An autodissemination strategy using entomopathogenic fungi and kairomonal attractants for managing thrips on grain legumes BK Mfuti 25076558 Thesis submitted for the degree Philosophiae Doctor in Environmental Sciences at the Potchefstroom Campus of the North-West University Promoter: Prof MJ du Plessis Co-promoter: Dr NK Maniania Assistant Promoter: Dr S Subramanian May 2016 i DEDICATION To my beloved wife Hermane Julienne Longi and my two sons Divin-Regis Kupesa and Joyce- Marlon Mfuti for their constant affection and motivation, To my father Kupesa, my mother Kolingila and the entire Kupesa family for their encouragement and moral support, I dedicate this dissertation. ii iii ACKNOWLEDGEMENTS My sincere thanks are addressed to Prof. Magdalena Johanna du Plessis for accepting to be my university supervisor. I really appreciate your invaluable contribution and guidance during the execution of this research project. Your criticism, remarks and kind advice have strongly guided me to become rigorous. I sincerely appreciate the way we interacted. It has given me confidence and strength for the accomplishment of this thesis. I am grateful to Dr. Nguya Kalemba Maniania for all the scientific knowledge especially in arthropod pathology that I have learnt from you. Your critical remarks and criticism have given me strength and showed me how to work hard. I deeply appreciate your mentorship and support in multiple ways for the achievement of this research project. Your door was always open for me for any discussion. I am very confident for my future career and I am very proud to be able to work with you. I also express my sincere thanks to Dr. Sevgan Subramanian for his invaluable suggestions, constructive advice and guidance during this research project. I deeply appreciate your mentorship and support in multiple ways for the achievement of this research project. That support has strengthened me from the beginning to the end of the present study. Further gratitude goes to Dr. Saliou Niassy for his guidance. You have shown me how to work hard and overcome difficulties. Your critical advice and support have helped me to fill quickly the gap on my adaptation to Anglophone world. iv I express my gratitude to the Biostatistics Unit, especially to Dr. Daisy Salifu for giving guidance on statistical data analysis, for which I am very grateful. I am very grateful to the International Centre of Insect Physiology and Ecology (icipe) who gave me this opportunity on behalf of the African Regional Postgraduate Program in Insect Science (ARPPIS) network partners, funded by the German Academic Exchange Service (DAAD). I sincerely acknowledge the financial support from the African Union (AU) through the African Union Research Grant Contract No. AURG/108/2012 and the German Federal Ministry for Economic Cooperation and Development (BMZ) through the grant Project number: 11.7860.7- 001.00, Contract number: 81141840 for the accomplishment of this work. I thank all colleagues and friends for their encouragement and support during my study, especially David Cham Tembong, Bayissa Wakuma, Tigist Tolosa Asefa, Andnet B., Tumahise Venasio, Ange Toe, Alex Muvea, Eric Ntiri, Rosaline Macharia, Briget Babadoe, Yvonne Ukamaka, David Omondi, San Pedro, Soul Midengoyi, Edoh Kokum, Thomas Franck, Caroline Foba, Dr. Johnson Nyasani, Dr. Akutse; Dr. Paulin Nana, Dr. David Tchouassi, Dr. Tanga Mbi. My special thanks to Dr. Didi Kiatoko and his entire family for their invaluable assistance and support. I am also grateful to my colleagues from my home research institute Institut National pour l’Etudes et la Recherche Agronomiques (INERA). I thank all the staff from the Thrips project, Arthropod Pathology Unit (APU) and Capacity Building for providing me with all the facilities needed during my study. I particularly thank Dr. v Rob Skilton, Lillian Igweta, Levi Odhiambo, Caroline Akal, Bernard Muia, Barbara Obonyo, Catherine Adongo, Peris Kariuki, Bernard Mulwa, Emmanuel Mlato, Alex Irungu Maina, Josua Matuku, Jane Kimemia, Lisa Omondi, and Mama Maggy Ochanda. Finally, I would like to express my gratitude to all my family members, particularly my parents Mr. Kupesa and Mrs Kolingila, my brothers (Odon Kupesa, Michel Kupesa, Lady Tuangaliye, Toussaint Kupesa, Jean Jules Kupesa, Antoine Kupesa and Junior Kupesa) and all my nephews and nieces who always supported and encourage me. God bless you all. vi ABSTRACT Grain legumes are among the key economical crops widely grown in western and eastern Africa as important sources of food and animal fodder. However, the production of grain legumes in Kenya is seriously affected by a complex of insect pests particularly thrips. Yield losses of 20 to 100% have often been reported in some areas. The bean flower thrips (BFT), Megalurothrips sjostedti is considered to be the most important thrips pest of grain legumes. Chemical control is still the main management strategy, with detrimental consequences on the environment, users and consumers. Entomopathogenic fungi (EPF) are among the most promising alternatives to chemical pesticides. Inundative sprays are the most common application techniques for EPF. Although efficient and environmentally safe, the performance of entomopathogenic fungi is affected by several environmental parameters such as UV light, temperature, drought and rain. In order to improve the efficacy of EPF, an autodissemination system has been developed for the management of thrips in greenhouses. In this system, thrips are attracted to an autoinoculator where they are infected with an EPF before returning to the environment to disseminate the EPF to conspecifics. It therefore provides promising prospects, but for effective control, the conidial persistence and thrips attraction need to be optimized, while the EPF and the semiochemical should be compatible. The objective of this study was therefore to optimize the autodissemination system for thrips management on grain legumes in Kenya. The semiochemical Lurem-TR, has been found to inhibit conidia of EPF when put together in an autoinoculation device. The effect of spatial separation of Lurem-TR on the persistence of conidia of EPF, Metarhizium brunneum and Metarhizium anisopliae was therefore evaluated to vii develop an autodissemination strategy for the management of M. sjostedti. Influence of spatial separation of the semiochemical on thrips attraction and conidial acquisition by thrips from the autoinoculation device was also investigated in the field. This study showed that conidia persistence of both fungal species increased with distance of separation from Lurem-TR. Attraction of thrips to the device also varied significantly according to distance between the device and semiochemical. More thrips were attracted when Lurem-TR was placed in a container below the device and at 10 cm distance from the device. Conidial acquisition by thrips was not significantly different between spatial separation treatments of conidia and Lurem-TR. Seven alternative thrips attractants, namely 4-anisaldehyde, ethyl benzoate, cis-jasmone, linalool, methyl anthranilate, trans-caryophyllene and phenylethanol were also screened for their compatibility with M. anisopliae ICIPE 69 in autodissemination devices and for their attraction to M. sjostedti in the field. Methyl anthranilate (MA) was found to be the attractant most compatible with M. anisopliae and its attractiveness to M. sjostedti was similar to that of Lurem- TR. The performance of the attractant, methyl anthranilate, was compared to the commercial attractant Lurem-TR in autoinoculation devices treated with M. anisopliae under field conditions for two seasons. Densities of M. sjostedti in plots with the two semiochemical-baited autoinoculation devices were less than in the control plots during both experimental seasons. Plots with MA-baited and Lurem-TR-baited devices had similar densities of M. sjostedti during both seasons. However in the second season thrips densities in plots with the Lurem-TR-baited devices did not differ significantly from the control plots. Conidial viability of M. anisopliae was viii significantly higher in semiochemical-free baited devices (control) than in semiochemical-baited devices in both seasons. Conidial germination decreased over time in all the treatments but remained above 45%, 12-15 days post-exposure. The average number of conidia acquired by a single M. sjostedti ranged between 2.0 and 10.0 x 103 conidia in both semiochemical-baited device treatments during both seasons. Significantly more conidia were acquired by single thrips in MA-baited devices compared to Lurem-TR baited devices during the podding stage of the crop during the second season. Significantly higher mortality of M. sjostedti was caused in field plots by Lurem-TR baited and MA-baited autoinoculation devices compared to mortality of M. sjostedti collected from the control plots in both seasons. Cowpea yield also differed significantly between the treatment plots. The highest yield was recorded in plots where MA- baited devices were placed. From this study, it could therefore be recommended that methyl anthranilate be used in autoinoculation devices for the management of M. sjostedti on grain legumes. The success achieved with MA in these trials resulted in the evaluation of this EPF for possible use in a spot spray strategy. The efficacy of spot spray and cover spray applications of M. anisopliae in combination with the thrips attractant Lurem-TR was compared in field experiments for the management of M. sjostedti on a cowpea crop in two seasons. Plants in the treatment plots where a spot spray application of M. anisopliae was done five days after the placement of Lurem-TR recorded the lowest densities of M. sjostedti. Fungal viability and thrips conidial acquisition did not differ between the two application methods. Compared to the control treatment plots, both application strategies resulted in yield increases of 34.1 and 46.2% with spot and cover spray treatments, ix respectively. The cost benefit analysis suggests that the spot spray application was more profitable due to the reduction in labour and the quantity of inoculum used. Key words: Biological control, cowpea, entomopathogenic fungus, grain legumes, lure and infect, Megalurothrips sjostedti, semiochemicals, thrips x TABLE OF CONTENTS DEDICATION................................................................................................................................ i DECLARATION AND APPROVAL.........................................................................................ii ACKNOWLEDGEMENTS ........................................................................................................ iii ABSTRACT .................................................................................................................................. vi TABLE OF CONTENTS ............................................................................................................. x LIST OF TABLES ...................................................................................................................... xv CHAPTER 1: GENERAL INTRODUCTION ........................................................................... 1 1.1. Introduction .................................................................................................................... 1 1.2. Problem statement and justification ............................................................................. 3 1.3. Objectives ........................................................................................................................ 4 1.3.1 General objective ............................................................................................................ 4 1.3.2 Specific objectives .......................................................................................................... 4 1.3.3 Research Hypotheses ...................................................................................................... 5 1.4 References ........................................................................................................................ 5 CHAPTER 2: LITERATURE REVIEW ................................................................................. 10 2.1 Thrips taxonomy and identification ............................................................................ 10 2.2 Geographical Distribution ............................................................................................ 10 2.3 Biology ............................................................................................................................ 12 2.4 Economic importance of thrips .................................................................................... 13 2.5 Control strategies for thrips ......................................................................................... 13 2.5.1 Chemical control........................................................................................................... 13 2.5.2 Intercropping ................................................................................................................ 14 2.5.3 Thrips monitoring and trapping .................................................................................... 14 2.5.4 Semiochemicals ............................................................................................................ 15 2.5.5 Biological control ......................................................................................................... 15 2.5.5.1 Predators .................................................................................................................... 15 2.5.5.2 Parasitoids .............................................................................................................. 16 xi 2.5.5.3 Entomopathogenic fungi ........................................................................................ 16 2.5.5.4 Current strategies for delivery of entomopathogenic fungi in the field ................. 17 2.6 References ...................................................................................................................... 18 CHAPTER 3: SPATIAL SEPARATION OF SEMIOCHEMICAL LUREM-TR AND ENTOMOPATHOGENIC FUNGI TO ENHANCE THEIR COMPATIBILITY AND INFECTIVITY IN AN AUTOINOCULATION SYSTEM FOR THRIPS MANAGEMENT .............................................................................................................. 29 Abstract ................................................................................................................................ 29 3.1 Introduction ................................................................................................................... 30 3.2 Materials and methods .................................................................................................. 32 Study site ................................................................................................................................ 32 Entomopathogenic fungi ........................................................................................................ 33 Semiochemical ....................................................................................................................... 34 3.2.1 Effect of spatial separation of Lurem-TR on the persistence of conidia of Metarhizium brunneum in the greenhouse .................................................................................................. 34 3.2.2 Effect of spatial separation of Lurem-TR on Metarhizium anisopliae conidia persistence in the field ........................................................................................................... 40 3.2.3 Attraction of Megalurothrips sjostedti and other pests ................................................ 40 3.2.4 Conidial acquisition by Megalurothrips sjostedti ........................................................ 40 3.2.5 Statistical analysis......................................................................................................... 41 3.3 Results ............................................................................................................................. 42 3.3.1 Effect of spatial separation of Lurem-TR on conidial viability in the greenhouse ...... 42 3.3.2 Effect of spatial separation of Lurem-TR on conidial viability in the field ................. 43 3.3.3 Effect of spatial separation of Lurem-TR on attraction of Megalurothrips sjostedti ... 49 3.3.4 Effect of spatial separation of Lurem-TR on conidial acquisition by Megalurothrips sjostedti .................................................................................................................................. 53 3.3.5 Effect of spatial separation of Lurem-TR on the attraction of other insects ................ 54 3.4 Discussion ....................................................................................................................... 57 3.5 Conclusion ...................................................................................................................... 59 3.6 References ...................................................................................................................... 60 xii CHAPTER 4: SCREENING OF ATTRACTANTS FOR COMPATIBILITY WITH METARHIZIUM ANISOPLIAE USE IN THRIPS MANAGEMENT ......................... 67 Abstract ................................................................................................................................ 67 4.1 Introduction ................................................................................................................... 68 4.2 Materials and methods .................................................................................................. 