001 H. miles 2h 100f 100f 100e 100e 4 4 4 4 24h 100f 100f 100e 100e 4 4 4 4 48h 95.8±1.8f 100f 100e 100e 4 4 4 4 72h 79.8±2.1e 86.5±2.4e 86.9±2.4d 89.7±1.9d 4 4 4 4 96h 40.8±1.5d 49.4±1.4d 51.8±1.5c 57.6±2.5c 2 3 3 3 B 120h 27.3±2.4c 35.9±2.2c 37.4±2.0b 37.4±2.0b 2 2 2 2 144h 10.5±0.6b 23.2±1.3b 32.3±1.8b 32.3±1.8b 1 2 2 2 168h 3.8±1.0a 11.5±1.2a 22.3±1.8a 22.3±1.8a 1 1 1 1 F 537.45 471.23 348.82 373.99 df 7 & 152 7 & 152 7 & 152 7 & 152 P < 0.0001 < 0.0001 < 0.0001 < 0.0001 *For each species, within column, means with different letters differed significantly (α = 0.05).

177 Chapter VII: Resistance management recommended rate, mortality of T. montdorensis was different among spinosad residues at each post-release exposure period. Mortality of T. montdorensis was highest and lowest when exposed to 2 h and 168 h old residues respectively. However, at 24 h or 48 h post-release exposure periods, there was no significant difference in T. montdorensis’ mortality when exposed to 2 h and 24 h old residues. In addition, when examined at 72 h or 96 h post-release exposure periods, T. montdorensis’ mortality did not differ between 2 h, 24 h and 48 h old residues. Similarly, at each post-release exposure period, mortality of N. cucumeris or H. miles was highest and lowest when exposed to 2 h and 168 h old residues respectively. However, at 24 h, 48 h or 72 h post-release exposure periods, N. cucumeris mortality did not differ between 2 h and 24 h residues. While at 96 h post-release exposure period, there was no difference in N. cucumeris’ mortality when exposed to 2 h, 24 h and 48 h old residues. For H. miles, mortality due to 2 h, 24 h and 48 h old residues was not different. Moreover, mortality of each species of predatory mites due to spinosad residues was increased as the mites remain for longer periods in the Petri dish.

Spinosad residues declined in toxicity with age (Table 7.2). According to the IOBC toxicity classification, residues aged 2 h to 48 h were harmful (>75% mortality) to all three species for as long as 96 hours after the mites were first exposed. Hypoaspis miles was the most sensitive to spinosad residues, with 96 h old residue classified as moderately harmful. Based on the IOBC toxicity ratings, spinosad caused no harmful effect to N. cucumeris five days post-release (rating 1= < 25% mortality) and was considered to be short-lived (persisting for less than five days). For T. montdorensis and H. miles, spinosad was classified as slightly persistent (persisting for 5- 15 days) with 120 h (5 days) old residue for T. montdorensis and 120 h (5 days) and 144 h (6 days) old residues for H. miles classified as slightly harmful (25-50% mortality).

The relationships between predatory mite mortality and spinosad residues, with mortality as a function of residue age (log hrs) are presented in Figure 7.2. The LT25 (25% mortality of non- target organisms caused by residual toxicity) indicates that the release of each species of predatory mite after a spinosad application would cause only 25% mortality, allowing 75% survival of predatory mites. The LT25 of spinosad differed for each species and was estimated as 2.16660 2.10669 6.1 days (146.76 h; Antilog10 ) for T. montdorensis, 5.3 days (127.85 h; Antilog10 ) for 2.21071 N. cucumeris and 6.8 days (162.45 h; Antilog10 ) for H. miles (Table 7.3).

178 Chapter VII: Resistance management

Table 7.3 Probit analysis (Abbott 1925) of the mortality of predatory mites exposed to spinosad residues (by consumption of intoxicated WFT larvae and simultaneous exposure to residues) of different ages.

LT 95% CL Species Slope ± SE 25 χ2 df P (hrs) Lower Upper T. montdorensis -4.55±0.25 146.76 141.51 152.82 12.64 6 0.0491 N. cucumeris -5.36±0.19 127.85 124.24 131.85 10.20 6 0.1167 H. miles -5.37±0.37 162.45 156.86 169.02 16.56 6 0.0111

1.0 A

0.5

0.0 1.0

B

0.5 Probit mortality Probit

0.0 1.0

C

0.5

0.0 1.0 1.5 2.0 2.5 3.0

Log10 (hrs)

Figure 7.2 Relationship of Log10 (hrs) and probit mortality of (A) T. montdorensis, (B) N. cucumeris and (C) H. miles when exposed to twice the recommended rate of spinosad residues with different ages.

179 Chapter VII: Resistance management

7.3.3 Efficacy of predatory mites with spinosad against WFT-resistant strain

7.3.3.1 Effect of spinosad and predatory mite releases on WFT adults

Predatory mites (T. montdorensis, N. cucumeris, and H. miles) were more effective at reducing WFT adult numbers after spinosad is applied (Figure 7.3). Since there was a statistically significant interaction between spray [spinosad, water (control)] and mite releases (no mites, T. montdorensis, N. cucumeris, H. miles) (Appendix 7.1), treatments (spinosad, water) were separately analysed at each observation [DAS = days after spray] (Appendix 7.2).

Spinosad Water

50 10 DAS 15 DAS 40 30 20 10

SE) 0  50 20 DAS 25 DAS 40 30 20 10 0 50 Number of WFT adults (Mean adults WFT of Number 30 DAS 35 DAS 40 30 20 10 0 No mitesTM NC HM No mitesTM NC HM

Figure 7.3 Mean numbers of WFT adults per plant sprayed with spinosad or water (control) in the presence or absence of predatory mite. TM = T. montdorensis, NC = N. cucumeris, HM = H. miles. DAS = Days after spray. Means with different letters differed significantly (α = 0.025).

There were no treatment differences in WFT adult numbers (plants assigned to mites release), prior to mite releases at 1 DAS (days after spray) (Fspinosad = 0.583, 16, P = 0.6368; Fwater = 2.573,

180 Chapter VII: Resistance management

16, P = 0.0905) and 5 DAS (Fspinosad = 0.463, 16, P = 0.7152; Fwater = 0.923, 16, P = 0.4551) (Figure 7.4). From 10 to 35 DAS, mean numbers of WFT adults were significantly different among predatory mite treatments (no mites, T. montdorensis, N. cucumeris, H. miles), except at 10 DAS in the control treatment (water) (Appendix 7.2; Figure 7.4B). Plants treated with ‘no mites’ had the highest numbers of WFT adults compared to plants that were treated with spinosad and received predatory mites (Figure 7.4A). Similarly, in the water treatment (control), plants treated with ‘no mites’ had the highest numbers of WFT adults compared to plants that received predatory mites (Figure 7.4B). In both spinosad and water (control) treatments, T. montdorensis was the most effective species at reducing WFT adult numbers, except at 10 and 15 DAS. Hypoaspis miles was most effective at 10 and 15 DAS in both spinosad and control (water) treatments, but the least effective species from 20 to 35 DAS compared with T. montdorensis and N. cucumeris.

No mites TM NC HM 40 A

30 SE)  20

10

0

40 B

30

20

Number of adults per plant (Mean plant per adults of Number 10

0 D1* D5* D10 D15 D20 D25 D30 D35

Figure 7.4 Numbers of WFT adults per plant (Y-axis) sprayed with spinosad (A) or water (B) with or without predatory mite. In each observation (D10 to D35, D = days after spray), means were separated by least square mean difference at α = 0.025. *WFT adults’ numbers per plant before mite release.

7.3.3.2 Effect of spinosad and predatory mite releases on WFT larvae

Predatory mites (T. montdorensis, N. cucumeris, and H. miles) were more successful at reducing WFT larvae numbers after spinosad is applied (Figure 7.5). Because of the statistically significant interaction between spray [spinosad, water (control)] and mite treatment (no mites, T. montdorensis, N. cucumeris, H. miles), effects of mite treatment on WFT larvae were

181 Chapter VII: Resistance management separately analysed for spinosad and water spray at each observation [DAS = days after spray] (Appendix 7.1 and 7.2).

Spinosad Water

40 10 DAS 15 DAS 30 b 20 b b

SE) b b

 b a b 10 a b a a a a a a 0 40 20 DAS 25 DAS 30 b b 20 a b b a a b b a b b 10 a a a a 0 40 30 DAS 35 DAS 30 a b a Number of WFT larvae per plant (Meanplant per ofWFTNumber larvae a a a 20 a a b b b b a 10 a a a 0 No mitesTM NC HM No mitesTM NC HM

Figure 7.5 Numbers of WFT larvae per plant with spinosad and water (control) in the presence or absence of predatory mites. DAS = days after spray. TM = T. montdorensis, NC = N. cucumeris, HM = H. miles. Means within group with different letters differed significantly (α = 0.025).