69 Study sites .............................................................................................................................. 69 Thrips attractants .................................................................................................................. 69 Crop ....................................................................................................................................... 72 Fungal culture ....................................................................................................................... 72 4.2.1 Effect of thrips attractants on conidial viability of Metarhizium anisopliae ................ 72 4.2.2 Effect of thrips attractants on germ tube length of Metarhizium anisopliae ................ 73 4.2.3 Effect of selected thrips attractants on the attraction of Megalurothrips sjostedti ....... 73 4.2.4 Statistical analysis......................................................................................................... 75 4.3 Results ............................................................................................................................. 75 4.3.1 Effect of thrips attractants on conidial viability of Metarhizium anisopliae ................ 75 4.3.2 Effect of thrips attractants on germ tube length of Metarhizium anisopliae ................ 78 4.3.3 Effect of selected thrips attractants on the attraction of Megalurothrips sjostedti ....... 81 4.4 Discussion ....................................................................................................................... 82 4.5 Conclusion ...................................................................................................................... 84 4.6 References ...................................................................................................................... 85 CHAPTER 5: FIELD EVALUATION OF METHYL ANTHRANILATE AS BAIT FOR MEGALUROTHRIPS SJOSTEDTI IN AUTOINOCULATION DEVICES ............... 92 Abstract ................................................................................................................................ 92 5.1 Introduction ................................................................................................................... 93 5.2 Materials and methods .................................................................................................. 95 Study site ................................................................................................................................ 95 Experimental crop ................................................................................................................. 97 Mass production of the fungus............................................................................................... 95 Semiochemicals ..................................................................................................................... 97 xiii 5.2.1 Effect of semiochemical-baited autoinoculation devices treated with Metarhizium anisopliae on Megalurothrips sjostedti density..................................................................... 97 5.2.2 Effect of semiochemical-baited autoinoculation devices treated with Metarhizium anisopliae on conidial persistence of Metarhizium anisopliae ............................................. 98 5.2.3 Effect of semiochemical-baited autoinoculation devices treated with Metarhizium anisopliae on conidial acquisition and mortality of Megalurothrips sjostedti ...................... 98 5.2.4 Cowpea yield ................................................................................................................ 99 5.2.5 Statistical analysis......................................................................................................... 99 5.3 Results ........................................................................................................................... 100 5.3.1 Effect of semiochemical-baited autoinoculation devices treated with Metarhizium anisopliae on Megalurothrips sjostedti density .................................................................. 100 5.3.2 Effect of semiochemical-baited autoinoculation devices treated with Metarhizium anisopliae on conidial viability of Metarhizium anisopliae ................................................ 103 5.3.3 Effect of semiochemical baited autoinoculation devices treated with Metarhizium anisopliae on conidial acquisition by and mortality of Megalurothrips sjostedti ............... 106 5.3.4 Cowpea yield .............................................................................................................. 106 5.4 Discussion ..................................................................................................................... 109 5.5 Conclusion .................................................................................................................... 111 5.6 References .................................................................................................................... 112 CHAPTER 6: IMPROVING APPLICATION OF FUNGUS-BASED BIOPESTICIDE IN COMBINATION WITH SEMIOCHEMICAL FOR THE MANAGEMENT OF BEAN FLOWER THRIPS ON COWPEA .................................................................. 117 Abstract .............................................................................................................................. 117 6.1 Introduction ................................................................................................................. 118 6.2 Materials and methods ................................................................................................ 119 Study site .............................................................................................................................. 119 The fungus............................................................................................................................ 120 Semiochemical ..................................................................................................................... 120 Experimental design ............................................................................................................ 120 6.2.1 Effect of fungal application strategy on Megalurothrips sjostedti density ................ 121 6.2.2 Effect of fungal application strategy on Metarhizium anisopliae conidial persistence ............................................................................................................................................. 122 xiv 6.2.3 Effect of fungal applications strategy on conidial acquisition ................................... 122 6.2.4 Cowpea yield .............................................................................................................. 123 6.2.5 Cost benefit analysis ................................................................................................... 123 6.2.6 Statistical analysis....................................................................................................... 124 6.3 Results ........................................................................................................................... 124 6.3.1 Effect of fungal application strategy on Megalurothrips sjostedti density ................ 124 6.3.2 Effect of fungal application strategy on Metarhizium anisopliae conidial persistence and Megalurothrips sjostedti conidial acquisition............................................................... 127 6.3.3 Effect of fungal application strategy on conidial acquisition ..................................... 129 6.3.4 Cowpea yield .............................................................................................................. 130 6.4 Discussion ..................................................................................................................... 132 6.5 Conclusion .................................................................................................................... 134 6.6. References ................................................................................................................... 146 CHAPTER 7: GENERAL DISCUSSION, CONCLUSIONS AND RECOMMENDATIONS .......................................................................................................................................... 141 7.1. General discussion ...................................................................................................... 141 7.2. Conclusions ................................................................................................................. 144 7.3. Recommendations ...................................................................................................... 145 7.4. References ................................................................................................................... 146 xv LIST OF TABLES Table 3.1: Repeated measures ANOVA table for the response variable: Metarhizium anisopliae conidial viability (A) and acquisition (B) in autoinoculation devices as affected by spatial separation of Lurem-TR position and Metarhizium anisopliae..................................................................................................................45 Table 3.2: Effect of spatial separation of Lurem-TR on the persistence of conidia of Metarhizium anisopliae (% germination) in autoinoculation devices over time…..48 Table 3.3: Repeated measures ANOVA table for the response variable: Megalurothrips sjostedti attraction (A) and other insect attraction (B) (log-transformed counts) in autoinoculation devices as affected by spatial separation of Lurem-TR position and Metarhizium anisopliae…………………………………………………….……..50 Table 3.4: Effect of spatial separation of Lurem-TR and Metarhizium anisopliae on Megalurothrips sjostedti attraction (mean number of thrips per trap) on autoinoculation devices over time………………………..………………………..52 Table 3.5: Effect of spatial separation of Lurem-TR position and Metarhizium anisopliae on conidial acquisition (mean number of spores per individual thrips) on autoinoculation devices over time……………………..…………………………..54 Table 3.6: Effect of spatial separation of Lurem-TR position and Metarhizium anisopliae on the attraction of other insects (mean number per trap) on autoinoculation devices over time……..…………………………………………………………………….56 Table 4.1: General information regarding the thrips attractant compounds that were evaluated ..................................................................................................................................71 xvi Table 4.2: Effect of thrips attractants on mean percentage conidial germination of Metarhizium anisopliae and germ tube length (µm) 8 days after exposure……………...………76 Table 4.3: Effect of thrips attractants on Metarhizium anisopliae conidial germination (%) over time….…..……………………………………………………………………77 Table 4.4: Effect of thrips attractants on Metarhizium anisopliae mean conidial germ tube length (µm) over time after exposure inside dessicators..........................................79 Table 5.1: Mean number of Megalurothrips sjostedti in plots with methyl anthranilate-baited and Lurem-TR-baited inoculation devices in two planting seasons.......................101 Table 5.2: Effect of semiochemical-baited autoinoculation devices treated with Metarhizium anisopliae on conidial persistence of M. anisopliae during flowering and podding stages of the two experimental seasons..................................................................104 Table 5.3: Metarhizium anisopliae conidial acquisition by single thrips from MA-baited and Lurem-TR-baited autoinoculation devices at flowering and podding stages.........107 Table 5.4: Megalurothrips sjostedti mortality from plots with MA-baited and Lurem-TR- baited autoinoculation devices as well as the control after seven days during the podding stage of the two seasons...................…….………...……………………108 Table 5.5: Mean cowpea yield (kg/ha) of plots with MA- baited and Lurem-TR-baited autoinoculation devices as well as the control during the second season..………108 Table 6.1: Bean flower thrips density per plant following spot and cover spray applications of Metarhizium anisopliae during flowering (from day 0 to day 9) and early podding (from day 15 to day 21) during the two seasons. ...................................................125 xvii Table 6.2: Conidial acquisition by Megalurothrips sjostedti following application of Metarhizium anisopliae as spot spray and cover spray during the flowering and podding stages of cowpea during two seasons ......................................................129 Table 6.3: Cost benefit analysis in US$ following application of Metarhizium anisopliae as spot and cover spray treatments in comparison with a control and traditional farmer’s practices.....………..................................................................................131 xviii LIST OF FIGURES Figure 2.1: Geographic distribution of Bean flower thrips, Megalurothrips sjostedti in Africa (a) and in east Africa (b) (Moritz et al., 2013). ……………………..…...………..11 Figure 2. 2: Female of the bean flower thrips, Megalurothrips sjosjedti……………………….13 Figure 3.1: Experimental design for the evaluation of the effect of distance separation of Lurem-TR on Metarhizium brunneum conidial persistence in the greenhouse........36 Figure 3.2: Description of spatial separation of Lurem-TR on Metarhizium anisopliae conidial persistence in an autoinoculation device in the field. Treatments: T1 – Direct exposure of conidia to Lurem-TR; T2 – conidia separated from Lurem-TR placed inside a small container fixed just below the device; T3 - conidia separated from Lurem-TR at 10 cm above the device; T4 - conidia separated from Lurem-TR at 20 cm above the device and T5 - control, device without Lurem- TR.............................................................................................................................39 Figure 3.3: Effect of spatial separation of Lurem-TR from Metarhizium brunneum (Met52) on conidial germination. Treatments: P0, P5, P10 and P20 are respectively Petri-dishes with conidia directly exposed, 5 cm above, 10 cm above and 20 cm above Lurem- TR. Lmin and Lmax represent the minimum and the maximum effect of Lurem-TR on inhibition of spore germination when placed in closed boxes with or without Lurem-TR. Control: Petri dish atomized with conidial suspension and germination determined immediately…………………………………………………………...46 Figure 3.4: Effect of spatial separation of Lurem-TR on conidial viability of Metarhizium anisopliae in autoinoculation devices. Bars denote means ± one standard error at xix P = 0.05 (Tukey HSD). Means (±SE) of three replicates of five autoinoculation devices……………………………………………………………………………..47 Figure 3.5: Effect of spatial separation of Lurem-TR and Metarhizium anisopliae on overall attraction of Megalurothrips sjostedti. Bars denote means ± one standard error at P = 0.05 (Tukey HSD). Means (±SE) of three replicates of five autoinoculation devices..….………………………………………………………………………...51 Figure 4.1: Megalurothrips sjostedti attracted to the surface of the blue sticky card baited with attractant suspension poured in 5 ml Eppendorf tube ...…………...………………74 Figure 4.2: Relationship between Metarhizium anisopliae conidial germination and germ tube length………………………………………………………………………………80 Figure 4.3: Mean number of Megalurothrips sjostedti attracted to blue sticky cards baited with methyl anthranilate, cis-jasmone, Lurem-TR and control. Means with the same letters are not significantly different according to the Student–Newman–Keuls test (SNK)………………………………………………………………...………….....81 Figure 5.1: Autodissemination device for spatial separation of the semiochemical and entomopathogenic fungi………………………..………………………………….96 Figure 5.2: Mean number of Megalurothrips sjostedi per plant during flowering (from day 3 to day 15) and podding (from day 21 to day 30) stages in plots with methyl anthranilate-baited (ADD-MA), Lurem-TR-baited (ADD-L) and control autoinoculation devices over two seasons. *Significance using Levene’s test..........................................................................................................................102 xx Figure 5.3: Effect of semiochemical-baited autoinoculation devices on conidial persistence of Metarhizium anisopliae during flowering and podding stages of cowpea over seasons……………………………………………………………..……………105 Figure 6.1: Megalurothrips sjostedti density per plant following spot and cover spray applications of Metarhizium anisopliae during flowering (from day 0 to day 9) and early podding (from day 15 to day 21) during season I (A) and season II (B). Bars denote means ± one standard error at P =0.05 (Tukey HSD test)……………..…….126 Figure 6.2: Conidial viability of Metarhizium anisopliae following spot and cover spray applications during flowering and podding stages of cowpea during season I (A) and season II (B)............................................................................................................128 1 CHAPTER 1: GENERAL INTRODUCTION 1.1 Introduction Grain legumes are cultivated on an estimated 27 million ha in Sub-Saharan Africa (SSA) with an estimated yield of 19 million metric tons (MT). In South Asia, 40 million hectares are planted with an estimated yield of 43 MT. The export value of grain legumes expressed in terms of global exports is estimated at 0.4% in SSA and 2% in South Asia, respectively (Abate et al., 2012). These grain legumes (Fabales: Fabaceae) include common beans, Phaseolus vulgaris L., cowpea Vigna unguiculata (L.) Walp. and pigeonpea, Cajanus cajan (L.) Willsp. These crops play an important role in tropical cropping systems in SSA (Singh and Van Emden, 1978). They are major sources of plant proteins, vitamins and animal fodder (Tarawali et al., 1997; Asiwe, 2009). Cowpea is among the most consumed grain legumes in eastern Africa (Uganda, Kenya and Tanzania) (Abate et al., 2012). Insect pests, especially thrips, are regarded as mainly responsible for the low yield of grain legumes (Rachie, 1985; Jackai and Daoust, 1986; Abate and Ampofo, 1996). Thrips have a very short life cycle and overlapping of generations is frequently observed (Mac Donald et al., 1998). High infestation levels may result in complete grain yield losses if no control measures are taken (Asiwe et al., 2005). The most common thrips species (Thysanoptera: Thripidae) on cowpea in East Africa include the bean flower thrips (BFT), Megalurothrips sjostedti Trybom, Frankliniella occidentalis Pergande, Frankliniella schultzei Trybom and Hydatothrips aldolfifriderici Karny (Singh and Allen, 1979). 