When plants were examined at one DAS, no WFT larvae were found. At five DAS prior to predatory mites release, there was no differences in WFT larvae numbers per plant among treatments (plants assigned to predatory mites release) in spinosad (F 3, 16 = 2.00, P = 0.1546) or water (control) (F 3, 16 = 0.80, P = 0.5111) treatments (Figure 7.6). From 10 to 35 DAS, mean numbers of WFT larvae per plant were significantly different among predatory mite treatments (no mites, T. montdorensis, N. cucumeris or H. miles) in both spinosad and water (control) treatments (Appendix 7.2; Figure 7.6). In spinosad treatment, plants treated with ‘no mites’ had the highest numbers of WFT larvae compared to plants received predatory mites. Similarly, for the water treatment (control), plants treated with ‘no mites’ had the highest numbers of WFT larvae compared to plants that received predatory mites. For both spinosad and water (control)

182 Chapter VII: Resistance management treatments, T. montdorensis was the most effective species in reducing WFT larvae, except at 15 DAS in spinosad treatment and 10 DAS in the water (control) treatment. Neoseiulus cucumeris was the most effective in reducing WFT larvae at 15 DAS and 10 DAS in spinosad and water (control) treatments respectively. H. miles was the least effective species in reducing WFT larvae compared with T. montdorensis and N. cucumeris.

No mites TM NC HM A 30 SE)  20

10

0 30 B

20

10 Number of WFT larvae per plant (Mean plant per larvae WFT of Number

0 D1* D5* D10 D15 D20 D25 D30 D35

Figure 7.6 Numbers of WFT larvae per plant with spinosad (A) and water (B) in the presence or absence of predatory mites (TM = T. montdorensis, NC = N. cucumeris, HM = H. miles). At each observation (D = days after spray), means were separated by least square mean difference at α = 0.025. *WFT nymphs per plant before release of mites.

7.4 Discussion

The present study suggests that to control 50% or more of a spinosad-resistant WFT strain, the recommended rate of spinosad (80 mL/100L, 0.096 g a.i./L) will need to be doubled (160 mL/100L, 0.192 g.a.i./L). However, by doubling the rate, toxicity to predatory mites (T. montdorensis, N. cucumeris and H. miles) released for biological control of WFT increased and caused 100% mortality. The predatory mites used in this study cannot tolerate direct contact with spinosad at the above rate.

Fresh to relatively fresh residues (2 to 72 h old) of double the recommended rate of spinosad were also harmful to T. montdorensis, N. cucumeris, and H. miles, causing 100% mortality. Spinosad is considered to be a short-lived insecticide that rapidly degrades in nature (Thompson et al. 2000, Williams et al. 2003). Under laboratory conditions; however, spinosad is considered

183 Chapter VII: Resistance management to be highly stable and capable of causing high mortality to a parasitoid, up to 1 month after being applied to foliage or artificial surfaces (Bernando and Viggiani 2000). Semi-field and field studies suggest that spinosad residues pose little or no toxicity to natural enemies after three to 7 days based on species (Boyd and Boethel 1998, Ruberson and Tillman 1999, Crouse et al. 2001). This study was conducted in the laboratory. However, the effect of spinosad on predatory mites suggests that the effects of spinosad residues decreases over time, and would pose little threat to the predator if the initial exposure to the pesticide occurs after a period of time. Based on residual threshold (LT25) calculations and confirmed with laboratory trials, mites can be released 5 to 7 days after a spinosad application. Neoseiulus cucumeris could be released after 5.3 days (127.85 h), T. montdorensis after 6.1 days (146.76 h), and H. miles after 6.8 days (162.45 h). Despite the increase in the rate of spinosad applied, the results presented here suggest that predatory mites (T. montdorensis, N. cucumeris and H. miles) can be integrated with an increased rate of spinosad application for the effective management of spinosad- resistant WFT.

In the bioassay, spinosad was classified to have short-lived persistence against T. montdorensis and N. cucumeris, while slightly persistent for H. miles. The variable effect of spinosad residues is not unexpected. There is variation in the residual effect of insecticides including spinosad on other species. It has been reported that LC50 values of spinosad exceeded 960 ppm for Podius nigrispinus (Dallas) (Hemiptera: Pentatomidae) (Torres et al. 1999), while LC50 was only 50 ppm for Podius maculiventris (Say) (Hemiptera: pentatomidae) (Viñuela et al. 1998). Similar to the present findings, Sáenz-de-Cabezón Irigaray et al. (2007) stated that the miticide, abamectin was short-lived to the western predatory mite, Galendromus (= Typhlodromus, = Metaseiulus) occidentalis Nesbitt (Acari: Phytoseiidae), and slightly persistent to Phytoseiulus persimilis Athias-Henriot (Acari: Phytoseiidae). Khan and Morse (2006) tested the impact of four pesticides on the predatory mite Euseius tularensis Congdon, and found residual effects if mites were released five to six days after spinosad was applied. But spinosad had no residual effect on E. tularensis if released seven days after the spinosad application (Khan and Morse 2006).

As with previous findings (Chapters 3, 4 and 5), the efficacy of the mites in reducing WFT differed. Typhlodromips montdorensis appears to be the most effective species in suppressing WFT, followed by N. cucumeris and H. miles. There may be a variety of reasons for such variation (Brødsgaard 1989, van Houten et al. 1995, Steiner and Goodwin 1998, 2001, Berndt et al. 2004, Messelink et al. 2006, Skirvin et al. 2007). Nevertheless all three predatory mites tested in this study provided better control of WFT when released after spinosad was applied. However, to get maximum suppression of WFT using these predatory mites, a further study detailing factors that influence their success needs to be carried out.

184 Chapter VII: Resistance management

In summary, the challenge for managing resistant pest populations is that management with insecticides often requires that the dose applied has to be increased in order to be effective (Broughton and Herron 2007). The consequential effects on natural enemy means that often the two tactics cannot be used together. However, the results herein suggest that spinosad-resistant WFT could be managed with an increased rate of spinosad and could be further reduced by releasing predatory mites. However, caution is required as direct exposure to spinosad or relatively fresh residues are toxic to T. montdorensis, N. cucumeris, and H. miles. Since predatory mites predate on either first instar larvae or pupal stages but not the adult stage, the adult WFT population needs to be reduced before predatory mites are released. As this study was conducted in a controlled environment, pesticide risk assessments need to be validated with semi-field and/or field studies.

7.5 Literature cited

Abbott, W. S. 1925. A method of computing the effectiveness of an insecticide. Journal of Economic Entomology 18: 265-267. Anh, K. S., S. Y. Lee, K. Y. Lee, Y. S. Lee, and G. H. Kim. 2004. Selective toxicity of pesticides and control effects of the two-spotted spider mite, Tetranychus urticae by predatory mite and pesticide mixture on rose. Korean Journal of Applied Entomology 43: 71-79. Bernando, U., and G. Viggiani. 2000. Effects of spinosad, a new insect control agent naturally derived on the mealybug parasitoids Leptomastix dactylopii Howard (Hymenoptera: Encyrtidae). Bulletin IOBC/WPRS 23: 81-84. Berndt, O., H.-M. Poehling, and R. Meyhofer. 2004. Predation capacity of two predatory laelapid mites on soil-dwelling thrips stages. Entomologia Experimentalis et Applicata 112: 107-115. Bielza, P., V. Quinto, J. Contreras, M. Torne, A. Martin, and P. J. Espinosa. 2007. Resistance to spinosad in the western flower thrips, Frankliniella occidentalis (Pergande), in greenhouses of southeastern Spain. Pest Management Science 63: 682- 687. Boyd, M. L., and D. J. Boethel. 1998. Residual toxicity of selected insecticides to heteropteran predaceous species (Heteroptera: Lygaeidae, Nabidae, Pentatomidae) on soybean. Environmental Entomology 27: 154-160. Bret, B. L., L. L. Larson, J. R. Schoonover, T. C. Sparks, and G. D. Thomson. 1997. Biological properties of Spinosad. Down to Earth 52: 6-13. Broadbent, A. B., and D. J. Pree. 1997. Resistance to insecticides in populations of Frankliniella occidentalis (Pergande) (Thysanoptera: Thripidae) from greenhouses in the Niagara region of Ontario. Canadian Entomologist 129: 907-913. Brødsgaard, H. F. 1989. Coloured sticky traps for Frankliniella occidentalis (Thysanoptera: Thripidae) in glasshouse. Journal of Applied Entomology 107: 136-140. Brødsgaard, H. F. 1994. Insecticide resistance in Europe and African strains of western flower thrips (Thysanoptera: Thripidae) tested in a new residue-on-glass test. Journal of Economic Entomology 87: 1141-1146. Broughton, S., and G. A. Herron. 2007. Frankliniella occidentalis (Pergande) (Thysanoptera: Thripidae) chemical control: insecticide efficacy associated with the three consecutive spray strategy. Australian Journal of Entomology 46: 140-145. Cisneros, J., D. Goulson, L. C. Derwent, D. I. Penagos, O. Hernandez, and T. Williams. 2002. Toxic effects of spinosad on predatory insects. Biological Control 23: 2002.