2 Chemical control is the main strategy for the management of thrips on grain legumes. However, most of the chemicals insecticides are toxic to humans and hazardous to the environment (Oparaeke, 2006; Nderitu et al., 2007). The effectiveness of synthetic chemicals is constrained and debatable due to the development of resistance to pesticide among thrips (Jensen, 1998; 2004; Espinosa et al., 2002), emergence of secondary pests (Graham-bryce, 1977) and the presence of toxic residues in the crop produce (Mitchell and Lykken, 1963). Hence, there is an urgent need for research on environmental-friendly alternatives. Entomopathogenic fungi (EPF) are among the alternatives being considered (Ekesi et al., 2002). A Metarhizium anisopliae (Metchnikoff) Sorokin based biopesticide has been developed by icipe and was commercialized for thrips control by Real IPM (www.realipm.com; Ekesi et al., 2009). EPF are generally applied using the conventional insecticide application approach, e.g. inundative sprays. However, this approach has a number of disadvantages including short persistence of the inoculum due to detrimental effects of solar radiation and high costs as a result of repeated applications and high volume of inoculums required (Inglis et al., 2000; Leland and Behle, 2004; Jaronski, 2010). Thrips generally respond to colour, odour, and shape (Terry, 1997; Teulon et al., 1999; Mainali and Lim, 2011). Coloured sticky traps were developed for monitoring of thrips in ornamental orchards and greenhouses (Cho et al., 1995) and blue sticky traps have been found to be the most attractive to M. sjostedti (Muvea et al., 2014). Semiochemicals (aggregation, pheromones or allelochemicals) have also been shown to attract thrips. For example, a commercial product Lurem-TR, whith the active compound, methyl isonicotinate, increase thrips catches up to 30 3 fold (Davidson et al., 2007; Teulon et al., 2010). Subsequently, the combination of semiochemicals and coloured sticky traps has become an important IPM tool for the management of thrips (Muvea et al., 2014; Niassy et al., 2012; Mfuti et al., 2016). Since EPF can be transmitted horizontally (Dimbi et al., 2013), their integration with semiochemicals provides new opportunities for use in an autodissemination /”lure and infect” device. This approach could be improved further to sustain both the thrips attraction and conidial persistence ensuring compatibility between EPF and the semiochemical. The presence of the semiochemical Lurem-TR in an autodissemination device has been reported to have a negative effect on the viability of conidia of Metarhizium anisopliae (Metchnikoff) Sorokin (Niassy et al., 2012). This finding led to the current study to investigate the compatibility between M. anisopliae and Lurem-TR in the autodissemination. 1.2. Problem statement and justification Biological control using predators and parasitoids is effective in screen houses but not in open fields. The use of EPF is therefore considered as a component for integrated thrips management under field conditions. EPF are generally applied through an inundative approach, which requires high volumes of inoculum, resulting in high costs. In addition, the short persistence of the inoculum in the field as a result of breakdown by solar radiation necessitates frequent applications, which further increases the cost. The high cost of biopesticides in general has always been considered as one of the limiting factors for their adoption by the small-scale farmers (Samuel and Graham, 2003). A 4 strategy by which insects are infected by a pathogen after being attracted to a semiochemical- baited inoculation device containing it, and disseminating the pathogen to other insects in the population after its return to the environment, could address the shortcomings described above. Alternatively, EPF could be used in combination with a semiochemical in spot spray applications, thereby reducing the quantity of inoculum needed and the resultant cost thereof. 1.3. Objectives 1.3.1 General objective To develop efficient, economical and sustainable strategies for the management of thrips on grain legumes using a “lure and infect’’ approach 1.3.2 Specific objectives The specific objectives were:  To investigate the compatibility between M. anisopliae and Lurem-TR in an autodissemination device for thrips management on grain legumes, using distance and time of separation  To identify other potential attractants that could be compatible with M. anisopliae  To evaluate the performance of the selected attractant in an autodissemination device for the management of thrips on grain legumes  To evaluate the efficacy and cost benefit of spot spray and cover spray applications of M. anisopliae through the use of the attractant Lurem-TR for the management of M. sjostedi on cowpea crops 5 1.3.3 Research Hypotheses  Distance separation of Lurem-TR from M. anisopliae in an autodissemination device will enhance their compatibility and infectivity for thrips management on grain legumes.  Thrips attractants (other than Lurem-TR) are compatible with M. anisopliae.  Alternative attractants to Lurem-TR will perform as well as Lurem-TR in an autoinoculation device for the management of thrips on grain legumes.  Spot spray and cover spray applications of M. anisopliae in combination with thrips attractants are effective and can be used for the management of M. sjostedti on cowpea. 1.4 References Abate, T., Alene, A. D., Bergvinson, D., Shiferaw, B., Silim, S., Orr, A. and Asfaw, S. (2012). Tropical grain legumes in Africa and South Asia. Knowledge and opportunities. ICRISAT-CIAT-IITA, Nairobi, Kenya. Abate, T. and Ampofo, J.K. (1996). Insect pests of beans in Africa: Their ecology and management. Annual Review of Entomology 41, 45-73. Asiwe, J.A.N. (2009). Needs assessment of cowpea production practices, constraints and utilization in South Africa. African Journal of Biotechnology 8, 5383-5388. Asiwe, J.A.N., Nokoe, S., Jackai, L.E.N. and Ewete, F.K. (2005). Does varying cowpea spacing provide better protection against cowpea pests? Crop Protection 24, 465-471. Cho, K., Eckel, C.S., Walgenbach, J.F. and Kennedy, G.G. (1995). Comparison of colored sticky traps for monitoring thrips populations (Thysanoptera: Thripidae) in staked tomato fields. Journal of Entomological Science 30, 176-190. 6 Davidson, M.M., Butler, R.C., Winkler, S. and Teulon, D.A.J. (2007). Pyridine compounds increase trap capture of Frankliniella occidentalis (Pergande) in a covered crop. New Zealand Plant Protection 60, 56-60. Dimbi, S., Maniania, N.K. and Ekesi, S. (2013). Horizontal transmission of Metarhizium anisopliae in fruit flies and effect of fungal infection on egg laying and fertility. Insects 4, 206-216. Ekesi, S., Maniania, N.K. and Lux, S.A. (2002). Mortality in three African tephritid fruit fly puparia and adults caused by the entomopathogenic fungi Metarhizium anisopliae and Beauveria bassiana. Biocontrol Science and Technology 12, 7-17. Espinosa, P.J., Bielza, P., Contreras, J. and Lacasa, A. (2002). Insecticide resistance in field populations of Frankliniella occidentalis (Pergande) in Murcia (south-east Spain). Pest Management Science 58, 967-971. Graham-bryce, I.J. (1977). Crop protection: A consideration of the effectiveness and disadvantages of current methods and of the scope for improvement. Philosophical Transaction of the Royal Society of London B: Biological Sciences 281, 163-179. Inglis, G.D., Ivie, T.J., Duke, G.M. and Goettel, M.S. (2000). Influence of rain and conidial formulation on persistence of Beauveria bassiana on potato leaves and colorado potato beetle larvae. Biological Control 18, 55-64. Jackai, L.E.N. and Daoust, R.A. (1986). Insect pests of cowpeas. Annual Review of Entomology 31, 95-119. Jaronski, S.T. (2010). Ecological factors in the inundative use of fungal entomopathogens. BioControl 55, 159-185. 7 Jensen, E.S. (2004). Insecticide resistance in the western flower thrips, Frankliniella occidentalis. Journal of Integrated Pest Management Review 5, 131-146. Jensen, S.E. (1998). Acetylcholinesterase activity associated with methiocarb resistance in a strain of western flower thrips, Frankliniella occidentallis (Pergande). Pesticide Biochemistry and Physiology 61, 191-200. Leland, J. and Behle, R.W. (2004). Formulation of the entomopathogenic fungus, Beauveria bassiana, with resistance to UV degradation for control of tarnished plant bug, Lygus lineolaris. Beltwide Cotton Conferences, San Antonio, USA. Mac Donald, J.R., Bale, S.J. and Walters, K.A.F. (1998). Effect of temperature on development of the western flower thrips Frankliniella occidentalis (Thysanoptera: Thripidae). European Journal of Entomology 95, 301-306. Mainali, B.P. and Lim, U.T. (2011). Behavioral response of western flower thrips to visual and olfactory cues. Journal of Insect Behavior 24, 436-446. Mfuti, D.K., Subramanian, S., Van Tol, R.W.H.M., Wiegers, G.L., De Kogel, W.J., Niassy, S., Du Plessis, H., Ekesi, S. and Maniania, N.K. (2016). Spatial separation of semiochemical Lurem-TR and entomopathogenic fungi to enhance their compatibility and infectivity in an autoinoculation system for thrips management. Pest Management Science, 72, 131- 139. Mitchell, L.E.and Lykken, L. (1963). Practical considerations in the degradation of pesticide chemical residues from forage crops. (In: Residue revue. Gunther, F.A., ed. Springer New York, p, 130-149). Muvea, A.M., Waiganjo, M.M., Kutima, H.L., Osiemo, Z., Nyasani, J.O. and Subramanian, S. (2014). Attraction of pest thrips (Thysanoptera: Thripidae) infesting french beans to 8 coloured sticky traps with Lurem-TR and its utility for monitoring thrips populations. International Journal of Tropical Insect Science 34, 197-206. Nderitu, J.H., Wambua, E.M., Olubayo, F., Kasina, J.M. and Waturu, C.N. (2007). Management of thrips (Thysanoptera: Thripidae) infestation on french beans (Phaseolus vulgaris L.) in Kenya by combination of insecticides and varietal resistance. Journal of Entomology 4, 469-473. Niassy, S., Maniania, N.K., Subramanian, S., Gitonga, L.M. and Ekesi, S. (2012). Performance of a semiochemical-baited autoinoculation device treated with Metarhizium anisopliae for control of Frankliniella occidentalis on french bean in field cages. Entomologia Experimentalis et Applicata 142, 97-103. Oparaeke, A.M. (2006). The sensitivity of Flower Bud Thrips, Megalurothrips sjostedti Trybom (Thysanoptera: Thripidae), on cowpea to three concentrations and spraying schedules of Piper guineense Schum and Thonn extracts. Plant Protection Science 42, 106-111. Rachie, K.O. (1985). Introduction. (In: Cowpea research, production and utilization. Singh, R.H., Rachie, K.O., ed. John Wiley & Sons, U.K, p. XXi-XXViii). Samuel, G.M. and Graham, A.M. (2003). Recent developments in sprayers for application of biopesticides. An overview. Biosystems Engineering 84, 119-125. Singh, S.R. and Allen, D.J. (1979). Cowpea pests and diseases, Manual series No. 2. IITA, Ibadan, Nigeria. Singh, S.R. and Van Emden, H.F. (1978). Insect pests of grain legumes. Annual Review of Entomology 24, 255-278. Tarawali, S., Singh, B. B., Peters, M. and Blade, S.F. (1997). Cowpea haulms as fodder. ( In : Singh, B.B., Mohan Raj, D.R., Dashiell, K.E. and Jackai, L.E.N., ed. Advances in 9 cowpea research. Co-publication of International Institute of Tropical Agriculture (IITA), and Japan International Research Center for Agricultural Sciences (JIRCAS), Ibadan, Nigeria. p. 313-325). Terry, L.I. (1997). Host selection, communication and reproductive behaviour. (In : Lewis, T., ed. Thrips as crop pests. CAB International, Wallingford, UK, p. 65-118). Teulon, D.A.J., Davidson, M.M., Nielsen, M., Perry, N., Van Tol, R. and de Kogel, W. (2010). The lure of scent: allelochemicals for thrips pest management. Journal of Insect Science 10, 49-50. Teulon, D.A.J., Hollister, B., Butler, R.C. and Cameron, E.A. (1999). Colour and odour responses of flying western flower thrips: wind tunnel and greenhouse experiments. Entomologia Experimentalis et Applicata 93, 9-19. 10 CHAPTER 2: LITERATURE REVIEW 2.1 Thrips taxonomy and identification Thrips belonging to order Thysanoptera, are present worldwide and only 5000 of an estimated 8000 extant species have been described (Mound and Houston, 1987). The Thysanoptera are divided into two suborders: Terebrantia and Tubilifera (Lewis, 1997; Mound, 2009). Most pest thrips species found on grain legumes belong to the suborder: Terebrantia. Among them, bean flower thrips (BFT), Megalurothrips sjostedti Trybom (Thysanoptera: Thripidae) is the most common thrips species found on cowpea (Vigna unguiculata L. Walp) in tropical Africa (Moritz et al., 2013). 2.2 Geographical Distribution Thrips are widespread throughout the world and are found in various habitats including forests, grasslands, and areas of low vegetation and deserts as well as on most cultivated crops. The different thrips species can be classified as phytophagous, carnivorous species, gall-makers or inquilines (Lewis, 1973). Species that feed on a wide range of plants and are crop pests are mostly in the family Thripidae (Moritz et al., 2004). Some flower thrips reproduce in flowers and feed on the cells of the flower tissue, on pollen grains and on small developing fruits (Mortiz et al., 2004). Many of the flower-dwelling species are partly predatory on small insects whilst other species primarily feed on leaves (Lewis, 1973; Mortiz et al., 2004). The Bean flower thrips, M. sjostedti occurs throughout tropical Africa (Figure 2.1) (Singh and Van Emden, 1978; Moritz et al., 2013). Adults can prevail in the dry savannah throughout the 11 year (Bottenberg et al., 1997), indicating a much higher degree of adaptability to unfavourable conditions, which might be a consequence of their capability to feed and reproduce on more diverse types of plants (Tamò et al., 1993). Legumes (Fabales: Fabaceae) are the main host plants of M. sjostedti and include cowpea [V. unguiculata], pigeon pea [Cajanus cajan (L.) Willsp], common beans/French beans [Phaseolus vulgaris L.] (Tamo et al., 1993; Moritz et al., 2013). They also attack other plant species which are considered as minor hosts such as groundnut [Arachis hypogaea] (Tamo et al., 1993) and wild host plants (Tamo et al., 1997). (a) ( (b) Figure 2.1: Geographic distribution of bean flower thrips, Megalurothrips sjostedti in Africa (a) and in east Africa (b) (Moritz et al., 2013). 12 2.3 Biology Development rate of thrips is highly dependent upon environmental conditions and nutrient quality of their food sources (Mound, 1997). All described genera of thrips are haplodiploid organisms capable of parthenogenesis, with some favoring arrhenotoky (unfertilized eggs develop into males) and others, thelytoky (unfertilized eggs develop into females) (Lewis, 1997; Kumm and Moritz, 2008). Thysanoptera species are hemimetabolous insects with an incomplete metamorphosis (Mound, 1997; 2005). Females of M. sjostedti undergo a pre-oviposition period which lasts from a day to a week during which their eggs mature, and before they mate. Although mated females of M. sjostedti laid eggs that produce both sexes, a very high percentage of their offspring is females (Lewis, 1997; Kumm and Moritz, 2008). The life cycle has six distinct stages. Eggs are very tiny (0.25 mm long and 0.1 mm wide). They are white when freshly laid and turn pale yellow toward maturation. Eggs are usually laid singly inside the plant tissue, and are therefore not visible (Lewis, 1997). They hatch within 3 to 20 days, depending on temperature. The first and second instars are very small (0.5 to 1.2 mm). They are wingless and usually lighter in colour than the adults. The larval stage lasts for 8 to more than 20 days in total, followed by non-feeding prepupal and pupal stages. The pupal stages are usually completed in the soil at the base of the plant. After 3 to 6 days, the adult thrips emerge (Mound and Kibby, 1998). Although most adult thrips possess long fringed wings, wingless adults also occur (Lewis, 1973; 1997; Mound, 1997). 13 2.4 Economic importance of thrips Yield losses caused by M. sjostedti have been estimated to be between 20 and 100% in various parts of Africa (Singh and Allen, 1980). In Kenya for instance, 94% yield loss has been reported on cowpea (Ampong-Nyarko et al., 1994). Thrips damage mainly the floral parts (flowers, buds and pods) of plants. Infested flower buds become brown and eventually abort leaving behind dark red scars (Singh et al., 1997). Damaged flowers are characterized by distortion, malformation and discoloration of floral parts (Singh and Van Emden, 1978). Figure 2. 2: Female of the bean flower thrips Megalurothrips sjostedti. 2.5 Control strategies for thrips 2.5.1 Chemical control Chemical insecticide application is the most widely used thrips control method. Diverse insecticides such as chlorpyrifos-methyl, methiocarb, methamidophos, acrinathrin, endosulfan, deltamethrin and formetanate are often used (Singh and Rachie, 1985; Jackai and Adalla, 1997). However, development of pest resistance to insecticides has resulted in higher dosages and more frequent insecticide applications with more environmental hazards and negative effects on 14 human, environment and non-target insect species. Rotation of insecticides with different modes of action has been suggested to reduce pest resistance (Alghali, 1992) but not all the farmers can afford it. 2.5.2 Intercropping Cultural control practices such as intercropping have been reported to reduce M. sjostedti infestations on crops (Kyamanywa and Ampofo, 1988; Kyamanywa and Tukahirwa, 1988; Kyamanywa et al., 1993; Ampong-Nyarko et al., 1994). For example, yield loss caused by M. sjostedti was reduced from 94% to 51% in cowpea/sorghum intercrop which also received chemical treatment (Ampong-Nyarko et al., 1994). Ekesi et al. (1999) also reported reductions in M. sjostedi numbers by 72 and 96% in cowpea monocrop and cowpea intercrop treated with M. anisopliae, respectively. 2.5.3 Thrips monitoring and trapping Early detection of thrips infestation could be crucial for their successful control. Visual inspection by tapping plants on a tray or checking flowers at regular time intervals are often used (Pearsall and Myers, 2000). Thrips monitoring should be done at least once a week, and more often when an infestation is detected. Coloured sticky cards are currently the best monitoring tool for thrips populations (Plimmer et al., 1982; Cho et al., 1995; Koschier et al., 2000; Muvea et al., 2014). Blue and yellow are the colours mostly recommended (Blumthal et al., 2005; Muvea et al., 2014). It is recommended that sticky traps should be placed above the crop canopy so that the bottoms of the traps are just above the crop, at a rate of one or two traps per 1,000 square feet (Greer and Diver, 2000). Regular monitoring is crucial for effective control. 15 2.5.4 Semiochemicals Thrips respond to olfactory cues (pheromones, semiochemiclas or allelochemicals) (De Kogel and Koschier, 2003; Kirk and Terry, 2003; Hamilton et al., 2005; Muvea et al., 2014). Subsequently, semiochemical-based products such as Lurem-TR and Thripline have been developed for use in thrips monitoring and management (Sampson and Kirk, 2013; Teulon et al., 2014; Broughton et al., 2015). These semiochemicals can be integrated with other control strategies to improve thrips management in horticulture (Suckling et al., 2012; Sampson and Kirk, 2013). Lurem-TR is a commercial semiochemical whose active ingredient is methyl- isonicotinate. It was previously reported to be effective in monitoring Frankliniella occidentalis (Pergande) (Thysanoptera: Thripidae) (Davidson et al., 2007) and several other pest thrips (Nielsen et al., 2010). More recently, it has been also reported to be effective against M. sjostedti populations (Muvea et al., 2014; Mfuti et al., 2016). 2.5.5 Biological control 2.5.5.1 Predators Larvae of thrips are easy prey for a wide range of general arthropod predators but those more specific to thrips include members of the Aeolothripidae, the anthocorid genera Orius and Montandoniola, the Cecidomyiid genus Thripsobremia and the Sphecidae genus Microstigmus (Mills, 1991). Some of them are commercially available and are currently used as biological control agents in a variety of crops (Driesche et al., 1998; Van Lenteren and Loomans, 1998; Loomans, 2003). In Africa, Fritzsche and Tamo (2000) reported Orius albidipennis Reuter (Heteroptera: Anthocoridae) to be a natural enemy of M. sjostedti on cowpea and other host plants. 16 2.5.5.2 Parasitoids Parasitoid species identified for M. sjostedti control include Ceranisus menes Walker (Hymenoptera: Eulophidae), (Diop, 1999), C. femoratus Gahan (Hymenoptera: Eulophidae) (Tamo et al., 1997; 2012), Megaphragma priesneri Kryger (Hymenoptera: Trichogrammatidae) and M. mymaripenne Timberlake (Hymenoptera: Trichogrammatidae) (Tamo et al., 1993; Loomans, 2003; Noyes, 2014). 2.5.5.3 Entomopathogenic fungi Entomopathogenic fungi (EPF) are among the entomopathogens being considered for biological control of thrips (Butt and Brownbridge, 1997; Ekesi and Maniania, 2002). EPF are generally applied through inundative sprays, which require high quantities of inocula, thereby encreasing its cost (Jaronski, 2010). The persistence of conidia applied on foliage is influenced by several environmental parameters such as UV light, rain, temperature (Inglis et al., 2000; Jaronski, 2010), which necessitates an improvement in application technique. There are many published reports on successful control of thrips by EPF (Ekesi et al., 1998, 1999; Maniania et al., 2003). A number of fungus-based products are now registered or marketed for the control of thrips worldwide (Faria and Wraight, 2007; Lacey et al., 2015). In Kenya, an isolate of M. anisoplaie ICIPE 69 is commercialized for the control of thrips by RealIPM (www.realipm.com; Ekesi et al., 2009). 17 2.5.5.4 Current strategies for delivery of entomopathogenic fungi in the field Entomopathogenic fungi are generally applied using inundative sprays similar to the conventional insecticide application approach (Jaronski, 2010). However, this technique has a number of shortcomings including the use of high volumes of inoculum, short persistence in the field due to breakdown by solar radiation which leads to repeated applications that are too expensive (Fargues et al., 1996; Inglis et al., 2000; Jaronski, 2010). Responses of insects to visual and olfactory cues are exploited for their management. For example, semiochemicals are used to lure large numbers of insects into a trap, inoculated with EPF as with termites (Alves et al., 2002). This strategy has led to the concept of autodissemination/autoinoculation. It consists of a semiochemical-baited inoculation device containing the pathogen. The insects are attracted to the device. On entering, they are infected with the pathogen and on return to the environment they disseminate the pathogen among the insects in the population (Vega et al., 2007). This strategy has been developed against a number of insects including fruit flies, Ceratitis spp. (Diptera: Tephritidae) (Dimbi et al., 2003), tsetse flies, Glossina spp. (Diptera: Glossinidae) (Maniania, 1998, 2002), pea leafminer, Liriomyza huidobrensis (Diptera: Agromyzidae) (Migiro et al., 2010) and recently against F. occidentallis (Thysanoptera: Thripidae) (Niassy et al., 2012). The cost of this technique is low in comparison to cover-spray applications. The integration of pheromones and kairomones in thrips management (Teulon et al., 2014; Broughton et al., 2015) therefore offers new perspectives for application of EPF for the control of thrips (Niassy et al., 2012; Mfuti et al., 2016). 18 2.6 References Alghali, A.M. (1992). Insecticide application schedules to reduce grain yield losses caused by insects of cowpea in Nigeria. International Journal of Tropical Insect Science13, 725- 730. Ampong-Nyarko, K., Reddy, K.V.S., Nyang, R.A. and Saxena, K.N. (1994). Reduction of insect pest attack on sorghum and cowpea by intercropping. Entomologia Experimentalis et Applicata 70, 179-184. Alves, S.B., Pereira, R.M., Lopes, R.B. and Tamai, M.A. (2002). Use of entomopathogenic fungi in Latin America. (In: Upadhyay, R.K., ed. Control of insect pests. Kluwer Academic/Plenum Publishers, p. 193-211). Blumthal, M.R., Cloyd, R.A., Spomer, L.A. and Warnock, D.F. (2005). Flower color preferences of western flower thrips. HortTechnology 15, 846-853. Bottenberg, H., Tamò, M., Arodokoun, D., Jackai, L.E.N., Singh, B.B. and Youm, O. (1997). Population dynamics and migration of cowpea pests in northern Nigeria: implications for integrated pest management. (In :Singh, B.B., Mohan Raj, D.R., Dashiell, K.E. and Jackai, L.E.N., ed. Advances in cowpea research. Co-publication of International Institute of Tropical Agriculture and Japan International Center for Agricultural Sciences. IITA, Ibadan, Nigeria, p. 271-284). Broughton, S., Cousins, D.A. and Rahman, T. (2015). Evaluation of semiochemicals for their potential application in mass trapping of Frankliniella occidentalis (Pergande) in roses. Crop Protection 67, 130-135. Butt, T.M. and Brownbridge, M. (1997). Fungal pathogens of thrips. (In: Lewis, T., ed. Thrips as crop pests. CAB International, Wallingford, UK, p. 399-433). 19 Cho, K., Eckel, C.S., Walgenbach, J.F. and Kennedy, G.G. (1995). Comparison of colored sticky traps for monitoring thrips populations (Thysanoptera: Thripidae) in staked tomato fields. Journal of Entomological Science 30, 176-190. Cook, S.M., Khan, Z.R. and Pickett, J.A. (2007). The use of Push-Pull strategies in integrated pest management. Annual Review of Entomology 52, 375-400. Davidson, M.M., Butler, R.C., Winkler, S. and Teulon, D.A.J. (2007). Pyridine compounds increase trap capture of Frankliniella occidentalis (Pergande) in a covered crop. New Zealand Plant Protection 60, 56-60. De Kogel, W.J. and Koschier, E.H. (2003). Thrips responses to plant odors. (In: Marullo, R. and Mound, L., ed. 7th International Symposium on Thysanoptera: Thrips, Plants, Tospoviruses. The millenial review. Reggio Calabria, Italy, p. 189-190). Dimbi, S., Maniania, N.K., Lux, S.A., Ekesi, S. and Mueke, J.M. (2003). Pathogenicity of Metarhizium anisopliae (Metsch.) Sorokin and Beauveria bassiana (Balsamo) Vuillemin to three adult fruit fly species: Ceratitis capitata (Wiedemann), C. rosa var. fasciventris Karsch and C. cosyra (Walker) (Diptera: Tephritidae). Mycopathologia 156, 375-382. Diop, K. (1999)."The biology of Ceranisus menes Walker (Hymenoptera: Eulophidae), a parasitoid of the bean flower thrips Megalurothrips sjostedti Trybom (Thysanoptera: Thripidae): a comparison between African and Asian populations." PhD dissertation, University of Ghana. Driesche, R.G.V., Heinz, K.M., van Lenteren, J.C., Loomans, A., Wick, R., Smith, T., Lopes, P., Sanderson, J.P., Daughtrey, M. and Browbridge, M. (1998). Western flower thrips in greenhouses: A review of its biological control and other methods. (In: 1998 Amherst, Massachusetts Mass Extension Floral Facts, University of Massachusetts, p. 32). 20 Ekesi, S. and Maniania, N.K. (2002). Metarhizium anisopliae: An effective biological control agent for the management of thrips in horti and floriculture in Africa. (In: Upadhyay, R.K., ed. Advances in Microbial Control of Insects Pests, New York. Kluwer Academic/Plenum publishers). Ekesi, S., Maniania, N.K. and Onu, I. (1999). Effects of the temperature and photoperiod on the development and ovoposition of the legume flower thrips, Megalurothrips sjostedti. Entomologia Experimentalis et Applicata 93, 149-155. Ekesi, S., Maniania, N.K., Onu, I. and Lohr, B. (1998). Pathogenicity of entomopathogenic fungi (Hyphomycetes) to the legumes flower thrips, Megalurothrips sjostedti (Thysanoptera: Thripidae). Journal of Applied Entomology 122, 629-634. El-Sayed, A.M., Suckling, D.M., Wearing, C. H. and Byers, J.A. (2006). Potential of mass trapping for long-term pest management and eradication of invasive species. Journal of Economic Entomology 99, 1550-1564. Fargues, J., Goettel, M.S., Smits, N., Ouedraogo, A., Vidal, C., Lacey, L.A., Lomer, C.J., Rougier, M. (1996). Variability in susceptibility to simulated sunlight of conidia among isolates of entomopathogenic Hyphomycetes. Mycopathologia 135, 171-181. Faria, M.R. and Wraight, S.P. (2007). Mycoinsecticides and Mycoacaricides: A comprehensive list with worldwide coverage and international classification of formulation types. Biological Control 43, 237-256. Fritzsche, M. and Tamo, M. (2000). Influence of thrips prey species on the life-history and behaviour of Orius albidipennis Reuter (Heterptera). Entomologia Experimentalis et Applicata 96, 111-118. 21 Greer, L. and Diver, S. (2000). Greenhouse IPM: Sustainable thrips control. Appropriate Technology Transfer for Rural Areas (ATTRA), NCAT Agriculture Specialists 148, 18. 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Assessing the attractiveness of volatile plant compounds to western flower thrips Frankliniella occidentalis. Journal of Chemical Ecology 26, 2643-2655. Kumm, S. and Moritz, G. (2008). First detection of Wolbachia in arrhenotokous populations of thrips species (Thysanoptera: Thripidae and Phlaeothripidae) and its role in reproduction. Environmental Entomology 37, 1422 - 1428. 22 Kuslitzky, W. (2003). New variant: Annotated list of hymenopterous parasitoids of thrips in Israel. 20th Conference of the Entomological Society of Israel. Phytoparasitica 31, 11- 12. Kyamanywa, S., Baliddawa, C.W. and Ampofo, K.J.O. (1993). Effect of maize plants on colonization of cowpea plants by bean flower thrips, Megalurothrips sjostedti. Entomologia Experimentalis et Applicata 69, 61-68. Kyamanywa, S. and Ampofo, J.K.O. (1988). Effect of cowpea/maize mixed cropping on the incident light at the cowpea canopy and flower thrips (Thysanoptera: Thripidae) population density. Crop Protection 7, 186-189. 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Pests, diseases, resistance and protection in cowpea. (In: Advances in Legume Science. Summerfield, R.A. and Bunting, H.H., ed. Royal Botanical Garden, Kew, Ministry of Agriculture, Fisheries and Food, p. 419-433). 26 Singh, S.R., and Van Emden, H.F. (1978). Insect pests of grain legumes. Annual Review of Entomology 24, 255-278. Singh, S.R. and Rachie, K.O. (1985). Introduction. (In: Cowpea Research, Production and Utilization. Singh S.R. and Rachie K.O., ed. John Wiley and Sons, New York). Suckling, D.M., Walker, J.T.S., Clare, G.K., Boyd Wilson, K.S.H., Hall, C., El-Sayed, A.M. and Stevens, P.S. (2012). Development and commercialisation of pheromone products in New Zealand. New Zealand Plant Protection 65, 267-273. Tamò, M., Srinivasan, R., Dannon, E., Agboton, C., Datinon, B., Dabire, C., Baoua, I., Ba, M. N., Haruna, B. and Pittendrigh, B. R. (2012). Biological control: A major component for the long‐term cowpea pest management strategy. 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Assessment of key factors responsible for the pest status of the bean flower thrips Megalurothrips sjostedti (Trybom) (Thysanoptera, Thripidae). Bulletin of Entomological Research 83, 251-258. Tagashira, E. and Hirose, Y. (2001). Development and reproduction of Ceranisus menes (Hymenoptera: Eulophidae), a larval parasitoid of thrips: effects of two host species, Frankliniella intonsa and Thrips palmi (Thysanoptera: Thripidae). Applied Entomology and Zoology 36, 237-241. Teulon, D.A.J., Castañé, C., Nielsen, M.-C., El-Sayed, A.M., Davidson, M.M., Gardner-Gee, R., Poulton, J., Kean, A.M., Hall, C., Butler, R.C., Sansom, C.E., Suckling, D.M. and Perry, N.B. (2014). Evaluation of new volatile compounds as lures for western flower thrips and onion thrips in New Zealand and Spain. New Zealand Plant Protection 67, 175-183. Teulon, D.A.J., Davidson, M.M., Nielsen, M., Perry, N., Van Tol, R. and De Kogel, W. (2010). The lure of scent: allelochemicals for thrips pest management. Journal of Insect Science 10, 49-50. Van Lenteren, J.C. and Loomans, A.J.M. (1998). Is there a natural enemy good enough for biological control of thrips? Proceedings of the 1998 Brighton conference on Pests and Diseases 2, 401-408. Van Tol, R.W.H.M., James, D.E., De Kogel, W.J. and Teulon, D.A.J. (2007). Plant odours with potential for a push-pull strategy to control the onion thrips, Thrips tabaci. Entomologia Experimentalis et Applicata 122, 69-76. Vega, F.E., Dowd, P.F., Lacey, L.A., Pell, J.K., Jackson, D.M. and Klein, M.G. (2007). Dissemination of beneficial microbial agents by insects. (In: Lacey, L.A. and Kaya, H.K., 28 ed. Field Manual of Techniques in Invertebrate Pathology 2nd edition Springer, Dordrecht, The Netherlands, p. 127-146). 29 CHAPTER 3: SPATIAL SEPARATION OF SEMIOCHEMICAL LUREM-TR AND ENTOMOPATHOGENIC FUNGI TO ENHANCE THEIR COMPATIBILITY AND INFECTIVITY IN AN AUTOINOCULATION SYSTEM FOR THRIPS MANAGEMENT Abstract The effect of spatial separation of the semiochemical Lurem-TR, which has been found to inhibit conidia of entomopathogenic fungi when put together, on the persistence of conidia of Metarhizium brunneum and M. anisopliae was evaluated in the greenhouse and field in order to develop an autodissemination strategy for the management of bean flower thrips, Megalurothrips sjostedti on cowpea crop. Influence of spatial separation of the semiochemical on thrips attraction and conidial acquisition by thrips from the autoinoculation device was also investigated in the field. Persistence of conidia of M. brunneum and M. anisopliae increased with distance of separation of Lurem-TR. Direct exposure of fungus without separation from Lurem- TR recorded the lowest conidial germination as compared to the other treatments. Attraction of thrips to the device also varied significantly according to distance between device and semiochemical, with a higher number of thrips attracted when Lurem-TR was placed in a container below the device and at 10 cm distance. There was no significant difference in conidia acquisition between spatial separation treatments of conidia and Lurem-TR. Attraction of other insect pests to the device did not significantly vary between treatments. Positive correlations were found between conidial acquisition and thrips attraction. This study suggests that spatial separation of fungal conidia from Lurem-TR in an autoinoculation device could provide a low- cost strategy for effective management of thrips in grain legume cropping systems. 30 Published as : MFUTI, K.D, SUBRAMANIAN, S., VAN TOL, R.W.H.M., WIEGERS, G.L., DE KOGEL, W.J., NIASSY, S., DU PLESSIS, H., EKESI, S. and MANIANIA, N.K. (2016) Spatial separation of semiochemical Lurem-TR and entomopathogenic fungi to enhance their compatibility and infectivity in an autoinoculation system for thrips management. Pest Management Science 72(1): 131-139. The greenhouse experiments included in the chapter were undertaken by co-authors of the manuscript Dr. R.W.H.M. van Tol, Dr. G.L. Wiegers and Dr. W.J. de Kogel in the Netherlands. 