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Contreras, J., D. Moreno, M. D. Hernandez, P. Bielza, and A. Lacasa. 2001. Preliminary study on insecticide resistance in Frankliniella occidentalis (Pergande) (Thysanoptera: Thripidae) in sweet pepper crops in Campo de Cartagena (Murcia), S. E. of Spain. Acta Horticulturae 559: 745-752. Crouse, G. D., T. C. Sparks, J. Schoonover, J. Gifford, J. Dripps, T. Bruce, L. L. Larson, J. Garlich, C. Hatton, R. L. Hill, T. V. Worden, and J. G. Martynow. 2001. Recent advances in the chemistry of spinosyn. Pest Management Science 57: 177-185. Espinosa, P. J., P. Bielza, J. Contreras, and A. Lacasa. 2002. Insecticide resistance in field populations of Frankliniella occidentalis (Pergande) in Murcia (south-east Spain). Pest Management Science 58: 967-971. Finney, D. J. 1971. Probit Analysis. Cambridge University press, Cambridge. Georghiou, G. P. 1990. Overview of insecticide resistance, pp. 18-41. In M. B. Green, H. M. LeBaron and W. K. Moberg [eds.], Managing resistance to agrochemicals: from fundamental research to practical strategies. American Chemical Society, Washington DC. GraphPad Software Inc 2007. GraphPad Prism computer program, version 5.0. By GraphPad Software Inc. Hassan, S. A., F. Bigler, H. Bogenschütz, E. Boller, J. Brun, J. N. M. Calis, J. Coremans- Pelseneer, C. Duso, A. Grove, U. Heimbach, N. Helyer, H. Hokkanen, G. B. Lewis, F. Mansour, L. Moreth, L. Polgar, L. Samsoe-Petersen, B. Sauphanor, A. Staubli, G. Sterk, A. Vainio, M. van de Veire, G. Viggiani, and H. Vogt. 1994. Results of the sixth joint pesticide testing programme of the IOBC/WPRS-working group <>. Entomophaga 39: 107-119. Healy, M. J. R., and L. R. Taylor. 1962. Tables of Power - Law transformations. Biometrika 49: 557-559. Herron, G. A., and G. C. Gullick. 2001. Insecticide resistance in Australian populations of Frankliniella occidentalis (Pergande) (Thysanoptera: Thripidae) causes the abandonment of pyrethroid chemicals for its control General and Applied Entomology 30: 21-26. Herron, G. A., and T. M. James. 2005. Monitoring insecticide resistance in Australian Frankliniella occidentalis Pergande (Thysanoptera: Thripidae) detects fipronil and spinosad resistance. Australian Journal of Entomology 44: 299-303. Herron, G. A., S. Broughton, and A. Clift. 2007. Frankliniella occidentalis (Pergande) (Thysanoptera: Thripidae) chemical control: residues associated with the three consecutive spray strategy. Australian Journal of Entomology 46: 146-151. Jensen, S. E. 2000. Insecticide resistance in the western flower thrips, Frankliniella occidentalis. Integrated Pest Management Reviews 5: 131-146. Khan, I., and J. G. Morse. 2006. Impact of citrus thrips chemical treatments on the predatory mite Euseius tularensis. Journal of Applied Entomology 130: 386-392. Kim, S. S., S. G. Seo, J. D. Park, S. G. Kim, and D. I. Kim. 2005. Effects of selected pesticides on the predatory mite, Amblyseius cucumeris (Acari: Phytoseiidae). Journal of Entomological Science 40: 107-114. Kongchuensin, M., and A. Takafuji. 2006. Effects of Some Pesticides on the Predatory Mite, Neoseiulus longispinosus (Evans) (Gamasina: Phytoseiidae). Journal of Acarology Society of Japan 15: 17-27. Lewis, T. 1998. Pest thrips in perspective, pp. 385-390, Proceedings The 1998 Brighton Conference- Pest and Diseases. British Crop Protection Council, Brighton, UK. Messelink, G. J., S. E. F. van Steenpaal, and P. M. J. Ramakers. 2006. Evaluation of phytoseiid predators for control of western flower thrips on greenhouse cucumber. Biocontrol 51: 753-768. Miles, M., R. Dutton, H. Vogt, U. Heimbach, and E. Vinuela. 2003. Testing the effects of spinosad to predatory mites in laboratory, extended laboratory, semi-field and field studies. Bulletin of IOBC/WPRS 26: 9-20. Quinn, G. P., and M. J. Keough. 2002. Experimental design and data analysis for biologists. University Press, Cambridge.

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Ruberson, J. R., and P. G. Tillman. 1999. Effect of selected insecticides on natural enemies in cotton: laboratory studies, pp. 1210-1212, Beltwide Cotton Conference. National Cotton Council of America, Memphis, TN. Sáenz-de-Cabezón Irigaray, F. J., F. G. Zalom, and P. B. Thompson. 2007. Residual toxicity of acaricides to Galendromus occidentalis and Phytoseiulus persimilis reproductive potential. Biological Control 40: 153-159. SAS 2002-2003. SAS 9.1 computer program, version 9.1. By SAS, Cary, NC, USA. Shipp, J. L., K. Wang, and G. Ferguson. 2000. Residual Toxicity of Avermectin b1 and Pyridaben to Eight Commercially Produced Beneficial Arthropod Species Used for Control of Greenhouse Pests. Biological Control 17: 125-131. Skirvin, D. J., L. Kravar-Garde, K. Reynolds, J. Jones, A. Mead, and J. Fenlon. 2007. Supplemental food affects thrips predation and movement of Orius laevigatus (Hemiptera: Anthocoridae) and Neoseiulus cucumeris (Acari: Phytoseiidae). Bulletin of Entomological Research 97: 309-315. Steiner, M. Y., and S. Goodwin. 1998. Development and marketing of an IPM package for western flower thrips, Frankliniella occidentalis (Pergande). Phase II: Evaluation and producing the natural enemies. HRDC/HSNA Report. NSW Agriculture, Gosford, Australia. Steiner, M. Y., and S. Goodwin. 2001. Development and marketing of an IPM package for western flower thrips, Frankliniella occidentalis. Phase III: Development and evaluation of usage protocols for newly developed western flower thrips biocontrol agents. HRDC report. NSW Agriculture, Gosford, Australia. Steiner, M. Y., S. Goodwin, T. M. Wellham, I. M. Barchia, and L. J. Spohr. 2003. Biological studies of the Australian predatory mite Typhlodromips montdorensis (Schicha) (Acari: Phytoseiidae), a potential biocontrol agent for western flower thrips, Frankliniella occidentalis (Pergande) (Thysanoptera: Thripidae). Australian Journal of Entomology 42: 124-130. Sterk, G., S. A. Hassan, M. Baillod, F. Bakker, F. Bigler, S. Blümel, H. Bogenschütz, E. Boller, B. Bromand, J. Brun, J. N. M. Calis, J. Coremans-Pelseneer, C. Duso, A. Garrido, A. Grove, U. Heimbach, H. Hokkanen, J. Jacas, G. Lewis, L. Moreth, L. Polgar, L. Rovestl, L. Samsoe-Petersen, B. Sauphanor, L. Schaub, A. Stăubli, J. J. Tuset, A. Vainio, M. van de Veire, G. Viggiani, E. Viñuela, and H. Vogt. 1999. Results of the seventh joint pesticide testing programme carried out by the IOBC/WPRS working group pesticides and beneficials. Biocontrol 44: 99-117. Tabashnik, B. E., and B. A. Croft. 1982. Managing pesticide resistance in crop arthropod complexes: interactions between biological and operational factors. Environmental Entomology 11: 1137-1144. Thompson, G. D., R. Dutton, and T. C. Sparks. 2000. Spinosad-a case study: an example from natural products discovery programme. Pest Management Science 56: 696-702. Tommasini, M. G., and S. Maini. 1995. Frankliniella occidentalis and other thrips harmful to vegetable and ornamental crops in Europe, pp. 1-42. In A. J. M. Loomans, J. C. Van Lenteren, M. G. Tommasini, S. Maini and J. Riudavets [eds.], Biological Control of Thrips, Wageningen Agricultural University, Wageningen, the Netherlands. Torres, J. B., P. de Clercq, and R. Barros. 1999. Effect of spinosad on the predator Podius nigrispinus and its lepidopreous prey. Mededelingen Facuteit Landbouwkundige en Toegepaste Biologische Wetenschappen, Universiteit Gent 64: 211-218. van Driesche, R. G., S. Lyon, and C. Nunn. 2006. Compatibility of spinosad with predacious mites (Acari: Phytoseiidae) used to control western flower thrips (Thysanoptera: Thripidae) in greenhouse crops. Florida Entomologist 89: 396-401. van Houten, Y. M., P. van Stratum, J. Bruin, and A. Veerman. 1995. Selection for non- diapause in Amblyseius cucumeris and Amblyseius barkeri and exploration of the effectiveness of selected strains for thrips control. Entomologia Experimentalis et Applicata 77: 289-295. Villanueva, R. T., and J. F. Walgenbach. 2005. Development, oviposition, and mortality of Neoseiulus fallacis (Acari: Phytoseiidae) in response to reduced-risk Insecticides. Journal of Economic Entomology 98: 2114-2120.