3.1 Introduction Grain legumes are among the key economical crops widely grown in eastern and western Africa as important sources of food and animal fodder (Abate et al., 2012; Tarawali et al., 1997). In Kenya, the annual bean production is estimated at 577,674 MT (USAID, 2013). However, the production of grain legumes is compromised by a complex of insect pests such as the legume pod borer, Maruca vitrata Fabricius (Lepidoptera: Pyralidae), bean stem maggots, Ophiomyia spp. (Diptera: Agromyzidae), aphids (Hemiptera: Aphididae) and thrips (Thysanoptera: Thripidae) (Abate and Ampofo, 1996). Among the thrips, the bean flower thrips (BFT), Megalurothrips sjostedti (Trybom) (Thysanoptera: Thripidae), is considered as the most important pest attacking the reproductive structures of grain legumes (Tamò et al., 1993). Damage by M. sjostedi includes early flower blemishes, abscission and necrosis with yield losses ranging between 20 to 100% (Singh and Allen, 1980). 31 Thrips are difficult to control owing to their cryptic flower dwelling behaviour and their minute size (Lewis, 1997). Chemical control is the most widely adopted management strategy by farmers who often resort to using obsolete or banned chemical pesticides with detrimental consequences to human, environmental and animal health (Nderitu et al., 2007). The introduction of stringent regulations by European importing countries such as the Maximum Residue Limit (MRL) has led to several crop rejections and economical losses. In addition, thrips have developed resistance to most of the chemical insecticides and hence the need to explore other control strategies including biological control (Brødsgaard, 1994; Espinosa et al., 2002; Jensen, 1998; 2004). Entomopathogenic fungi (EPF) are among the most promising alternatives to synthetic chemical pesticides (Butt and Brownbridge, 1997; Ekesi et al., 2001; Niassy et al., 2012b). Fungal-based biopesticides for control of thrips are commercially available and include Metarhizium anisopliae (Metschnikoff) Sorokin ICIPE 69 marketed as Campaign® by the RealIPM, Kenya. The most common application technique of EPF is through inundative sprays (Hajek and St- Leger, 1994). However, EPF conidia applied on foliage have short persistence owing to environmental factors such as UV light, temperature and rain (Daoust and Pereira, 1986; Hong et al., 1999; Inglis et al., 2000; Jaronski, 2010). For instance, Ekesi et al. (2001) reported persistence of M. anisopliae conidia for 3-4 days on cowpea leaves. Such short persistence in the field requires frequent applications of EPF, resulting in higher inoculum requirement and high costs. Another application technique referred to as autodissemination or autoinoculation consisting of attracting insects to an autoinoculator where they are infected with a pathogen before returning to the environment to disseminate the pathogen to conspecifics is also being 32 considered (Vega et al., 2007). This approach has already been tested against Frankliniella occidentalis Pergande on French bean (Niassy et al., 2012a) and is based on combined use of visual cues (blue color), the semiochemical attractant Lurem-TR and the entomopathogenic fungus M. anisopliae. However, Lurem-TR was found to negatively affect conidial germination and infectivity of M. anisopliae in field cages (Niassy et al., 2012a). Introduction of Lurem-TR in a dessicator containing a culture of M. anisopliae resulted in complete inhibition of its germination after 48 hrs, confirming field results (S. Niassy, pers. observation). In order to improve the performance of autodissemination device for thrips management, we explored the effect of distance separation of Lurem-TR from fungal conidia on the persistence of M. brunneum in greenhouse and M. anisopliae under field conditions. We also evaluated the influence on thrips attraction and conidial acquisition in various distance separation treatments under field conditions. 3.2 Materials and methods Study site The study was conducted in the greenhouse at Plant Research International, Wageningen, The Netherlands (51.986: 5.663, 13 m above sea level) (T = 20 °C, 16:8 L:D photoperiod), and in the field of Kamiti, Kiambu County, Kenya (1.191S: 36.883E, 1640 m above sea level) and at icipe, Nairobi (1.221S, 36.896E; 1616 m above sea level). In the greenhouse, the experiments intended to assess the effect of Lurem-TR on the persistence of M. brunneum while experiments in the field assessed the effect of Lurem-TR on the persistence of M. anisopliae strain ICIPE 69, attraction of thrips and other insects, and conidial acquisition by thrips. Experiments were carried 33 out during the dry season of May-August 2013. Average temperatures and relative humidity of 20.8 ˚C and 74.2%, respectively, were recorded in the experimental field. Entomopathogenic fungi Conidia of M. brunneum were obtained from the commercial product BIO1020 (strain Met52) (Bayer CropScience, The Netherlands). They were cultured on Sabouraud dextrose agar medium (SDA) at 25-27 ˚C, pH = 5.6±0.2 (Cooke et al., 2002). Conidia were harvested from the plate and suspended in 0.01% Triton X-100 and conidial concentration determined using a haemocytometer (Fuchs-Rosenthal 0.2 mm). A spore suspension of approximately 109 conidia ml-1 was prepared and stored for 2 days at 5 oC until use in the experiment. Metarhizium anisopliae isolate ICIPE 69 is commercially available and marketed as Campaign® by the RealIPM, Kenya. Conidia of M. anisopliae were mass-produced on long rice substrate in Milner bags (60 cm long by 35 cm wide). Rice was autoclaved for 1 h at 121 oC and inoculated with a 3- day-old culture of blastospores (Jenkins and Goettel, 1997). The rice containing fungal spores was then allowed to dry for five days at room temperature. Conidia were harvested by sifting the substrate through a 295-µm mesh sieve and stored for 2 days at 5 oC until use. Conidial viability was determined before any experiment by spread-plating 0.1ml of the suspension (3 x 106 conidia ml-1) on Sabouraud Dextrose Agar (SDA) plates. Sterile microscope cover slips were placed on each plate. Plates were then incubated at 24-28 oC, 12:12 L:D photoperiod and examined after 16-20 hours. Percentage germination was determined by counting the number of germ tubes formed among 100 random conidia for each plate at 400 x under a light microscope (Goettel and Inglis, 1997). Conidial germination of approximately 90% was considered acceptable. 34 Semiochemical Lurem-TR, a commercial semiochemical whose active ingredient is methyl-isonicotinate, previously reported to be effective in monitoring thrips populations was used in this study (Davidson et al., 2007). It was obtained from Pherobank (Wageningen, The Netherlands). 3.2.1 Effect of spatial separation of Lurem-TR on the persistence of conidia of Metarhizium brunneum in the greenhouse Four 9 cm Petri dishes without cover were placed at 0, 5, 10 and 20 cm, corresponding to treatments P0, P5, P10 and P20, respectively, on a rack with platforms connected with a stick in such a way that all platforms/Petri-dishes were vertically under each other (Figure 3.1). Lurem- TR was placed above the top Petri dish (P0). Petri dishes contained water agar (1.5% w/w) on which eight cover slips of 10 mm diameter (0.79 cm2) previously atomized with a spore suspension of M. brunneum were placed. Atomization was done by spraying 4 ml conidial suspension (approximately equivalent to 600 l/ha) of M. brunneum on eight glass cover slips placed on Petri dishes without water agar at a pressure of 7.5 bar using a Potter Precision Laboratory Spray Tower (Burkard Manufacturing Co Ltd., Rickmansworth, United Kingdom). Petri dishes were allowed to dry for 20-30 min, after which cover slips were transferred to the Petri-dishes containing water agar and then placed in the rack. The treated Petri dishes were exposed to Lurem-TR for 24 h. As a control, a Petri dish was atomized with conidial suspension as described above and allowed to dry, and conidial germination determined immediately. All treatments were replicated two times and repeated four times. 35 To determine the maximum effect of Lurem-TR on inhibition of conidial germination, in addition to the four treatments described above, Petri dishes were prepared as detailed above and placed in closed boxes (diameter 10 cm, height 10 cm) with or without Lurem-TR. After 24 h the spore germination was determined. The persistence of conidia was determined after a period of 24 h for all the treatments including the control. Conidial viability was determined according to an adapted method of Faria et al. (2010). Each cover slip with conidia was removed from the Petri dish, placed in a 10-ml Greiner tube containing 1 ml of 0.01% Triton X-100 water solution and vortexed for 20s to dislodge conidia. From each Greiner tube, three samples of each 10 μl were pipetted separately on one glass slide covered with a thin layer of SDA and incubated in a closed container on humidified filter paper in the dark for 24 h at 25 oC. Percentage germination was determined by pipetting one droplet of lactophenol on each sample after 24 h, covering it with a cover slip and counting the number of germinating and non-germinating conidia (minimum count was 200 spores per droplet). 36 Figure 3.1: Experimental design for the evaluation of the effect of distance separation of Lurem- TR on Metarhizium brunneum conidial persistence in the greenhouse. 37 Field experiment with autoinoculation device The effect of spatial separation of Lurem-TR on the persistence of M. anisopliae, attraction of M. sjostedi and other insects, and conidial acquisition by thrips was evaluated in field experiments. Cowpea, Vigna unguiculata L. Walp variety Ken-Kunde1, was planted in 10 m2 plots with inter- and intra- row spacing of 10 and 45 cm, respectively. No fertilizers, organic matter or insecticides were applied during the experimental period. The autoinoculation device used in the present study and procedure for the inoculation of device is as described by Niassy et al. (2012a). Briefly, a Lynfield trap (11 cm diameter x 10 cm height) was perforated with six entry ⁄ exit holes (2 x 3 cm) near the top and bottom of the bottle at alternate positions. Velvet (8 x 8.5 cm) and blue netting (3.5 x 11 cm) were wrapped around a smaller inner cylindrical bottle (5.2 cm diameter x 6 cm high) that was then hung inside the trap. The semiochemical dispenser used to lure thrips was placed in different positions (see Figure 3.2). Approximately, 2–3 g of dry conidia was spread evenly on the velvet cloth of the autoinoculation device. Blue netting was then wrapped around the velvet cloth containing spores and tightened with two office pins. The device was then hung at crop canopy level (35 cm). The following treatments were used in the field with the autoinoculation device: T1 – Direct exposure of fungal conidia to Lurem-TR; T2 – Conidia separated from Lurem-TR placed inside a small container fixed just below the device, hereafter also referred to 0 cm; T3 - Conidia separated from Lurem-TR at 10 cm above the device; T4 - Conidia separated from Lurem-TR at 20 cm above the device and T5 - Control (device without Lurem-TR) (Fig. 3.2). Treatments were laid out in a complete randomized block design with three blocks as replicates. The blocks and 38 treatments were separated by a distance of at least 15 m to avoid interferences between treatments and within blocks. Each of the five treatments was deployed in a single plot so there were five plots and these were repeated three times. For conidial viability, five treatments replicated four times were used, giving a total of 20 experimental units. The experiment on thrips conidial acquisition and attraction, five treatments were replicated three times (15 experimental plots in total). Experiments were conducted during peak flowering stage of the crop which corresponds to the period of peak infestation of the crop by thrips necessitating control measures. The crop was planted 14 June 2013, and experiments run from July to August 2013. The flowering stage occurred from 24 July 2013 to 7 August 2013 while the podding stage started from 7August 2013 up to harvest. 39 Figure 3.2: Description of spatial separation of Lurem-TR on Metarhizium anisopliae conidial persistence in an autoinoculation device in the field. Treatments: T1 – Direct exposure of conidia to Lurem-TR; T2 – conidia separated from Lurem-TR placed inside a small container fixed just below the device; T3 - conidia separated from Lurem-TR at10 cm above the device; T4 - conidia separated from Lurem-TR at 20 cm above the device and T5 - control, device without Lurem-TR. 40 3.2.2 Effect of spatial separation of Lurem-TR on Metarhizium anisopliae conidia persistence in the field The persistence of conidia of M. anisopliae was evaluated for a period of two weeks after the onset of the experiment. At three–day intervals, samples of conidia were collected from the autoinoculation devices from the five treatments using a moist cotton bud. The end of the cotton bud was cut, suspended in 10-ml 0.05% (wt ⁄vol) Triton X-100 and vortexed for 1 min to dislodge conidia. A sample of 100 µl was spread-plated on SDA and incubated for 16 h at 25 ± 2 oC and L12:D12 photoperiod. Germination of conidia was determined as described above. 3.2.3 Attraction of Megalurothrips sjostedti and other pests A blue sticky card (5 cm × 10 cm) was fixed to the side of the autoinoculaion device with or without Lurem-TR to determine the number of insects, including M. sjostedti visiting the device. The sticky cards were replaced every three days. Kerosene was used to dissolve the glue on the sticky cards and insects were removed with a fine brush. Thrips specimens were then cleared, mounted on slides and identified as described in the Lucid Key Pest thrips of the world and Pest thrips of east Africa (Moritz et al., 2004; Moritz et al., 2013). The number of thrips and other insect pests such as leaf miners and bean stem maggot were recorded. 3.2.4 Conidial acquisition by Megalurothrips sjostedti To assess the number of conidia acquired by a single thrips visiting the autoinoculation device, 5 to 10 cowpea plants from a distance of 2 m around the autoinoculation device were randomly sampled using a whole plant tapping technique (Pearsall and Myers, 2000). The latter consists of 41 tapping plants on a white barber tray (25 x 45 cm) where the tray is held underneath the selected plant, while the plant is tapped gently by hand (5 taps). In each treatment, five cowpea plants were sampled around the autoinoculation device (1-2 m radius) and 20 insects were collected in separate glass containers (10-ml) using an aspirator. Containers were labelled and stored in the fridge for immobilization. Insects were transferred individually into 2-ml cryogenic tubes containing 1 ml of sterile 0.05% Triton X-100. The tube was vortexed for 2–3 min to dislodge conidia from the insect and the concentration of conidia was determined using a Neubauer haemocytometer. 3.2.5 Statistical analysis In the greenhouse experiment, differences in germination rate of conidia of M. brunneum between treatments were assessed by linear logistic regression analysis of the observed counts of germinated spores over the total number of spores examined for the replicate. The data Y were treated as pseudo-binomial data, taking the variance to be proportional to binomial variance, i.e. var (Y) = σ2np (1-p). Here p (045% after 12-15 days post-exposure. In field cage studies, Niassy et al. (2012) reported that conidial viability was not affected in autoinoculation device without a semiochemical, 7 days post-treatment, but in the Lurem-TR- 110 baited autoinoculation device, conidial viability decreased from 80 to 6% at 2 and 7 days post- inoculation, respectively. The difference between the two studies is that conidia were directly exposed to Lurem-TR (Niassy et al., 2012) while they were spatially separated in the present study thereby enhancing compatibility (see Mfuti et al., 2016). Maniania (2002) reported that conidia of M. anisopliae could retain up to 60% of their viability after a 31-day exposure in a contamination device. The mean number of conidia acquired by a single thrips in both semiochemical-baited devices varied between 2.0 x 103 and 10.0 x 103 conidia and was lower than what was reported by Niassy et al. (2012) for F. occidentalis (Pergande) (Thysanoptera: Thripidae). The study conducted by Niassy et al. (2012) was, however, conducted in experimental cages where multiple infections are possible compared to the present study which was conducted under field conditions. These results have shown that MA is as effective as Lurem-TR in terms of conidial acquisition. Mortality of M. sjostedi in the two semiochemical-baited treatments ranged between 40-46%. However, in the presence of the semiochemical, Dimbi et al. (2003) reported mortality between 70 and 93% of fruit flies Ceratitis rosa (Karsch) and C. fasciventris (Bezzi) (Diptera : Tephritidae) after being attracted to M. anisopliae-treated autoinoculators baited with brewer’s yeast in a field cage. In field cage studies, Niassy et al. (2012) reported thrips mortality ranging between 41.7 - 59.3% in autoinoculation device with or without semiochemicals. Similarly to conidial acquisition, differences in mortality among the different studies can be attributed to differences in experimental conditions (field cage vs open field) and difference in target insects. 