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Viñuela, E., M. P. Medina, M. Schneider, M. González, F. Budia, A. Adán, and P. Delestal. 2001. Comparison of side effects of spinosad, tebufenozide, and azadirachtin on the predators Chrysoperla carnea and Podisus maculiventris and the parasitoids Opius concolor and Hyposoter didymator under laboratory conditions. Bulletin IOBC/WPRS 24: 25-34. Viñuela, E., A. Adan, M. Gonzalez, F. Budia, G. Smagghe, P. de Clercq, H. Vogt, and P. Del Estal. 1998. Spinosa y azadiractina: effects de dos plaguicidas de origen natural en el chinche depredator Podius maculiventris (Say) (Hemiptera: pentatomidae). Bulletin Sanidad Vegetal Plagas 24: 57-66. Williams, T., J. Valle, and E. Vinuela. 2003. Is the naturally derived insecticide Spinosad compatible with insect natural enemies? Biocontrol Science and Technology 13: 459- 475. Zar, J. H. 1999. Biostatistical Analysis. Prentice Hall International, Upper Saddle River, New Jersey. Zhao, G. Y., W. Liu, and C. O. Knowles. 1994. Mechanisms associated with diazinon resistance in western flower thrips. Pesticide Biochemistry and Physiology 49: 13-23. Zhao, G. Y., W. Liu, J. M. Brown, and C. O. Knowles. 1995. Insecticide resistance in field and laboratory strains of western flower thrips (Thysanoptera: Thripidae). Journal of Economic Entomology 88: 1164-1170. Zilahl-Balogh, G. M. G., J. L. Shipp, C. Cloutier, and J. Brodeur. 2007. Predation by Neoseiulus cucumeris on western flower thrips, and its oviposition on greenhouse cucumber under winter vs. summer conditions in a temperate climate. Biological Control 40: 160-167.

188 Chapter VIII

General discussion and conclusion

8.1 General discussion

In this thesis, I hypothesised that chemical control, biological control, and resistant host-plants could be effectively combined to reduce pest numbers on a horticultural commodity. I sought to test this hypothesis with a significant and difficult to manage pest in a crop with high aesthetic value, and in Australia, where few alternatives to pesticides are available for management. Since its first detection in 1895 in California, western flower thrips, Frankliniella occidentalis (Pergande) (Thysanoptera: Thripidae) has spread throughout the world and is considered a significant pest. Frankliniella occidentalis is hard to control with pesticides and this has led to increased efforts towards the implementation and use of integrated pest management (IPM). In Australia, an IPM approach for F. occidentalis that incorporates multiple tactics has been difficult, because few alternatives to pesticides have been available besides pesticides. This is now changing now with the commercial availability of predatory mites, a predatory bug, predatory beetle and nematodes (Australian Biological Control 2009). However, to use biological and chemical control effectively, the tactics need to be carefully applied. Frankliniella occidentalis is a pest of many commercial crops and is particularly damaging to strawberry. Several varieties of strawberry are grown in Australia but none has been explicitly developed for resistance to F. occidentalis. The aim of this body of work was to evaluate the compatibility of chemical (spinosad) and biological control (predatory mites), along with host- plant resistance for the management of F. occidentalis in strawberry.

To test the hypothesis, this project sought first to establish the influence of strawberry cultivars on feeding preference and oviposition preference and performance of F. occidentalis. Secondly, this study tested the efficacy of three commercially available predatory mites [Typhlodromips montdorensis (Schicha) (Phytoseiidae), Neoseiulus cucumeris (Oudemans) (Phytoseiidae), and Hypoaspis miles (Berlese) (Laelapidae)] with or without spinosad along with three strawberry cultivars (Camarosa, Camino Real, and Albion) against F. occidentalis. Thirdly, this project investigated the efficiency of single- versus multiple-species release of predatory mites with or without spinosad in the management of the F. occidentalis population. Fourthly, this project sought to test the compatibility of predatory mites and the release the protocol for the management of F. occidentalis in low tunnel-grown strawberry. Fifthly, this study evaluates the residual threshold of spinosad for the predatory mites. Lastly, this study investigates the possibility of the integration of these Chapter VIII Summary predatory mites with an increased application rate of spinosad [twice the recommended rate for F. occidentalis] to control a spinosad-resistant F. occidentalis strain.

8.2 Findings and recommendations

8.2.1 Strawberry cultivars distinctively influence western flower thrips’ olfactory and feeding preference and oviposition preference and performance

Host-plant resistance to arthropod pests generally plays an important role in pest management programs (White 1969, Wellington 1977, White 1993, Parrella and Lewis 1997) and is considered a key method for pest control in many crops with a low economic threshold (Schoonhoven et al. 1998). To my knowledge, no strawberry cultivars have been tested for their resistance to F. occidentalis. I tested three strawberry cultivars developed by the University of California for differences in their suitability as hosts of F. occidentalis. These were Camarosa and Camino Real, which are short-day varieties, and Albion, an ever-bearing (day-neutral) variety. These cultivars are grown commercially in Australia and overseas (California Strawberry Commission 2009).

The experiments conducted in this study suggest that different cultivars influence the feeding and oviposition behaviour of F. occidentalis. Of the three cultivars tested, there was clear variation in thrips behaviour and survival. Camarosa was most preferred by F. occidentalis adults and it appeared to favour F. occidentalis development. The cultivar Camino Real was the least preferred cultivar and the most unfavourable for F. occidentalis. There may be other cultivars that are less favourable to F. occidentalis than Camino Real. Growers currently base their cultivar selections on marketable yield, fruit colour, fruit size, taste and resistance to diseases such as verticillium wilt and phytophthora crown rot (Phillips and Reid 2008). Further screening of these cultivars would enable resistance information to be provided to growers, and allow them to include thrips resistance in their decision-making. This study did not determine the specific factors or characteristics that influence thrips preference and performance, so future work should be conducted in this area. In addition, host-plant resistance to herbivores is influenced by seasonal changes, and this should also be addressed.

8.2.2 Biological control: multiple species versus single species

All three predatory mites tested are effective predators of F. occidentalis. Typhlodromips montdorensis was the most effective at reducing F. occidentalis, followed by N. cucumeris and H. miles. Multiple-species release of predatory mites provided better management of F.

190 Chapter VIII Summary occidentalis than releases of a single species of predatory mites. As an integrated approach, there is a growing trend to use two or more species of natural enemies to suppress insect pest populations (Premachandra et al. 2003, Avilla et al. 2004, Blümel 2004, Brødsgaard 2004, Chau and Heinz 2004, Chow and Heinz 2004, Hoddle 2004, Shipp and Ramakers 2004, Thoeming and Poehling 2006, Chow et al. 2008). By maintaining a lapse of time between spinosad application and predatory mite release, both methods can be incorporated to reduce F. occidentalis numbers. Management can be further improved when combined with the cultivar Camino Real. However, multiple-species release of natural enemies in pest a management program is not always quantitatively validated (Blockmans and Tetteroo 2002, Skirvin et al. 2006). Some of the studies support the premise that species are compatible (Gillespie and Quiring 1992, Wittmann and Leather 1997, Brødsgaard and Enkegaard 2005), whilst others suggest otherwise (Magalhăes et al. 2004, Sanderson et al. 2005). This study found that any combination of multiple-species release of predatory mites (T. montdorensis, N. cucumeris, and H. miles) performed better in reducing F. occidentalis than respective single species. However, predatory mite species varied in their compatibility. In this study, when combined together, T. montdorensis and N. cucumeris were less effective than other combinations. However, they performed better when applied in triple-species combination (second-best combination). When predatory mites were released in double-species combinations, their numbers (as a proportion of total mites released) were higher than in triple-species combination. This may increase the chance of interspecific competition as the mites share the same resource. Further research should study the optimal release rate, which might eliminate interspecific competition. One of the limitations of the predatory mites used in this study is that they are either larval or pupal predators. This means that adult F. occidentalis escaped predation. Therefore, future study should explore combining mite releases with other predators such as the anthocorid bug, Orius armatus, which preys on both larval and adult stage or other natural enemies (Baez et al. 2004).

8.2.3 Combining chemical and biological control

In addition to host-plant resistance, it has been argued that the integration of chemical and biological control could provide better management of arthropod pests (Funderburk et al. 2000, Ludwig and Oetting 2001, Ludwig 2002, Thoeming and Poehling 2006, Funderburk 2009) than either tactic alone. Spinosad is a biopesticide classified as an environmentally and toxicologically reduced-risk chemical (Cleveland et al. 2002, Thompson et al. 2002). It is registered as an IPM-compatible insecticide in Australia (Thompson and Hutchins 1999). Spinosad is regarded to have low to moderate toxicity to predatory mites but the toxicity varies from species to species (Williams et al. 2003, Cote et al. 2004, Jones et al. 2005).

191 Chapter VIII Summary

Therefore, for successful integration of chemical and biological control, it is important to evaluate the residual threshold of a chemical on its respective natural enemies.