111 Cowpea yield was higher in plots with the MA autoinoculation baited device compared to the Lurem-TR baited autoinoculation device and the control device. Bud formation and flowering phase of the crops are considered as critical crop growth stages for M. sjostedi management. Alghali (1991) reported that bean flower thrips can cause a yield reduction of 44% on cowpea at bud formation and flowering stages. Ezueh, (1981) reported that yield loss of 100% can occur on cowpea production if no control measures are taken. The higher yield in treated plots observed in this study can be linked to the higher mortality of M. sjostedi in treated plots as compared to the control. Among the treatments the increase in yield was more perceptible in the treatment MA- baited inoculation device as compared to the inoculation device baited with Lurem-TR. This could be attributed to the better conidial acquisition in MA treatments, which could have resulted in better secondary spread of the entomopathogen. The underlying factors for greater efficiency of MA as compared to Lurem-TR need further scrutiny. 5.5 Conclusion The current study demonstrated the field efficacy of methyl anthranilate that is compatible with M. anisopliae and effective in attracting and infecting M. sjostedi. Hence, it could be used as alternative attractant to commercial semiochemical Lurem-TR in an autoinoculation device for M. sjostedi management instead of cover spray applications of biopesticides. 112 5.6 References Abate, T. and Ampofo, J.K. (1996). Insect pests of beans in Africa: Their ecology and management. Annual Review of Entomology 41, 45-73. Adipala, E., Omongo, C.A., Sabiti, A., Obuo, J.E, Edema, R., Bua, B., Atyang, A., Nsubuga, E.N. and Ogenga-latigo, M.W. (1999). Pests and diseases on cowpea in Uganda: Experiences from a diagnostic survey. African Crop Science Journal 7, 465-478. Alghali, A.M. (1991). Integrated pest management strategy for cowpea production under residual soil moisture in the Bida area of northern Nigeria. Tropical Pest Management 37, 224- 227. Daoust, R.A. and Pereira, R.M. (1986). Stability of entomopathogenic fungi, Beauveria bassiana and Metarhizium anisopliae on beetle-attracting tubers and cowpea foliage in Brazil. Environmental Entomology 15, 1237-1243. Dimbi, S., Maniania, N.K., Lux, S.A., Ekesi, S. and Mueke, J.M. (2003). Pathogenicity of Metarhizium anisopliae (Metsch.) Sorokin and Beauveria bassiana (Balsamo) Vuillemin to three adult fruit fly species: Ceratitis capitata (Weidemann), C. rosa var. fasciventris Karsch and C. cosyra (Walker) (Diptera: Tephritidae). Mycopathologia 156, 375-382. Ekesi S., Maniania, N.K., Ampong-Nyarko, K. and Akpa, A.D. (2001). Importance of timing of application of the entomopathogenic fungus, Metarhizium anisopliae for the control of legume flower thrips, Megalurothrips sjostedti and its persistence in cowpea. Archive Phytopathology and Plant Protection 33, 431-445. 113 Ekesi, S., Maniania, N.K. and Lux, S.A. (2002). Mortality in three African tephritid fruit fly puparia and adults caused by the entomopathogenic fungi Metarhizium anisopliae and Beauveria bassiana. Biocontrol Science and Technology 12, 7-17. Espinosa, P.J., Bielza, P., Contreras, J. and Lacasa, A. (2002). Insecticide resistance in field populations of Frankliniella occidentalis (Pergande) in Murcia (south-east Spain). Pest Management Science 58, 967-971. Ezueh, M.I. (1981). Nature and significance of pre-flowering damage by thrips to cowpea. Entomologia Experimentalis et Applicata 29, 305-312. Fargues, J., Goettel, M.S., Smits, N., Ouedraogo, A., Vidal, C., Lacey, L.A., Lomer, C.J. and Rougier, M. (1996). Variability in susceptibility to simulated sunlight of conidia among isolates of entomopathogenic Hyphomycetes. Mycopathologia 135, 171-181. Fritsche, M.E. and Tamo, M. (2000). Influence of thrips species on the life history and behaviour of Orius albidipennis. Entomologia Experimentalis et Applicata 96, 111-118. Hamilton, J.G.C., Hall, D.R. and Kirk, W.D.J. (2005). Identification of a male produced aggregation pheromone in the western flower thrips, Frankliniella occidentalis. Journal of Chemical Ecology 31, 1369-1379. Inglis, G.D., Ivie, T.J., Duke, G.M. and Goettel. M.S. (2000). Influence of rain and conidial formulation on persistence of Beauveria bassiana on potato leaves and colorado potato beetle larvae. Biological Control 18, 55-64. Jackai, L.E.N. and Daoust, R.A. (1986). Insect pests of cowpeas. Annual Review of Entomology 31, 95-119. Jaronski, S.T. (2010). Ecological factors in the inundative use of fungal entomopathogens. BioControl 55, 159-185. 114 Jensen, S.E. (2004). Insecticide resistance in the western flower thrips, Frankliniella occidentalis. Journal of Integrated Pest Management Reviews 5, 131-146. Kimani, P.M., Nyende, A.B. and Silim, S. (1994). Development of early maturing fusarium wilt resistant pigeonpea cultivars. African Crop Science Journal 2, 35-41. Krueger, S., Subramanian, S., Niassy, S. and Moritz, G.B. (2015). Sternal gland structures in males of bean flower thrips, Megalurothrips sjostedti, Poinsettia thrips, Echinothrips americanus in comparison with those of western flower thrips, Frankliniella occidentalis (Thysanoptera:Thripidae). Arthropod Structure and Development 44, 455-467. Leland, J. and Behle, R.W. (2004). Formulation of the entomopathogenic fungus, Beauveria bassiana, with resistance to UV degradation for control of tarnished plant bug, Lygus lineolaris. Beltwide Cotton Conferences, San Antonio, USA. Maniania, N.K. (2002). A low-cost contamination device for infecting adult tsetse flies, Glossina spp., with the entomopathogenic fungus Metarhizium anisopliae in the field. Biocontrol Science and Technology 12, 59-66. Mergeai, G., Kimani, P., Mwang’ombe, A., Olubayo, F., Smith, C., Audi, P., Baudoin, J-P. and Le Roi, A. (2001). Survey of pigeonpea production systems, utilization and marketing in semi-arid lands of Kenya. Biotechnology, Agronomy, Society and Environment 5, 145- 153. Mfuti, D.K., Subramanian, S., van Tol, R.W.H.M., Wiegers, G.L., De Kogel, W. J., Niassy, S., Du Plessis, H., Ekesi, S. and Maniania, N.K. (2016). Spatial separation of semiochemical Lurem-TR and entomopathogenic fungi to enhance their compatibility and infectivity in an autoinoculation system for thrips management. Pest Management Science 72, 131- 139. 115 Moritz, G., Brandt, S., Triapitsyn, S. and Subramanian, S. (2013). Identification and information tools for pest thrips in East Africa. QAAFI Biological Information Technology (QBIT), The University of Queensland, Brisbane, Australia. ISBN 978-1-74272-0687-8: http://thripsnet.zoologie.uni-halle.de/key-server-neu/. Muvea, A.M., Waiganjo, M.M., Kutima, H.L., Osiemo, Z., Nyasani, J.O. and Subramanian, S. (2014). Attraction of pest thrips (Thysanoptera: Thripidae) infesting French beans to coloured sticky traps with Lurem-TR and its utility for monitoring thrips populations. International Journal of Tropical Insect Science 34, 197-206. Niassy, S., Ekesi, S., Maniania, N.K., Moritz, G.B., de Kogel, W. J. and Subramanian, S. (2015). Aggregation ecology in bean flower thrips, Megalurothrips sjostedti Trybom (Thysanoptera: Thripidae). Entomologia Experimentalis et applicata DOI: 10.1111/eea.12383. Niassy, S., Maniania, N.K., Subramanian, S., Gitonga, L.M. and Ekesi, S. (2012). Performance of a semiochemical-baited autoinoculation device treated with Metarhizium anisopliae for control of Frankliniella occidentalis on French bean in field cages. Entomologia Experimentalis et Applicata 142, 97-103. Ogah, E.O. (2011). Assessing the impact of varietal resistance and planting dates on the incidence of African yam bean flower thrips (Megalurothrips sjostedti, Hochst. Ex. A. Rich) in Nigeria. Asian Journal of Plant Sciences 10, 370-375. Oparaeke, A.M. (2006 ). The sensitivity of flower bud thrips, Megalurothrips sjostedti Trybom (Thysanoptera: Thripidae), on Cowpea to three concentrations and spraying schedules of Piper guineense Schum and Thonn extracts. Plant Protection Science 42, 106-111. 116 Pearsall, I.A. and Myers, J.H. (2000). Population dynamics of western flower thrips (Thysanoptera: Thripidae) in British Columbia. Journal of Economic Entomology 93, 264-275. Rachie, K.O. (1985). Introduction. (In: Cowpea research, production and utilization (Singh, R.H. and Rachie, K.O., ed.) John Wiley & Sons, U.K, p. XXi-XXViii). R Development Core Team (2014). A Language and environment for statistical computing. R Foundation for Statistical Computing ed., Vienna, Austria. Saidi, M., Itulya, F.M., Aguyoh, J.N. and Ngouajio, M. (2010). Effects of cowpea leaf harvesting initiation time and frequency on tissue nitrogen content and productivity of a dual- purpose cowpea–maize intercrop. Hortscience 45, 369-375. Smits, N., Fargues, J., Rougier, M., Goujet, R. and Itier, B. (1996). Effect of temperature and solar radiation interactions on the survival of quiescent conidia of the entomopathogenic fungus Paecilomyces fumusoroseus (Wize) Brown and Smith. Mycopathologia 135, 163-170. Vega, F.E., Dowd, P.F., Lacey, L.A., Pell, J.K., Jackson, D.M. and Klein, M.G. (2007). Dissemination of beneficial microbial agents by insects. (In: Field Manual of Techniques in Invertebrate Pathology 2nd edition, Lacey, L.A. and. Kaya, H.K., ed. Springer, Dordrecht, The Netherlands, p. 127-146). 117 CHAPTER 6: IMPROVING APPLICATION OF FUNGUS-BASED BIOPESTICIDE IN COMBINATION WITH SEMIOCHEMICAL FOR THE MANAGEMENT OF BEAN FLOWER THRIPS ON COWPEA Abstract The efficacy of spot spray and cover spray applications of Metarhizium anisopliae (Metsch.) Sorok. in combination with the thrips attractant Lurem-TR (methyl-isonicotinate), was compared in field experiments for the management of the bean flower thrips (BFT), Megalurothrips sjostedti (Trybom) (Thysanoptera: Thripidae) on a cowpea crop over two seasons. Treatments were applied five days after the placement of Lurem-TR in the field. During the first season, M. sjostedti densities were lower in spot spray (10.1±4.3 thrips) and cover spray (11.5±4.8 thrips) treatments than in the control treatment (41.8±15.2 thrips). In the second season, the lowest M. sjostedti density of 32.5±6.0 thrips were recorded in the spot spray treatment, followed by cover spray with 40.9±7.0 thrips recorded. The highest M. sjostedti density of 67.4±10.3 thrips was recorded in the control treatment. Fungal viability and thrips conidial acquisition did not differ between the two application methods. Both application strategies resulted in a yield increase of 34.1 and 46.2% compared to the control with the spot and cover spray treatments, respectively. The cost benefit analysis indicated more profits with the spot spray than cover spray application due to the reduction in labour and the quantity of inoculum used. Spot spray application of biopesticides could therefore be a more viable option for small-scale farmers for the management of M. sjostedti on cowpea. 118 6.1 Introduction Cowpea, Vigna unguiculata L. Walp. (Fabales: Fabaceae), is an important food and cash crop in different parts of the tropics (Quin et al., 1997). It occupies a vital place in human nutrition as sources of protein, vitamins and minerals. In Kenya, cowpea is among the most consumed grain legumes, but the recorded yield is low and the demand can therefore not be satisfied (Mergeai et al., 2001). Annual production of cowpea in Kenya declined from about 83,000 MT in 2007 to about 48,000 MT in 2008, despite an increase in area planted from around 130,000 in 2007 to about 148,000 hectares over the same period (Belmain et al., 2013; Kiprotich et al., 2015). Insect pests are the main factor responsible for the low grain legume production (Abate et al., 2012; Ajeigbe et al., 2012). The bean flower thrips (BFT), Megalurothrips sjostedti (Trybom) (Thysanoptera: Thripidae), is considered as major pest attacking the reproductive structures of cowpeas during plant development ( Ezueh, 1981). Megalurothrips sjostedti can cause yield losses ranging from 20 to 100% (Singh and Allen, 1980). The control of M. sjostedti relies heavily on the use of synthetic chemical insecticides (Jackai and Daoust, 1986; Abate and Ampofo, 1996), which is associated with numerous problems such as safety of workers health risks to consumers and livestock, and environmental contamination (Nderitu et al., 2007). Furthermore, intense applications of chemical insecticides can result in residue problems on harvested beans (Löhr, 1996). The use of entomopathogenic fungi (EPF) in horticulture is gaining momentum in Africa over the last few years (Ekesi et al., 2001; Ekesi and Maniania, 2007). Biopesticides are generally applied through inundative sprays, which requires high quantities of inoculum. In addition, their short persistence in the field due to solar radiations 119 means that repeated applications are often needed, resulting in high costs. Therefore, there is the need to find an alternative application strategy to overcome these limitations. The use of semiochemicals to lure large number of insects in a trap that can then be inoculated with EPF has been reported in the case of termites (Alves et al., 2002). This technique is attractive as it has low cost in comparison to the broadcast application. The integration of pheromones and kairomones in thrips management (Teulon et al., 2014; Broughton et al., 2015) can offer new perspectives for application of EPF for the control of thrips (Niassy et al., 2012; Mfuti et al., 2016). It has been reported that the presence of Lurem-TR attracts high numbers of M. sjostedti adults in baited traps (Muvea et al., 2014). It is therefore hypothesized that spot spray application of EPF in combination with thrips attractant could reduce the quantity of inoculum, thereby the cost of the thrips management. The objective of this study was therefore to evaluate the efficacy and cost benefit analysis of spot spray and cover spray applications of the entomopathogenic fungus M. anisopliae through the use of the attractant Lurem-TR for the management of M. sjostedti on cowpea crop. 6.2 Materials and methods Study site A field experiment was conducted from June to December 2014 at Mbita Thomas Odhiambo Campus, situated in the eastern shores of Lake Victoria (0 ̊ 26’ 06.19” S, 34 ̊ 12’ 53.13” E; 1,137 m above sea level) in western Kenya. The vegetation type around the campus is mainly savannah grassland with mixed combretum and acacia trees to the north and papyrus along the shores of the lake. The experiment was conducted in two seasons. In the first season, cowpea was planted on 6 June 2014 and the experiment was conducted from July to August 2014. It coincided with 120 the dry season with low infestation of M. sjostedti. However, the season was characterized by high cowpea aphid infestation. In the second season, cowpea was planted on 10 August 2014 and the experiment was conducted from October to December 2014. This coincided with the short rainy season with high M. sjostedti infestation. The fungus Metarhizium anisopliae isolate ICIPE 69 was obtained from Real IPM Ltd. in Kenya. It is registered in Kenya and other African countries and commercialized under the trade name of Campaign®. Emulsifiable formulation of the fungus (1 x 109 CFU ml-1) was applied at the recommended dose of 200 ml ha-1, corresponding to 2.0 x 1014 conidia ha-1, using a CP15 knapsack sprayer (Cooper Pergler, Sussex, UK) with an output of 350 litres ha-1. Spores were checked for viability before application and conidial germination over 85% was considered acceptable. Semiochemical Lurem-TR was used in the present study as previously described in chapter 3, section 3.2. Experimental design Cowpea variety Ken-Kunde 1 was planted with an intra-row spacing of 50 cm and inter-row spacing of 20 cm. Irrigation was provided once a week and weeding was done regularly. No fertilizers, organic matter or synthetic chemical insecticides were applied during the experimental periods. The experiment was conducted during the flowering and early podding stages which coincided with higher M. sjostedti populations (Ezueh, 1981; Nyasani et al., 2013). 121 The field was divided into four blocks, each with three equal size experimental plots (7 x 7 m2). Treatments were arranged in these plots in a complete randomized block design. Blocks were separated by 5 m while within blocks, plots were separated by 2 m to avoid interference of treatments. The treatments were: T1: Blue card+Lurem-TR (Control); T2: Blue card+Lurem-TR+M. anisopliae applied in spot spray; T3: Blue card+Lurem-TR+M. anisopliae applied in cover spray. For spot spray, the fungus was applied on a 9-m2 area around the Lurem-TR attractant placed at the centre of each plot within the block. In the cover spray treatment, the fungus was applied as a full cover spray in the plot i.e. 48 m2. Preliminary data on the attraction of M. sjostedti population to Lurem-TR-baited sticky cards indicated that the optimal time to attract ~50% of M. sjostedti was between 3 and 5 days after deployment of the baited cards in fields (Niassy et al. unpublished). The fungus was applied five days after deployment of Lurem-TR in the plots at the beginning of flowering (which ran from day 0-9 of the experiment) and then again at the beginning of the podding stage (which ran from day 9-21 of the experiment). No fungus was applied in the control plot. For assessing conidial persistence and acquisition, data from treatments with M. anisopliae applied in cover and spot sprays only were analysed. 6.2.1 Effect of fungal application strategy on Megalurothrips sjostedti density The number of M. sjostedti per plant was recorded every three days for a period of 9 days during both the flowering and podding stages. Random sampling of five cowpea plants was done in each treatment plot using the whole plant tapping technique. Plants were tapped gently with the 122 hand, five times, on a white barber tray (25 x 45 cm) held underneath the selected plant (Pearsall and Myers, 2000). The tray was cleaned after each sampling. 