I found that a spinosad application followed by the release of commercially available predatory mites could effectively reduce the F. occidentalis population but only if there was sufficient time between spraying and release. As expected, the direct toxicity of recommended rate of spinosad (F. occidentalis management, 80 ml/100 L, 0.096 g. a.i./L) as well some aged residues were toxic to T. montdorensis, N. cucumeris and H. miles, causing substantial mortality. Thresholds for residual toxicity of spinosad LT25 (lethal time for 25% mortality) were estimated as 4.2 days (101.63h), 3.2 days (77.72) and 5.8 days (138.83 h) for T. montdorensis, N. cucumeris and H. miles respectively. At this time, spinosad posed no significant negative effect to mites. Based on the bioassays, spinosad was characterised as short- lived (degraded quickly) for T. montdorensis and N. cucumeris (degrade quickly) and slightly persistent for H. miles. It is important to note that residual threshold increased when predatory mites fed on intoxicated F. occidentalis and were simultaneously exposed to spinosad residues. This was estimated as 5.4 days (129.67), 4 days (95.09), and 6.1 days (146.68 h) for T. montdorensis, N. cucumeris, and H. miles respectively. Natural enemies subjected to multiple routes of exposure to pesticides may respond in unexpected ways that would be impossible to predict based on single-route exposure (Kunkel et al. 2001). Thus, estimates of residual thresholds should be based on persistence and harmfulness of indirect and contact toxicity, not contact toxicity alone.

Spinosad can degrade quickly in nature (Thompson et al. 2000, Williams et al. 2003). The bioassays conducted in this study were in the laboratory under controlled conditions. However, chemical degradation may be influenced by environmental factors such as temperature, photoperiod, and seasonal variation (van de Veire et al. 2002, Zilahl-Balogh et al. 2007). Persistence of a chemical also varies from season to season (van de Veire et al. 2002, Zilahl- Balogh et al. 2007). Spinosad may thus have more or less of a detrimental effect on the predatory mites than presented in these experiments.

8.2.4 IPM of western flower thrips in low tunnel-grown strawberry

Control of F. occidentalis and other thrips with natural enemies in glasshouse crops is successful in many parts of the world (Ramakers 1988). However, little is known about the efficacy of predatory mites in semi-open or open field conditions. It has been established that predatory mites are effective in glasshouse conditions where temperatures and humidity are higher. Can predatory mites be effective in low tunnel-grown crops during spring, when

192 Chapter VIII Summary conditions are much cooler? I found that predatory mites reduced F. occidentalis in low tunnel grown strawberry. The mites T. montdorensis, N. cucumeris and H. miles can be used in combination in low tunnels for F. occidentalis management even in the relatively cooler conditions. Gillespie and Ramey (1988) reported that N. cucumeris can survive at a constant temperature of 9⁰C for months and oviposit within three days when returned to room temperature (20-22°C). Typhlodromips montdorensis did not diapause when reared under 10⁰C with long day conditions (Steiner et al. 2003) and can survive at 8⁰C or above (Steiner and Goodwin 2002). In the low tunnel, the plastic cover keeps the tunnel warmer than the ambient air, which might provide a suitable environment for predatory mites at least during the time of year that the experiment was conducted. Longer-term studies on the population dynamics of the mites in the field environment should be conducted to determine the temperature range at which they can sustain their populations and remain active.

As an integrated approach, this study tested whether the use of predatory mites before or after spinosad application would be the better option for F. occidentalis management in low-tunnel grown strawberry. Spraying of spinosad followed by the release of predatory mites is more effective as shown in this strategy, spinosad provided effective reduction of F. occidentalis while having no significant detrimental effects on natural enemies. In the present study, the numbers of predatory mites were significantly lower in the ‘mites then spinosad’ treatment compared to the ‘spinosad then mites’ treatment, suggesting that there is a detrimental effect of spinosad on predatory mites when applied after the mites are released. In a low tunnel environment, the integration of spinosad and predatory mites was effective at reducing F. occidentalis population below the economic threshold (5 or more thrips per flower in 45% of flowers, (Steiner and Goodwin 2005)), but only for a limited period. Thus, there might be a need for a further application of either spinosad or of predatory mites or both to maintain the F. occidentalis population below the economic threshold throughout the cropping period.

8.2.5 Control of spinosad-resistant western flower thrips strain

Given the current high reliance on spinosad for F. occidentalis control and the increasing probability that resistance in Australian populations will spread (Herron et al. 2007), it is important to explore ways in which spinosad can be used with biological control agents effectively when resistance is evident. One approach is to use an initial high dose of an insecticide to reduce the resistant population, and thereafter use natural enemies to maintain the population at a low level. However, use of the high dose of an insecticide would likely increase the detrimental effect of a pesticide to natural enemies, particularly if there is any residual activity of the insecticide. This study found that twice the recommended rate of spinosad

193 Chapter VIII Summary

(recommended rate for F. occidentalis control, 160 mL/100 L, 0.192 g a.i./L) is very toxic to T. montdorensis, N. cucumeris and H. miles, causing 100% mortality. Moreover, the residues (depending on age) were also very toxic to the predatory mites. Despite the increased rate, spinosad can degrade rapidly by photolysis and is short-lived. It persisted for less than five days for N. cucumeris, and was classified as slightly persistent to T. montdorensis and H. miles. The residual threshold (LT25) of twice the recommended rate was found to be 5.3 days (127.85 h) 6.1 days (146.76 h), and 6.8 days (162.45 h) for N. cucumeris, T. montdorensis, and H. miles. By maintaining a lapse of 6-7 days between spray and predatory mite release, an increased rate of spinosad could be used for F. occidentalis management when required.

This study only tested the compatibility of predatory mites with spinosad for WFT management. However, the prey-predator ratio in which optimal management could be achieved was not determined. Strawberry crops are often infested by other pests such as spider mites, rootworm, leafrollers, root weevils , crown borers. These pests might also require chemical or biological control. Therefore, compatibility of these predatory mites with other chemicals, or other beneficials used for other pest control may also need to be evaluated.

8.3 Conclusions

This thesis evaluated the compatibility of an existing chemical (spinosad) and predatory mites (T. montdorensis, N. cucumeris and H. miles), combined with cultivar resistance for the management of F. occidentalis in strawberry. This thesis provides information for F. occidentalis management in strawberry, which can also be adopted for pest management in other horticultural crops where F. occidentalis is a problem. Differential performance of F. occidentalis on strawberry cultivars has been demonstrated in this thesis, suggesting that host- plant resistance could be used. Cultivar selection for thrips management has not been considered for the Australian strawberry industry. Likewise, careful integration of biological and chemical control has not been previously used for F. occidentalis management in Australia. The findings presented here provide multifaceted support towards an effective, integrated approach to manage F. occidentalis in field-grown strawberries. Pests continue to develop resistance to insecticides and this is the case with F. occidentalis and spinosad. I demonstrated that resistant strains of a pest could be managed by incorporating a pesticide with natural enemies, thus retaining both pesticides and natural enemies in the pest management toolbox. Where F. occidentalis has already begun to develop resistance to spinosad in Australia, it is critical that we find alternatives to its management.

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8.4 Literature cited

Australian Biological Control. 2009. Good bugs. The Australian Biological Control Association Inc. (ABC). Avilla, J., R. Albajes, O. Alomar, C. Castane, and R. Gabarra. 2004. Biological control of whiteflies on vegetable crops, pp. 171-184. In K. M. Heinz, R. G. van Driesche and M. P. Parrella [eds.], Biocontrol in protected culture. Ball Publishing, Batavia, Illinois. Baez, I., S. R. Reitz, and J. E. Funderburk. 2004. Predation by Orius insidiosus (Heteroptera: Anthocoridae) on life stages and species of Frankliniella flower thrips (Thysanoptera: Thripidae) in pepper flowers. Environmental Entomology. 33: 662-670. Blockmans, K. J. F., and A. N. M. Tetteroo. 2002. Biological pest control in eggplants in the Netherlands. IOBC/WPRS Bulletin 13: 71-75. Blümel, S. 2004. Biological control of aphids on vegetable crops, pp. 297-312. In K. M. Heinz, R. G. van Driesche and M. P. Parrella [eds.], Biocontrol in protected culture. Ball Publishing, Batavia, IL. Brødsgaard, H. F. 2004. Biological control of thrips on ornamental crops, pp. 253-264. In K. M. Heinz, R. G. van Driesche and M. P. Parrella [eds.], Biocontrol in protected culture. Ball Publishing, Batavia, Illinois. Brødsgaard, H. F., and A. Enkegaard. 2005. Intraguild predation between Orius majusculus (Reuter) (Hemiptera: Anthocoridae) and Iphiseius degenerans Berlese (Acarina: Phytoseiidae). I.B.O.C./W.P.R.S. Bulletin 28: 19-22. California Strawberry Commission. 2009. California strawberries: A healthy indulgence. California Strawberry Commission, California. Chau, A., and K. M. Heinz. 2004. Biological control of aphids on ornamental crops, pp. 277- 295. In K. M. Heinz, R. G. van Driesche and M. P. Parrella [eds.], Biocontrol in protected culture. Ball Publishing, Batavia, Illinois. Chow, A., and K. M. Heinz. 2004. Biological control of leafminers on ornamental crops, pp. 221-238. In K. M. Heinz, R. G. van Driesche and M. P. Parrella [eds.], Biocontrol in protected culture. Ball Publishing, Batavia Illinois. Chow, A., A. Chau, and K. M. Heinz. 2008. Compatibility of Orius insidiosus (Hemiptera: Anthocoridae) with Amblyseius (Iphiseius) degenerans (Acari: Phytoseiidae) for control of Frankliniella occidentalis (Thysanoptera: Thripidae) on greenhouse roses. Biological control 44: 259-270. Cleveland, C. B., G. A. Bormett, D. G. Saunders, F. L. Powers, A. S. McGibbon, G. L. Reeves, L. Rutherford, and J. L. Balcer. 2002. Environmental fate of spinosad. 1. Dissipation and degradation in aqueous systems. Journal of Agricultural and Food Chemistry 50: 3244-3256. Cote, K. W., P. B. Schultz, and E. E. Lewis. 2004. Using acaricides in combination with Phytoseiulus persimilis Athias-Henriot to suppress Tetranychus urticae Koch populations. Journal of Entomological Science 39: 267-274. Funderburk, J. 2009. Management of the western flower thrips (Thysanoptera: Thripidae) in fruiting vegetables. Florida Entomologist 92: 1-6. Funderburk, J. E., J. Stavisky, and S. Olson. 2000. Predation of Frankliniella occidentalis (Thysanoptera: Thripidae) in field peppers by Orius insidiosus (Hemiptera: Anthocoridae). Environmental Entomology 29: 376-382. Gillespie, D. R., and C. A. Ramey. 1988. Life history and cold storage of Amblyseius cucumeris (Acarina: Phytoseiidae). Journal of Entomological Society of British Columbia 85: 71-76. Gillespie, D. R., and D. J. M. Quiring. 1992. Competition between Orius tristicolor (White) (Hemiptera: Anthocoridae) and Amblyseius cucumeris (Oudemans) (Acari: Phytoseiidae) on Frankliniella occidentalis (Pergande) (Thysanoptera: Thripidae). The Canadian Entomologist 124: 1123-1128. Herron, G., M. Steiner, B. Gollnow, and S. Goodwin. 2007. Western flower thrips (WFT) insecticide resistance management plan. New South Wales Department of Primary Industries.