6.2.2 Effect of fungal application strategy on Metarhizium anisopliae conidial persistence The persistence of conidia of M. anisopliae was evaluated from samples taken on the day of fungal application (day 0), day 1 and 4 after fungal applications. Samples were collected randomly by cutting three cowpea leaves from plants in plots that received fungus treatments. Leaves were cut in small pieces and suspended in 10-ml 0.05% (wt ⁄vol) Triton X-100 and vortexed for 1 min to dislodge the conidia. A sample of 100 µl was spread-plated on SDA plates containing chloramphenicol (500 µg/ml) to inhibit growth of bacterial contaminants (Inglis et al., 2012). It was incubated for 16 h at 25 ± 2 ˚C in total darkness. Germination of conidia was determined as described above (see chapter 3, section 3.2). The sampling for cover spray treatments was done at random over the entire plot, while for the spot spray; it was done in the 9m2 area around the traps. 6.2.3 Effect of fungal applications strategy on conidial acquisition Twenty adult M. sjostedti were randomly collected from 5 cowpea plants in fungus-treated plots using an aspirator to assess the number of conidia acquired by single insect. Megalorothrips sjostedti were transferred to glass containers which were labelled and stored in the fridge to immobilize the insects. They were then transferred individually into 2-ml cryogenic tubes containing 1 ml of sterile 0.05% Triton X-100. The tubes were vortexed for 2–3 min to dislodge conidia from the thrips and the concentration of conidia was determined using a Neubauer haemocytometer. 123 6.2.4 Cowpea yield Due to high infestation of cowpea aphid during the first season, the yield could not be assessed. Therefore, cowpea yield was only obtained during the second season. Pods from each plot were harvested 90 days after planting, sundried on a large cement surface for three weeks and weighed with a Mettler PM 15 balance. The yield obtained from the grains was calculated in Kg/ha. 6.2.5 Cost benefit analysis The net benefit was calculated using the partial budgeting procedure (El-Deep Soha, 2014) which assesses the costs and benefits associated with a specific change in an individual enterprise within the business operation. The procedure focuses specifically on the implications of the intended change in a business operation by comparing the benefits and costs resulting from implementing the alternative with respect to the current practice. Partial budget, like an enterprise budget, is based on a unit (a one crop farm). Only variable input costs are used in a partial budget. The net benefit is the difference between the gross farm gate benefit and total variable input costs. The cost benefit analysis was calculated taking into account the traditional farmer’s practices where no blue sticky card and semiochemical attractants are used. Grain yields of crops grown under low-input conditions are low and unstable. Grain yields of between 350 and 540 kg ha-1 were reported in Kenya (Ekesi et al., 1998, 1999; Katungi et al., 2009). The grain yield of 350 kg ha-1 was therefore used for the cost benefit analysis calculation. 124 6.2.6 Statistical analysis All data were tested for normality and homogeneity of variance using Shapiro–Wilk and Bartelett tests (Shapiro and Wilk, 1965; Snedecor and Cochran, 1989). Due to the over dispersion of M. sjostedti between treatments, data were subjected to the negative binomial of generalize linear model. The means were compared with Tukey HSD test at 5% significance level. Repeated measures ANOVAs were used to analyze M. anisopliae conidial persistence and conidial acquisition. Yield data were subjected to one way ANOVA and the means compared with Tukey HSD test at 5% significance level. All data analyses were performed using R (Version 3.1.3, 2015) statistical software (R Development core Team 2014). 6.3 Results 6.3.1 Effect of fungal application strategy on Megalurothrips sjostedti density The density of M. sjostedti was higher in the control than in the fungus-treated plots in both seasons (season I: F = 59.5; df = 2,61; P<0.001; season II: F = 85.5; df = 2,61; P<0.001). There was no significant difference in M. sjostedti density between spot spray and cover spray application plots in season I (Table 6.1). There was, however, a significant difference in M. sjostedti density between spot spray and cover spray application plots in season II, with the spot spray treatment having the lowest number of thrips (Table 6.1). A significant difference was detected over time during the two experimental seasons (season I: F = 4.2; df = 6,61; P<0.001; season II: F = 26.4; df = 6,61; P<0.001)(Figure 6.1). Megalurothrips sjostedti density was higher in the control than in cover and spot spray treatments at all sampling days in season I (Figure 6.1A) and season II (Figure 6.1B). 125 Table 6.1: Megalurothrips sjostedi density per plant following spot and cover spray applications of Metarhizium anisopliae during flowering (from day 0 to day 9) and early podding (from day 15 to day 21) during the two seasons. Treatments Mean M. sjostedti density/plant ± SE Season I Season II Control 41.8±15.2a 67.4±10.3a Cover spray 11.5±4.8b 40.9±7.0b Spot spray 10.1±4.3b 32.5±6.0c F = 59.5; df = 2, 61; P < 0.001 F = 85.5; df = 2, 61; P < 0.001 Means (±SE) followed by the same letters within the column are not significantly different according to Tukey’s HSD 126 Figure 6.1: Megalurothrips sjostedti density per plant following spot and cover spray applications of Metarhizium anisopliae during flowering (from day 0 to day 9) and early podding (from day 15 to day 21) during season I (A) and season II (B). Bars denote means ± one standard error at P =0.05 (Tukey HSD test). 0 10 20 30 40 50 60 70 80 90 100 0 3 6 9 15 18 21 N u m b er o f M . sj o st ed ti /p la n t Day after treatment Control Cover spray Spot spray 0 10 20 30 40 50 60 70 80 90 100 0 3 6 9 15 18 21 N u m b er o f M . sj o st ed ti /p la n t Day after treatment Control Cover spray Spot spray B A 127 6.3.2 Effect of fungal application strategy on Metarhizium anisopliae conidial persistence and Megalurothrips sjostedti conidial acquisition There was no significant difference in conidial persistence of M. anisopliae between the two fungal application strategies in both seasons. Season I (Flowering: F = 0.2; df = 1, 11; P = 0.7; Podding: F= 0.0; df = 1, 11; P = 1.0), season II (Flowering: F = 0.1; df = 1, 11; P = 0.8; Podding: F= 0.2; df = 1, 11; P = 0.7). There was also no significant treatment x time interaction during flowering (F = 0.2; df = 1, 30; P = 0.6) as well as during podding (F= 0.0; df = 1, 30; P = 0.9) of season I.Conidial viability did, however, decrease significantly over time during both flowering (F = 40.7; df = 1, 30; P < 0.0001) and podding (F= 515.1; df = 1, 30; P < 0.0001) (Figure 6.2 A). For example, conidial germination dropped from 93.4% at day 0 to 58.6 and 47.9% after one day in spot spray and cover spray treatments, respectively, and to 32.0% after 4 days in both treatments during the flowering stage of season 1 (Figure 6.2 A). During the podding stage, conidial germination reduced from 87.0% on day 0 to between 42.2% and 45.6% on day 1 and to 13.7% on day 4 post-treatment in both spray applications (Figure 6.2 A). There was a significant treatment x time interaction during flowering of season II (F = 6.3; df = 1, 30; P < 0.01), but not during the podding stage of season II (F= 0.0; df = 1, 30; P = 1.0). Conidial viability of the fungus, regardless of the method of application decreased significantly over time (Flowering: F = 16.5; df = (1, 30); P < 0.0001; Podding: F=93.0; df = (1, 30); P < 0.0001) (Figure 6.2 B). A drastic decrease in conidial germination was observed one day after application of the fungus during the flowering stage of season II with a decreased from 85.0% at day 0 to 26.2% in both treatments. Germination further decreased 4 days after treatment, to 17.0% in the cover spray and 9.4% in spot spray treatments respectively (Figure 6.2 B). During the podding stage, conidial viability decreased from 84.6% on day 0 to 47.0 and 45.0% one day after treatment in spot spray 128 and cover spray treatments, respectively, and to 6.1%, 4 days post-treatment for both application methods (Figure 6.2 B). A: Season I B: Season II 0 10 20 30 40 50 60 70 80 90 100 0 1 4 % G er m in a ti o n ± S E Days after treatment in flowering stage Cover spray Spot spray 0 10 20 30 40 50 60 70 80 90 100 0 1 4 Days after treatment in podding stage 0 10 20 30 40 50 60 70 80 90 100 0 1 4 % G er m in a ti o n ± S E Days after treatment in flowering stage Cover spray Spot spray 0 10 20 30 40 50 60 70 80 90 100 0 1 4 Days after treatment in podding stage Figure 6.2: Conidial viability of Metarhizium anisopliae following spot spray and cover spray applications during the flowering and podding stages of cowpea during season I (A) and season II. (B). 129 6.3.3 Effect of fungal application strategy on conidial acquisition No significant differences in the number of conidia acquired by M. sjostedti were observed between the two application strategies during the flowering stage of both seasons and the podding stage of season 2 (Table 6.2). There was, however, a significant difference in the number of conidia acquired by M. sjostedti between the cover and spot spray application treatments during the podding stage of season 1 (Table 6.2). Table 6.2: Conidial acquisition by Megalurothrips sjostedti following application of Metarhizium anisopliae as spot and cover sprays during flowering and podding of cowpea over two seasons. Season Crop stage Days Cover spray (x 104conidia) Spot spray (x 104 conidia) Season I Flowering 1 1.4±0.8 0.5±0.3 4 0.4±0.2 0.5±0.3 F = 1.2; df = 1, 11; P = 0.3 Podding 1 0.3±0.1 1.1±0.8 4 0.3±0.1 0.4±0.2 F = 5.6; df = 1, 11; P < 0.05 Season II Flowering 1 0.3±0.2 0.5±0.2 4 1.3±0.8 0.5±0.3 F = 0.4; df = 1, 11; P = 0.5 Podding 1 0.8±0.1 1.0±0.4 4 0.9±0.7 0.1±0.1 F = 0.5; df = 1, 11; P = 0.6 130 6.3.4 Cowpea yield The yield was significantly high (F =5.8; df = 2, 6; P < 0.04) in both cover (1430.7±114.2 kg) and spot spray (1312.1±87.7 kg) treatments than in the control treatment plots (976.8±105.2 kg). There was, however, no significant difference in yield of the fungus treatment plots. 6.3.5 Cost benefit analysis in US$ following application of M. anisopliae as spot and cover sprays compared to current farmer’s practices Variables that were considered in the cost benefit analysis are listed in Table 6.3. The total input cost for crops treated with cover and spot sprays is estimated at US$ 674 and US$ 611.4 respectively. The input cost for the control plots is estimated at US$ 597 and US$ 155 for traditional farmer’s practices (Table 6.3). The gross income is estimated at US$ 3,148 and US$ 2,887 for crops receiving cover and spot spray, respectively, US$ 2,153 for crops produced in the control plots and US$ 770 for crops produced by using the traditional farmer’s practices (Table 6.3). The net benefit is therefore higher in the cover and spot spray treatments than in the control plots (Table 6.3). In comparison with the traditional farmer’s practices and with the control, the cost benefit analysis showed that the rate of return was higher for the spot spray treatment than the cover spray (Table 6.3). 131 Table 6.3: Cost benefit analysis in US$ following application of Metarhizium anisopliae as spot and cover spray treatments in comparison with a control treatment and traditional farmer’s practices. Variables Cover spray Spot spray Traditional farmer’s practices Control Area for calculation 1 ha 1 ha 1 ha 1ha Fungal application area 1 ha 0.1875ha Gross income Average cowpea yield (kg/ha) 1,430.7 1,312.1 350.0 978.6 Unit selling price of dry cowpea seeds (US$/kg) 2.2 2.2 2.2 2.2 Revenue (US$/ha) 3,147.5 2,886.6 770.0 2,153.0 Variable input costs Technological cost (US$) Attractant (US$/ha) 383.6 383.6 383.6 Blue card (US$/ha) 58.4 58.4 58.4 Cost of oil formulation of fungal spores (US$/ha) x 2 sprays 50.0 9.4 0.0 Total technological cost (US$/ha) 492.0 451.4 0.0 442.0 Seed cost (US$/ha) 73.0 73.0 73.0 73.0 Labor cost(US$/ha) 109.0 87.0 82.0 82.0 Total variable input costs 674.0 611.4 155.0 597.0 Net benefit (US$) 2,473.5 2,275.2 615 1,556.0 Change in net benefit relative to the traditional farmer’s practices# and to the control* #1,858.5 *917.5 #1,660.2 (719.2) Change in total variable input cost relative to the traditional farmer’s practices# and to the control* #519.0 *77.0 #456.4 *14.4 Rate of return to the traditional farmer’s practices# and to the control* #3.58 *11.91 #3.64 *49.94 132 6.4 Discussion This study demonstrated that a combination of Lurem-TR with M. anisopliae applied as cover and spot sprays significantly reduced M. sjostedti densities on cowpea during both seasons. With the exception of the second season during which a greater reduction in M. sjostedti numbers occurred in spot spray than in cover spray treatments. Both the fungus treatments reduced the M. sjostedti numbers significantly during the first season. The use of spot sprays can therefore overcome the shortcomings of cover spray applications, e.g. high volume of inoculum needed to achieve effective mortality of the target pest (Samuel and Graham, 2003; Jaronski, 2010). This is evidenced by the low input costs (fungus application, total technological and labour costs) recorded in the spot spray treatment compared to that of the cover spray. In comparison with the traditional farmer’s practices, the cost benefit analysis showed a rate of return of 358% for the cover and 364% for the spot spray treatments. In other words, an investment of US$1 in fungus application on cowpea recoups the US$1 and gives an additional US$ 3.58 and US$ 3.64 benefit for the cover ans spot sprays, respectively. However, when compared with the control used in this study, US$ 1 recoups the US$ 1 and results in an additional US$ 11.91 and US$ 49.94 respectively, if M. anisopliae is applied as cover and spot spray treatments. Despite the fact that the total variable inputs were less in the traditional farmer’s practices, the rate of return was much lower due to biotic pressure on cowpea plants, resulting in lower productivity. It is expected that this rate of return will increase over time since some of the materials such as blue cards can be reused, resulting in lower input costs for the farmer. Fungal conidia acquired by individual thrips were similar in both application methods and are lower than reported in literature. For instance, Mfuti et al. (2016) reported conidial acquisition in 133 the range of 3.0 and 10.0 x 104 by single adult M. sjostedti depending on distance of separation of Lurem-TR and M. anisopliae in autoinoculation devices. Similar results were reported by Ugine et al. (2005) with B. bassiana against adult F. occidentalis. In another study, Niassy et al. (2012) observed that the overall mean number of conidia of M. anisopliae acquired per single F. occidentalis adults were 5.0 ± 0.6 x 104 in field cages with Lurem-TR-baited device while 2.2 ± 0.4 x 104 conidia in cages without semiochemical 7 days post-inoculation. A correlation between conidial acquisition and mortality was reported by Migiro et al. (2010) and Niassy et al. (2012). However, Ugine et al. (2005) noted that the rate of conidial acquisition on the bodies of F. occidentalis decreased significantly as the density of conidia on the leaf disk surface increased. The relatively lower number of conidia acquired by M. sjostedti in the present study compared to other studies could also be explained by the flower dwelling behaviour of M. sjostedti considering that experiments were conducted during flowering and the early podding stage. Conidial viability decreased drastically after application in both cover and spot spray treatments. This could be explained by the fact that conidia in both treatments were subjected to the same abiotic factors such as solar radiation which is known to affect their persistence (Jaronski, 2010). Results from this study are in agreement with that of Fargues et al. (1996) who reported that survival of conidia of entomopathogenic ascomycetes species decreased with increased exposure to sunlight. For example, exposure for 2 h or more was found to be detrimental to all isolates tested. Under simulated sunlight in the laboratory, the same authors observed that persistence of conidia of Beauveria bassiana isolate LRC 26 were reduced by 64% after only 1 h exposure and none survived 4 hours of continuous exposure. Under natural conditions, conidial populations of the same isolate reduced much slower, 99% and 75-90% after 4 days on the top of the canopy of 134 crested wheatgrass Triticum aestivum (Poales: Poaceae) and alfalfa Medicage sativa (Fabales: Fabaceae), respectively. Similar results were reported with M. anisopliae on cowpea leaves (Ekesi et al., 2001). The decrease in viability found in this study, is likely to favour spot spray application as smaller quantities of inoculum is used compared to cover spray applications. The first season was characterized by a low M. sjostedti population but the crop was highly infested by cowpea aphid Aphis craccivora (Hemiptera: Aphididae). Since no action was taken to control this insect pest, the grain yield was not recorded. As a result, the cowpea yield was recorded and the cost benefit analysis was calculated for the second season only. The cowpea yield in plots that received the spot and cover spray applications increased respectively with 42.6 and 34.1. The grain yields obtained in fungus treatments in this study are similar to yields reported by Saxena and Kidiavai (1997) following three applications of cypermethrin to the cowpea crop during the long rainy cropping season at the same site. 6.5 Conclusion The current study showed that spot spray application of EPF in combination with an attractant is as effective as cover sprays in reducing M. sjostedti populations on cowpea. It requires less inoculum and labour as compared to cover spray applications. Moreover, spot spray applications recorded higher rates of returns than cover spray and this could increase over time with depreciation of materials used. This approach seems to be cost-effective and could be adopted by small-scale farmers if disseminated at large scale through technology transfer and sensitization campaigns. 