195 Chapter VIII Summary

Hoddle, M. S. 2004. Biological control of whiteflies on ornamental crops, pp. 149-170. In K. M. Heinz, R. G. van Driesche and M. P. Parrella [eds.], Biocontrol in protected culture. Ball Publishing, Batavia, Illinois. Jones, T., C. Scott-Dupree, R. Harris, L. Shipp, and B. Harris. 2005. The efficacy of spinosad against the western flower thrips, Frankliniella occidentalis, and its impact on associated biological control agents on greenhouse cucumbers in southern Ontario. Pest Management Science 61: 179-185. Kunkel, B. A., D. W. Held, and D. A. Potter. 2001. Lethal and sublethal effects of bendiocarb, halofenozide and imidacloprid on Harpalus pennsylvanicus (Coleoptera: Carabidae) following different modes of exposure in turf grass. Journal of Economic Entomology 94: 60-67. Ludwig, S. W. 2002. Impact of spinosad on Orius insidiosus populations on greenhouse Marigolds, pp. 3, First floriculture industry research and scholarship trust. Texas A & M Agricultural Research and Extension Centre Overton, Tx and Kelli Hoover, Department of Entomology, The Pennsylvania State University, Pennsylvania. Ludwig, S. W., and R. D. Oetting. 2001. Effect of spinosad on Orius insidiosus (Hemiptera: Anthocoridae) when used for Frankliniella occidentalis (Thysanoptera: Thripidae) control on greenhouse pot chrysanthemums. Florida Entomologist 84: 311-313. Magalhăes, S., C. Tudorache, M. Montserrat, R. van Maanen, M. W. Sabelis, and A. Janssen. 2004. Diet of intraguild predators affects antipredator behaviour in intraguild prey. Behavioural Ecology 16: 364-370. Parrella, M. P., and T. Lewis. 1997. Integrated pest management (IPM) in field crops, pp. 595-638. In T. Lewis [ed.], Thrips as crop pests. CAB International, Wallingford, Oxon, UK. Phillips, D., and A. Reid. 2008. New strawberry varieties for WA - trial results in 2005, 2006 and 2007, pp. 9. Department of Agriculture and Food Western Australia, Kensington, South Perth WA. Premachandra, W. T. S. D., C. Borgemeister, O. Berndt, and R.-U. Ehilers. 2003. Combined release of entomopathogenic nematodes and the predatory mite Hypoaspis aculeifer to control soil-dwelling stages of western flower thrips Frankliniella. Biocontrol 48: 529-541. Ramakers, P. M. J. 1988. Population dynamics of the thrips predators Amblyseius mckenziei and Amblyseius cucumeris (Acarina: Phytoseiidae) on sweet pepper. Netherland Journal of Agricultural Science 36: 247-252. Sanderson, J. P., H. F. Brødsgaard, and A. Enkegaard. 2005. Preference assessment of two Orius spp for Neoseiulus cucumeris vs Frankliniella occidentalis, pp. 221-224, I.B.O.C./W.P.R.S. Bulletin. Schoonhoven, L. M., T. Jermy, and J. J. A. van Loon. 1998. Insect-Plant Biology. Chapman & Hall, London. Shipp, J. L., and P. M. J. Ramakers. 2004. Biological control thrips on vegetable crops, pp. 265-276. In K. M. Heinz, R. G. van Driesche and M. P. Parrella [eds.], Biocontrol in protected culture. Ball Publishing, Batavia, Illinois. Skirvin, D. J., L. Kravar-Garde, K. Reynolds, J. Jones, J. J. Reynolds, and M. E. de Courcy Williams. 2006. The influence of pollen on combining predators to control Frankliniella occidentalis in ornamental chrysanthemum crops. Biocontrol Science and Technology 16: 99-105. Steiner, M. Y., and S. Goodwin. 2002. Development of a new thrips predator, Typhlodromips montdorensis (Schicha) (Acari: Phytoseiidae) indigenous to Australia. Bulletin of IOBC/WPRS 25: 245-247. Steiner, M. Y., and S. Goodwin. 2005. Management of western flower thrips, Frankliniella occidentalis (Pergande) (Thysanoptera: Thripidae), in hydroponic strawberry crops: using yellow sticky traps to determine action thresholds. Australian Journal of Entomology 44: 288-292. Steiner, M. Y., S. Goodwin, T. M. Wellham, I. M. Barchia, and L. J. Spohr. 2003. Biological studies of the Australian predatory mite Typhlodromips montdorensis (Schicha) (Acari: Phytoseiidae), a potential biocontrol agent for western flower thrips,

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Frankliniella occidentalis (Pergande) (Thysanoptera: Thripidae). Australian Journal of Entomology 42: 124-130. Thoeming, G., and H.-M. Poehling. 2006. Integrating soil-applied azadiractin with Amblyseius cucumeris (Acari: Phytoseiidae) and Hypoaspis miles (Acari: Laelapidae) for the management of Frankliniella occidentalis (Thysanoptera: Thripidae). Environmental Entomology 35: 746-756. Thompson, D. G., B. J. Harris, L. J. Lanteigne, T. M. Buscarini, and D. T. Chartrand. 2002. Fate of spinosad in litter and soils of a mixed conifer stand in the Acacian forest region of New Brunswick. Journal of Agricultural and Food Chemistry 50: 790-795. Thompson, G., and S. Hutchins. 1999. Spinosad. Pesticide Outlook 10: 78-81. Thompson, G. D., R. Dutton, and T. C. Sparks. 2000. Spinosad-a case study: an example from natural products discovery programme. Pest Management Science 56: 696-702. van de Veire, M., M. Klein, and L. Tirry. 2002. Residual activity of abamectin and spinosad against the predatory bug Orius laevigatus. Phytoparasitica 30: 525-528. Wellington, W. G. 1977. Returning the insect to insect ecology: some consequences for pest management. Environmental Entomology 6: 1-8. White, T. C. R. 1969. An index to measure weather-induced stress of trees associated with outbreaks of psyllids in Australia. Ecology 50: 905-909. White, T. C. R. 1993. The inadequate environment: nitrogen and the abundance of animals. Springer Verlag, New York. Williams, T., J. Valle, and E. Vinuela. 2003. Is the naturally derived insecticide Spinosad compatible with insect natural enemies? Biocontrol Science and Technology 13: 459- 475. Wittmann, E. J., and S. R. Leather. 1997. Compatibility of Orius laevigatus Fieber (Hemiptera: Anthocoridae) with Neoseiulus (Amblyseius) cucumeris Oudemans (Acari: Phytoseiidae) and Iphiseius (Amblyseius) degenerans Berlese (Acari: Phytoseiidae) in the biocontrol of Frankliniella occidentalis Pergande (Thysanoptera: Thripidae). Experimental and Applied Acarology 21: 523-538. Zilahl-Balogh, G. M. G., J. L. Shipp, C. Cloutier, and J. Brodeur. 2007. Predation by Neoseiulus cucumeris on western flower thrips, and its oviposition on greenhouse cucumber under winter vs. summer conditions in a temperate climate. Biological Control 40: 160-167.

197 APPENDICS

Appendix 3.1 Summary of repeated measures ANOVAs tested the effect of cultivars and predatory mites with or without spinosad on mean numbers of WFT adults per plant at days 7, 14, and 21.

Source df (Num & Den) F value P value All cultivars Cultivar 2, 216 2245.37 <0.0001 Spray 1, 216 4062.00 <0.0001 Mites release 3, 216 516.31 <0.0001 Day 2, 432 4776.66 <0.0001 Cultivar*spray 2, 216 3.28 0.0395 Cultivar*mites release 6, 216 5.99 <0.0001 Cultivar*day 4, 432 14.14 <0.0001 Spray*mites release 3, 216 21.93 <0.0001 Spray*day 2, 432 256.04 <0.0001 Mites release*day 6, 432 144.53 <0.0001 Cultivar*spray*mites release 6, 216 2.38 0.0303 Cultivar*Spray*day 4, 432 14.64 <0.0001 Cultivar*mites release*day 12, 432 4.88 <0.0001 Spray*mites release*day 6, 432 7.32 <0.0001 Cultivar*spray*mites release*day 12, 432 9.26 <0.0001

Camarosa Spray 1, 72 1191.60 <0.0001 Mites release 3, 72 102.90 <0.0001 Day 2, 144 1600.00 <0.0001 Spray*mites release 3, 72 6.62 0.0005 Spray*day 2, 144 21.43 <0.0001 Mites release*day 6, 144 38.83 <0.0001 Spray*mites release*day 6, 144 9.75 <0.0001

Camino Real Spray Mites release 1, 72 1651.47 <0.0001 Day 3, 72 2209.65 <0.0001 Spray*mites release 2, 144 1241.62 <0.0001 Spray*day 3, 72 9.77 <0.0001 Mites release*day 2, 144 111.28 <0.0001 Spray*mites release*day 6, 144 45.93 <0.0001 6, 144 4.29 0.0005 Albion Spray 1, 72 1259.91 <0.0001 Mites release 3, 72 227.12 <0.0001 Day 2, 144 2107.92 <0.0001 Spray*mites release 3, 72 10.61 <0.0001 Spray*day 2, 144 185.13 <0.0001 Mites release*Day 6, 144 79.63 <0.0001 Spray*mites release*day 6, 144 13.02 <0.0001

xvii Appendix 3.2 Summary of repeated measures ANOVAs tested the effect of cultivars and predatory mites with or without spinosad on mean numbers of WFT larvae per plant at days 7, 14, and 21.

Source df (Num & Den) F value P value All cultivars Cultivar 2, 216 447.65 <0.0001 Spray 1, 216 143.41 <0.0001 Mites release 3, 216 181.43 <0.0001 Day 2, 432 254.80 <0.0001 Cultivar*spray 2, 216 241.03 <0.0001 Cultivar*mites release 6, 216 2.81 0.0457 Cultivar*day 4, 432 51.50 <0.0001 Spray*mites release 3, 216 3.40 0.0186 Spray*day 2, 432 9.17 0.0001 Mites release*day 6, 432 22.80 <0.0001 Cultivar*spray*mites release 6, 216 4.96 <0.0001 Cultivar*Spray*day 4, 432 15.43 <0.0001 Cultivar*mites release*day 12, 432 1.20 0.2782 Spray*mites release*day 6, 432 2.91 0.0087 Cultivar*spray*mites release*day 12, 432 2.31 0.0073

Camarosa Spray 1, 72 62.75 <0.0001 Mites release 3, 72 74.66 <0.0001 Day 2, 144 17.62 <0.0001 Spray*mites release 3, 72 6.78 0.0004 Spray*day 2, 144 9.32 0.0002 Mites release*day 6, 144 12.45 <0.0001 Spray*mites release*day 6, 144 3.78 0.0010

Camino Real Spray 1, 72 556.01 <0.0001 Mites release 3, 72 48.08 <0.0001 Day 2, 144 52.67 <0.0001 Spray*mites release 3, 72 4.35 0.0071 Spray*day 2, 144 19.52 <0.0001 Mites release*day 6, 144 5.55 <0.0001 Spray*mites release*day 6, 144 2.56 0.0217

Albion Spray 1, 72 22.40 <0.0001 Mites release 3, 72 63.53 <0.0001 Day 2, 144 264.96 <0.0001 Spray*mites release 3, 72 2.60 0.0590 Spray*day 2, 144 10.88 <0.0001 Mites release*Day 6, 144 7.73 <0.0001 Spray*mites release*day 6, 144 2.76 0.0114

xviii Appendix 4.1 Summary of repeated measures ANOVAs tested the effect of cultivars and predatory mites (combined application) with or without spinosad on mean numbers of WFT (adults and larvae) on days 6, 9, 12, 15, 18, and 21.

Source df (Num & Den) F value P value WFT adults 1, 144 5129.93 <0.0001 Spray 7, 144 585.99 <0.0001 Mites release 5, 720 855.30 <0.0001 Day 7, 144 14.06 <0.0001 Spray*mites release 5, 720 10.47 <0.0001 Spray*day 35, 720 56.53 <0.0001 Mites release*day 35, 720 2.52 <0.0001 Spray*mites release*day Spinosad Mites release 7, 72 193.12 <0.0001 Day 5, 360 312.30 <0.0001 Mites release*day 35, 360 23.94 <0.0001 Water Mites release 7, 72 469.60 <0.0001 Day 5, 360 596.93 <0.0001 Mites release*day 35, 360 37.21 <0.0001

WFT larvae Spray 1, 144 2964.87 <0.0001 Mites release 7, 144 134.16 <0.0001 Day 5, 720 46.62 <0.0001 Spray*mites release 7, 144 19.86 <0.0001 Spray*day 5, 720 77.28 <0.0001 Mites release*day 35, 720 9.64 <0.0001 Spray*mites release*day 35, 720 1.75 0.0052 Spinosad Mites release 7, 72 46.71 <0.0001 Day 5, 360 14.13 <0.0001 Mites release*day 35, 360 6.64 <0.0001 Water Mites release 7, 72 90.18 <0.0001 Day 5, 360 88.97 <0.0001 Mites release*day 35, 360 5.21 <0.0001

xix Appendix 5.1 Summary of ANOVA results for WFT on flower and fruit of strawberry grown in low tunnels before spray/mite treatments.

Source of variations Num df Den df F value P value Adults Flower Tunnel (between) 3 21 2.58 0.0815 Mites treatment (within tunnel) 7 21 1.17 0.3607 Times (weeks) 2 62 6.24 0.0034 Fruit Tunnel (between) 3 21 1.90 0.1606 Mites treatment (within tunnel) 7 21 1.21 0.3404 Times (weeks) 2 62 5.21 0.0088 Nymphs Flower Tunnel (between) 3 21 1.14 0.3559 Mites treatment (within tunnel) 7 21 1.12 0.3876 Times (weeks) 2 62 2.29 0.0385 Fruit Tunnel (between) 3 21 2.11 0.1291 Mites treatment (within tunnel) 7 21 1.53 0.2114 Times (weeks) 2 62 2.64 0.0187

xx Appendix 5.2 Summary of ANOVA results for WFT adults on flower and fruit (split-plot repeated measures ANOVA) of strawberry grown in low tunnels.

Source of variations Num df Den df F value P value Flowers Tunnel 3 6 0.90 0.4942 Spray regime 2 6 361.64 <0.0001 Predatory mites 7 63 344.31 <0.0001 Times (weeks) 4 288 564.74 <0.0001 Spray regime*predatory mites 14 63 10.73 <0.0001 Spray regime*times 8 288 110.72 <0.0001 Predatory mites*times 28 288 50.66 <0.0001 Spray regime*predatory mites*times 56 288 18.93 <0.0001 Water spray Tunnel 3 21 2.13 <0.0001 Predatory mites 7 21 145.73 <0.0001 Times (weeks) 4 96 331.68 <0.0001 Predatory mites*Times 28 96 32.71 <0.0001 Spinosad sprayed then mites released Tunnel 3 21 1.97 0.1494 Predatory mites 7 21 765.74 <0.0001 Times (weeks) 4 96 40.44 <0.0001 Predatory mites*times 28 96 101.42 <0.0001 Mites released then spinosad sprayed Tunnel 3 21 1.88 0.1640 Predatory mites 7 21 41.57 <0.0001 Times (weeks) 4 96 562.37 <0.0001 Predatory mites*times 28 96 7.04 <0.0001

Fruits Tunnel 3 6 1.72 0.2617 Spray regime 2 6 698.13 <0.0001 Predatory mites 7 63 130.16 <0.0001 Times (weeks) 4 288 70.61 <0.0001 Spray regime*predatory mites 14 63 9.63 <0.0001 Spray regime*times 8 288 118.24 <0.0001 Predatory mites*times 28 288 26.03 <0.0001 Spray regime*predatory mites*times 56 288 3.78 <0.0001 Water spray Tunnel 3 21 2.88 0.0602 Predatory mites 7 21 13.62 <0.0001 Times (weeks) 4 96 1.98 0.1029 Predatory mites*times 28 96 3.22 <0.0001 Spinosad sprayed then mites released Tunnel 3 21 0.67 0.5789 Predatory mites 7 21 284.76 <0.0001 Times (weeks) 4 96 74.93 <0.0001 Predatory mites*Times 28 96 32.59 <0.0001 Mites released then spinosad sprayed Tunnel 3 21 0.70 0.5607 Predatory mites 7 21 85.62 <0.0001 Times (weeks) 4 96 103.29 <0.0001 Predatory mites*times 28 96 47.34 <0.0001

xxi Appendix 5.3 Summary of repeated measures ANOVA results for WFT larvae on flower and fruits of strawberry grown in low tunnels.