135 6.6 References Abate, T. and Ampofo, J.K. (1996). Insect pests of beans in Africa: Their ecology and management. Annual Review of Entomology 41, 45-73. Abate, T., Alene, A.D., Bergvinson, D., Shiferaw, B., Silim, S., Orr, A. and Asfaw, S. (2012). Tropical grain legumes in Africa and South Asia. Knowledge and opportunities. ICRISAT-CIAT-IITA, Nairobi, Kenya. Adipala, E., Omongo, C.A., Sabiti, A., Obuo, J.E, Edema, R., Bua, B., Atyang, A., Nsubuga, E.N. and Ogenga-latigo, M.W. (1999). Pests and diseases on cowpea in Uganda: Experiences from a diagnostic survey. African Crop Science Journal 7, 465-478. Ajeigbe, H.A., Adamu, R.S. and Singh, B.B. (2012). Yield performance of cowpea as influenced by insecticide types and their combinations in the dry savannas of Nigeria. African Journal of Agricultural Ressources 7, 5930 -5938. Alves, S.B., Pereira, R.M., Lopes, R.B. and Tamai, M.A. (2002). Use of entomopathogenic fungi in Latin America. (In: Control of insect pests. Upadhyay, R.K., ed. Kluwer Academic/Plenum Publishers, p. 193-211). Belmain, S.R., Haggar, J., Holt, J. and Stevenson, P.C. (2013). Managing legume pests in sub- Saharan Africa. (In: Challenges and prospects for improving food security and nutrition through agro-ecological intensification. Chatham Maritime (United Kingdom). Natural Resources Institute, University of Greenwich. 34p). Broughton, S., Cousins, D.A. and Rahman, T. (2015). Evaluation of semiochemicals for their potential application in mass trapping of Frankliniella occidentalis (Pergande) in roses. Crop Protection 67, 130-135. 136 Ekesi, S. and Maniania, N.K. (2007). Use of entomopathogenic fungi in biological pest management Research Signpost, Kerala, India, 330p . Ekesi, S., Maniania, N.K., Ampong-Nyarko, K. and Akpa, A.D. (2001). Importance of timing of application of the entomopathogenic fungus, Metarhizium anisopliae for the control of legume flower thrips, Megalurothrips sjostedti and its persistence on cowpea. Archives of Phytopathology and Plant Protection 33, 431-445. Ekesi, S., Maniania, N.K., Ampong-Nyarko, K. and Onu, I. (1999). Effect of intercropping cowpea with maize on the performance of Metarhizium anisopliae against the legume flower thrips, Megalurothrips sjostedti, and some predators. Environemental Entomology 28, 1154-1161. Ekesi, S., Maniania, N.K., Ampong-Nyarko, K. and Onu, I. (1998). Potential of the entomopathogenic fungus, Metarhizium anisopliae (Metsch.) Sorokin for control of the legume flower thrips, Megalurothrips sjostedti (Trybom) on cowpea in Kenya. Crop Protection 17, 661-668. El-Deep Soha, M. (2014). The partial budget analysis for sorghum farm in Sinai Peninsula, Egypt. Annals of Agricultural Science 59, 77-81. Ezueh, M.I. (1981). Nature and significance of pre-flowering damage by thrips to cowpea. Entomologia Experimentalis et Applicata 29, 305-312. Fargues, J., Goettel, M.S., Smits, N., Ouedraogo, A., Vidal, C., Lacey, L.A., Lomer, C.J. and Rougier, M. (1996). Variability in susceptibility to simulated sunlight of conidia among isolates of entomopathogenic Hyphomycetes. Mycopathologia 135, 171-181. 137 Inglis, G.D., Enkerli, J. and Goettel, M.S. (2012). Laboratory techniques used for entomopathogenic fungi:Hypocreales. (In: Manual of techniques in invertebrate pathology, 2nd edn. Lacey, L.A., ed. Academic Press, San Diego, p. 189–253). Jackai, L.E.N. and Daoust, R.A. (1986). Insect pests of cowpeas. Annual Review of Entomology 31, 95-119. Jaronski, S.T. (2010). Ecological factors in the inundative use of fungal entomopathogens. BioControl 55, 159-185. Katungi, E., Farrow, A., Chianu, J., Sperling, L. and Beebe, S. (2009). Common bean in Eastern and Southern Africa: A situation and outlook analysis. International Centre for Tropical Agriculture(CIAT). Southern Africa Bean Research Network (SARN). Kiprotich, M.J., Mamati, E., Bikketi, E. (2015). Effect of climate change on cowpea production in Mwania watershed: A case of Machakos County. Journal of Research in International Edution 3, 287-298. Löhr, B. (1996). Integrated Pest Management in French beans in Kenya: pastachieved and some thoughts about the flower thrips problem. (In: Proceedings of the Biennial Crop Protetion Conference, 27-28 March, 1996, Nairobi. Kenya Agriculatural Research Insitute, Nairobi). Mergeai, G., Kimani, P., Mwang’ombe, A., Olubayo, F., Smith, C., Audi, P., Baudoin, J.-P. and Le Roi, A. (2001). Survey of pigeonpea production systems, utilization and marketing in semi-arid lands of Kenya. Biotechnology, Agronomy, Society and Environment 5, 145- 153. Mfuti, D.K., Subramanian, S., van Tol, R.W.H.M., Wiegers, G.L., De Kogel, W.J., Niassy, S., Du Plessis, H., Ekesi, S. and Maniania, N.K. (2016). Spatial separation of semiochemical 138 Lurem-TR and entomopathogenic fungi to enhance their compatibility and infectivity in an autoinoculation system for thrips management. Pest Management Science, 72: 131- 139. Migiro, L.N., Maniania, N.K., Chabi-Olaye, A. and Van den Berg, J. (2010). Pathogenicity of entomopathogenic fungi Metarhizium anisopliae and Beauveria bassiana (Hypocreales: Clavicipitaceae) isolates to the adult pea leafminer (Diptera: Agromyzidae) and prospects of an autoinoculation device for infection in the field. Environmental Entomology 39, 468-475. Muvea, A.M., Waiganjo, M.M., Kutima, H.L., Osiemo, Z., Nyasani, J.O. and Subramanian, S. (2014). Attraction of pest thrips (Thysanoptera: Thripidae) infesting French beans to coloured sticky traps with Lurem-TR and its utility for monitoring thrips populations. International Journal of Tropical Insect Science 34, 197-206. Nderitu, J.H., Wambua, E.M., Olubayo, F., Kasina, J.M. and Waturu, C.N. (2007). Management of thrips (Thysanoptera: Thripidae) infestation on french beans (Phaseolus vulgaris L.) in Kenya by combination of insecticides and varietal resistance. Journal of Entomology 4, 469-473. Niassy, S., Maniania, N.K., Subramanian, S., Gitonga, L.M. and Ekesi, S. (2012). Performance of a semiochemical-baited autoinoculation device treated with Metarhizium anisopliae for control of Frankliniella occidentalis on French bean in field cages. Entomologia Experimentalis et Applicata 142 97-103. Nyasani, J.O., Meyhöfer, R., Subramanian, S. and Poehling, H.-M. (2013). Seasonal abundance of western flower thrips and its natural enemies in different French bean agroecosystems in Kenya. Journal of Pest Science 86, 515-523. 139 Pearsall, I.A. and Myers, J.H. (2000). Population dynamics of western flower thrips (Thysanoptera: Thripidae) in British Columbia. Journal of Economic Entomology 93, 264-275. Quin, F.M. (1997). Introduction. (In: Advances in cowpea research. Co-publication of International Institute of Tropical Agriculture (IITA) and Japan International Research Center for Agricultural Sciences (JIRCAS). Singh, B.B., Mohan Raj, D.R., Dashiell, I.E., Jackai, L.E.N.,ed. Ibadan, Nigeria, p. ix-xv). R Core Development team. (2014). A Language and environment for statistical computing R. Foundation for statistical computing, Vienna, Austria, 2014. Samuel, G.M. and Graham, A.M. (2003). Recent developments in sprayers for application of biopesticides. An overview. Biosystems Engineering 84, 119-125. Saxena, R.C. and Kidiavai, E.L. (1997). Neem seed extract spray applications as low-cost inputs for management of the flower thrips in the cowpea crop. Phytoparasitica 25: 99-110. Shapiro, S.S. and Wilk, M.B. (1965). An analysis of variance test for normality (complete samples). Biometrika 52, 591-611. Singh, S.R. and Allen, D.J. (1980). Pests, diseases, resistance and protection in cowpea. (In: Advances in Legume Science. Summerfield, R.A. and Bunting, H.H., eds. Royal Botanical Garden, Kew, Ministry of Agriculture, Fisheries and Food, p. 419-433). Snedecor, G.W. and Cochran, W.G. (1989). Statistical methods, Eighth ed. University Press, Iowa State. Teulon, D.A.J., Castañé, C., Nielsen, M.-C., El-Sayed, A.M., Davidson, M.M., Gardner-Gee, R., Poulton, J., Kean, A.M., Hall, C., Butler, R.C., Sansom, C.E., Suckling, D.M. and Perry, N.B. (2014). 4-Pyridyl carbonyl and related compounds as thrips lures: effectiveness for 140 onion thrips and New Zealand flower thrips in field experiments. Journal of Agriculture and Food Chemistry 55, 6198-6205. Ugine, T.A., Wraight, S.P. and Sanderson, J.P. (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. 141 CHAPTER 7: GENERAL DISCUSSION, CONCLUSIONS AND RECOMMENDATIONS 7.1. General discussion In eastern Africa and in Kenya in particular, chemical control remains the main option for the management of the bean flower thrips (BFT), Megalurothrips sjostedti (Trybom) (Thysanoptera: Thripidae). However, the frequent use of these synthetic chemical insecticides is associated with several problems such as health risks to the users and consumers, non-target organisms and environmental contamination, in addition to resistance of thrips to these chemical insecticides (Oparaeke, 2006; Alao et al., 2011). Biological control using EPF is among the alternatives being considered (Ekesi and Maniania, 2007). EPF are generally applied using inundative sprays. However, this approach has a number of disadvantages including short persistence of the inoculum due to detrimental effects of solar radiation and high costs as a result of repeated applications (Inglis et al., 2000; Leland and Behle, 2004; Jaronski, 2010). In order to overcome these shortcomings, autodissemination devices have been developed for the management of several insects including thrips (Niassy et al., 2012). Autodissemination consists of attracting insects, using visual and chemical cues to a killing agent, namely an entomopathogenic fungi in this study. Attracted insects are infected with fungal conidia before returning to the environment where they can infect mates during mating or casual contacts. For thrips, the attractant is both visual (blue color) and may also be through a kairomone such as Lurem-TR, a methyl isonicotinate. However, Lurem-TR was found to negatively affect conidial viability of the M. anisopliae isolate ICIPE 69 (Niassy et al., 2012). Therefore, there is need to find the most appropriate way to combine the two agents or to identify 142 attractants that are compatible with M. anisopliae and which could be used both in autodissemination and “spot spray” approaches for the management of bean flower thrips (BFT), Megalurothrips sjostedti on grain legumes. The objective of this study was to develop efficient, economical and sustainable strategies for the management of M. sjostedti on grain legumes using the “lure and infect’’ approach. Influence of spatial separation of the semiochemical on thrips attraction and conidial acquisition by thrips from the autoinoculation device was investigated in the laboratory and in the field (Chapter 3). The results of this study demonstrated that the persistence of conidia of M. brunneum and M. anisopliae increased with distance of separation from Lurem-TR. Direct exposure of the fungus without separation from Lurem-TR recorded the lowest conidial germination as compared with the other treatments. Attraction of thrips to the device also varied significantly according to distance between the device and the semiochemical, with a higher number of thrips attracted when Lurem-TR was placed in a container below the device and at a 10 cm distance. No significant difference in conidial acquisition by M. sjostedti resulted from differences between spatial separation treatments of conidia and Lurem-TR. Positive correlations were found between conidial acquisition and thrips attraction. These results have confirmed the antifungal effect of Lurem-TR on the fungus as reported earlier by Niassy et al. (2012). Spatial separation of fungal conidia from Lurem-TR in an autoinoculation device could therefore provide a low-cost strategy for effective management of thrips in grain legume cropping systems. Seven thrips attractants were screened in the laboratory for their compatibility with M. anisopliae isolate ICIPE 69, in terms of conidial germination and germ tube length, before possible 143 integration in autodissemination devices (Chapter 4). Conidial germination of M. anisopliae was significantly higher and germ tube length, significantly longer in the control, followed by methyl anthranilate, cis-jasmone and trans-caryophellene, and were found to be compatible with M. anisopliae. The lowest conidial germination and shortest germ tube length were obtained when conidial spores were exposed to Lurem-TR which further confirmed its antifungal properties (Niassy et al., 2012; Mfuti et al., 2016). Under field conditions, methyl anthranilate was as attractive as Lurem-TR to M. sjostedi. Methyl anthranilate has been reported to be attractive to four other flower thrips species, namely Thrips hawaiiensis, T. coloratus, T. flavus, and M. distalis, irrespective of sex (Murai et al., 2000; Imai et al., 2001), and it could therfore be considered as a potential attractant candidate. The evaluation of semiochemical baited aoutinoculation devices (methyl anthranilate and Lurem- TR) in a field experiment resulted in a significant reduction in M. sjostedti numbers in cowpea plots (Chapter 5). Conidial viability of M. anisopliae was, however, significantly higher in semiochemical-free baited devices (control) than in semiochemical-baited devices in both seasons. The average number of conidia acquired by single M. sjostedti varied between 2.0 and 10.0 x 103 conidia in both semiochemical-baited device treatments during both seasons and was not significantly different, except for the podding stage in the second season where significant differences were found between treatments. Mortality of M. sjostedti in the two semiochemical- baited treatments ranged between 40-46%. Dimbi et al. (2003) reported Ceratitis rosa (Karsch) and C. fasciventris (Bezzi) (Diptera: Tephritidae) mortality of 70-93% and Niassy et al. (2012) 59% mortality of Frankliniella occidentalis (Thysanoptera: Thripidae) in the presence of a semiochemical in field cages. Similarly to conidial acquisition, the lowest level of mortality in 144 the, current study despite the presence of a semiochemical can be ascribed to infection levels under field conditions as compared to field cages. Cowpea yield varied significantly between treatments, with the highest yield recorded in plots containing a methyl anthranilate-baited device. However, no significant difference in the yield of plots containing semiochemical free and Lurem-TR-baited devices was found. It can therefore be recommended from this study that methyl anthranilate should be used in autoinoculation devices for the management of M. sjostedti on grain legumes. The efficacy of M. anisopliae applied in spot and cover sprays in combination with the thrips attractant, Lurem-TR was also evaluated in the field for two seasons (Chapter 6). Megalurothrips sjostedti densities were lower in spot and cover spray treatments than in the control treatment in both seasons, resulting in a yield increase. However, the cost benefit analysis following a procedure by El-Deep Soha (2014) procedure suggests that a spot spray application was more profitable due to a reduction in labour cost and the quantity of inoculum used. 7.2. Conclusions With regard to the results obtained from all the objectives, we concluded that: (i) Separating fungal conidia from Lurem-TR in an autoinoculation device could provide effective management of thrips in grain legume cropping systems. (ii) Considering the attraction of other insect pests such as whiteflies, bean flies and leafminer flies to semiochemical-baited autoinoculation devices, the strategy could be an alternative control option not only for thrips but also for other insect pests of grain legumes. 145 (iii) Methyl anthranilate is as effective as Lurem-TR and can be recommended as an alternative thrips attractant for use in autoinoculation devices and spot spray applications in combination with M. anisopliae for the control of M. sjostedti. (iv) Spot spray is a more profitable application strategy than cover spray of M. anisopliae due to a reduction in labour cost and the quantity of inoculum used. It should therefore be a more viable option for small-scale farmers and adoption should be facilitated by technology transfer campaigns. 7.3. Recommendations (i) Since the autoinoculation strategy is effective in controlling thrips and other insect pests of grain legumes, surveys should be carried out to investigate farmers’ perception to enable an eventual adoption of the strategy by farmers. (ii) Methyl anthranilate could be used in spot spray applications of M. anisopliae or in the autoinoculation strategy for the control M. sjostedti. (iii) The prototype of an autoinoculation device designed up to now seems to be cumbersome. A need therefore exists to develop a simple design of an autoinoculation device. The semiochemical included in the spot spray strategy is expensive. Potential for its reuse and replacement of the semiochemical with a colour attractant alone need to be investigated. (iv) Electrostatic formulation of conidia of the fungus should also be considered as it could improve the adherence of conidia to the host cuticle. 146 7.4. References Alao, F.O. and Adebayo, T.A. (2011). 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