Source of variations Num df Den df F value P value Flowers Tunnel 3 6 1.80 0.1781 Spray regime 2 6 150.17 <0.0001 Predatory mites 7 63 430.84 <0.0001 Times (weeks) 4 288 569.23 <0.0001 Spray regime*predatory mites 14 63 20.77 <0.0001 Spray regime*times 8 288 24.03 <0.0001 Predatory mites*times 28 288 274.06 <0.0001 Spray regime*predatory mites*times 56 288 37.11 <0.0001 Water spray Tunnel 3 21 2.85 0.0619 Predatory mites 7 21 332.62 <0.0001 Times (weeks) 4 96 407.16 <0.0001 Predatory mites*times 28 96 276.74 <0.0001 Spinosad sprayed then mites released Tunnel 3 21 2.53 0.0848 Predatory mites 7 21 93.85 <0.0001 Times (weeks) 4 96 154.91 <0.0001 Predatory mites*times 28 96 86.57 <0.0001 Mites released then spinosad sprayed Tunnel 3 21 1.24 0.3203 Predatory mites 7 21 219.91 <0.0001 Times (weeks) 4 96 191.86 <0.0001 Predatory mites*times 28 96 157.72 <0.0001

Fruits Tunnel 3 6 0.15 0.9279 Spray regime 2 6 104.20 <0.0001 Predatory mites 7 63 483.08 <0.0001 Times (weeks) 4 288 303.37 <0.0001 Spray regime*predatory mites 14 63 25.43 <0.0001 Spray regime*times 8 288 59.89 <0.0001 Predatory mites*times 28 288 53.76 <0.0001 Spray regime*predatory mites*times 56 288 9.15 <0.0001 Water spray Tunnel 3 21 1.91 0.1583 Predatory mites 7 21 175.90 <0.0001 Times (weeks) 4 96 13.55 <0.0001 Predatory mites*times 28 96 107.71 <0.0001 Spinosad sprayed then mites released Tunnel 3 21 1.83 0.1726 Predatory mites 7 21 238.96 <0.0001 Times (weeks) 4 96 233.93 <0.0001 Predatory mites*times 28 96 31.33 <0.0001 Mites released then spinosad sprayed Tunnel 3 21 1.19 0.3370 Predatory mites 7 21 113.75 <0.0001 Times (weeks) 4 96 103.60 <0.0001 Predatory mites*times 28 96 24.23 <0.0001

xxii Appendix 5.4 ANOVA results for predatory mites, T. montdorensis and N. cucumeris per flower/fruit when applied in different combinations in different spray treatments.

Source of variations Num df Den df F value P value Flower Tunnel 3 6 3.06 0.1131

Mite species 1 11 1.42 0.2592 Spray 2 6 38.63 0.0004 Times (tunnel) 3 63 1.55 0.2097 Spray*times 6 63 0.97 0.4551 Fruit Tunnel 3 6 1.05 0.4358 Single sp release sp Single Mite species 1 11 29.39 0.0002 Spray 2 6 299.7 0.0008 Times (tunnel) 3 63 2.58 0.0615 Spray*times 6 63 0.43 0.8562 Flower Tunnel 3 6 0.58 0.6497 Mite species 1 11 10.53 0.0078 Spray 2 6 11.95 0.0081 Times (tunnel) 3 63 1.89 0.1404 Spray*times 6 63 0.22 0.9680 Fruit Tunnel 3 6 1.24 0.3753 Mite species 1 11 9.99 0.0092 Spray 2 6 13.95 0.0062

Double sp release (Tm*Nc) sp release Double Times (tunnel) 3 63 1.28 0.2882 Spray*times 6 63 0.07 0.9986 Flower Tunnel 3 6 0.62 0.6179 Mite species 1 11 4.58 0.0556 Spray 2 6 30.70 0.0007

Times (tunnel) 3 63 1.58 0.2040 Spray*times 6 63 0.50 0.8089 Fruit

Nc*Hm) Tunnel 3 6 1.32 0.3259 Mite species 1 11 4.69 0.0534 Spray 2 6 21.41 0.0019 Times (tunnel) 3 63 1.83 0.1516 Double sp release (Tm*Hm, (Tm*Hm, sp release Double Spray*times 6 63 0.90 0.4992

Flower Tunnel 3 6 0.57 0.6556 Mite species 1 11 0.15 0.7070 Spray 2 6 8.04 0.0201 Times (tunnel) 3 63 0.18 0.9121 Spray*times 6 63 0.08 0.9982 Fruit Tunnel 3 6 1.35 0.3454 Mite species 1 11 0.15 0.7078 Spray 2 6 8.03 0.0201 Times (tunnel) 3 63 0.22 0.8801

Triple sp release (Tm*NC*Hm) release sp Triple Spray*times 6 63 0.09 0.9974 *indicates species combinations. Tm = T. montdorensis, Nc = N. cucumeris, Hm = H. miles.

xxiii Appendix 5.5 Repeated measures ANOVA results of the effects of spray, mite species combination (double) and time (weeks) on T. montdorensis (when applied with N. cucumeris and H. miles) and N. cucumeris (when applied with T. montdorensis and H. miles) in double- species combinations (Repeated measures ANOVA with split-plot design).

Source of variations Num df Den df F value P value T. montdorensis Flowers Tunnel 3 6 0.20 0.8954 Spray 2 6 15.88 0.0040 Mite combination 1 9 48.66 <0.0001 Times (tunnel) 3 54 2.41 0.0770 Spray*mite combination 2 9 2.93 0.1047 Spray*times 6 54 0.43 0.8557 Mite combination*Times 3 54 0.46 0.7093 Spray*mite combination*times 6 54 0.60 0.7283 Fruits Tunnel 3 6 0.09 0.9613 Spray 2 6 8.41 0.0182 Mite combination 1 9 11.46 0.0081 Times (tunnel) 3 54 2.12 0.1088 Spray*mite combination 2 9 0.15 0.8656 Spray*times 6 54 0.25 0.9555 Mite combination*times 3 54 0.04 0.9903 Spray*mite combination*times 6 54 0.08 0.9979

N. cucumeris Flowers Tunnel 3 6 1.39 0.3332 Spray 2 6 11.76 0.0084 Mite combination 1 9 20.43 0.0014 Times (tunnel) 3 54 2.34 0.0836 Spray*mite combination 2 9 2.73 0.1183 Spray*times 6 54 0.20 0.9766 Mite combination*times 3 54 0.08 0.9725 Spray*mite combination*times 6 54 0.31 0.9300 Fruits Tunnel 3 6 1.53 0.2467 Spray 2 6 11.57 0.0009 Mite combination 1 9 14.42 0.0018 Times (tunnel) 3 54 0.79 0.5073 Spray*mite combination 2 9 1.03 0.3799 Spray*times 6 54 0.59 0.7368 Mite combination*times 3 54 0.35 0.7927 Spray*mite combination*times 6 54 0.50 0.8055

xxiv Appendix 7.1 ANOVA results showing the interaction of spray (spinosad, water) and predatory mite (no mites, T. montdorensis, N. cucumeris, H. miles) on WFT adults and larvae.

Obs. Source F df P Adults 10 DAS 16.75 < 0.0001 15 DAS 6.96 0.0010 20 DAS 4.75 0.0075 25 DAS 3.28 0.0333 30 DAS 8.45 0.0003 35 DAS 14.50 < 0.0001 Spray*mites 3, 32 Larvae 10 DAS 38.43 < 0.0001 15 DAS 7.86 0.0005 20 DAS 7.89 0.0004 25 DAS 17.99 < 0.0001 30 DAS 26.97 < 0.0001 35 DAS 12.16 < 0.0001

xxv Appendix 7.2 ANOVA results of the effects of predatory mites (no mites, T. montdorensis, N. cucumeris, H. miles) on WFT adults and larvae in spinosad- and water-treated plants.

Obs. Source F df P Adults Spinosad 10 DAS 21.70 < 0.0001 15 DAS 554.34 < 0.0001 20 DAS 51.72 < 0.0001 25 DAS 30.85 < 0.0001 30 DAS 56.80 < 0.0001 35 DAS 112.26 < 0.0001 Water 10 DAS 2.38 0.1082 15 DAS 70.70 < 0.0001 20 DAS 58.51 < 0.0001 25 DAS 48.61 < 0.0001 30 DAS 38.88 < 0.0001 35 DAS 104.33 < 0.0001 Predatory mites 3, 16 Larvae release Spinosad 10 DAS 137.59 < 0.0001 15 DAS 31.69 < 0.0001 20 DAS 25.89 < 0.0001 25 DAS 213.26 < 0.0001 30 DAS 61.49 < 0.0001 35 DAS 136.26 < 0.0001 Water 10 DAS 25.64 < 0.0001 15 DAS 47.78 < 0.0001 20 DAS 59.92 < 0.0001 25 DAS 69.59 < 0.0001 30 DAS 51.21 < 0.0001 35 DAS 89.75 < 0.0001

xxvi