Efficacy of entomopathogenic nematodes for control of Tuta absoluta in South Africa O Coleman orcid.org 0000-0002-3383-8980 Dissertation accepted in fulfilment of the requirements for the degree Master of Science in Environmental Sciences with Integrated Pest Management at the North-West University Supervisor: Prof MJ Du Plessis Co-supervisor: Prof H Fourie Graduation October 2020 24093807 i ACKNOWLEDGEMENTS I want to express my sincere appreciation and gratitude to my supervisor and co-supervisor, Prof Hannalene du Plessis and Prof Driekie Fourie, who provided me with this great opportunity to further my studies. I am very grateful for the excellent guidance throughout my M.Sc. study culminating in the writing of this dissertation. Their patience, motivation and the guidance are greatly appreciated. Prof A.P. Malan, from the Stellenbosch University, for her guidance, patience and advice; without it we would have been at a great loss. Also, a big thank you for the provision of all the EPN specimens which she always provided on request for use in this study. I also want to thank Mrs. Helena Strydom for her patience with all my requests and the administrative assistance she provided. I would like to say thank you to my parents, Mariana and JC Coleman for standing behind me in the pursuit of my studies. The patience and support provided by you have kept me going and motivated to finish my M.Sc. The love and guidance you have provided me has made me the person I am today. I also want to thank my family, friends and colleagues for the support and encouragement throughout my studies: The people from the Integrated Pest Management program of the North-West University, standing together and helping each other. A special thanks to my friend Brendon Mann for always providing words of encouragement and keeping me motivated when times seemed bleak. My sisters that always believed in me and for having the highest admiration for what I have accomplished. ii DECLARATION BY THE CANDIDATE I, O. Coleman, declare that the work presented in this MSc thesis is my own work, that it is not been submitted for any degree or examination at any other University and that all the sources I have used or cited have been acknowledged by the complete reference. Signature………………………………….. Date……1…8/0…5/…20…20… …………… DECLARATION AND APPROVAL BY SUPERVISORS We declare that the work presented in this thesis was carried out by the candidate under our supervision and we approve this submission. Prof MJ du Plessis Unit for Environmental Sciences and Management, North West University, Private Bag, X6001, Potchefstroom, 2520, South Africa. Signature … …………………………… Date 18/05/2020………. Prof H. Fourie Unit for Environmental Sciences and Management, North West University, Private Bag, X6001, Potchefstroom, 2520, South Africa. Signature …………………………….. Date …18… M…a…y 2…0…20… …… iii ABSTRACT The South American tomato pinworm, Tuta absoluta (Meyrick) (Lepidoptera: Gelechiidae) is one of the most devastating pests of tomato (Solanum lycopersicon L.) in South America, Europe, the Middle East and Africa. Current management tactics of T. absoluta consist mainly of monitoring with sex pheromone traps and application of insecticides. Resistance to various insecticide groups has, however, been reported in Asia, Europe and South America. Development of integrated pest management (IPM) strategies for this pest is therefore important. There is currently no tomato cultivar commercially available which is resistant to T. absoluta, and parasitoids from only four families are known as biological control agents of T. absoluta in Africa. A variety of insect pests are controlled with entomopathogens such as fungi, bacteria and nematodes, which are used as biopesticides. Although entomopathogenic nematodes (EPNs) were initially applied as soil applications against pests, investigations to use EPNs as foliar applications also received renewed interest. In Europe, Heterorhabditis bacteriophora Poinar, 1976, Steinernema feltiae (Filipjev, 1934) Wouts, MráÏcek, Gerdin and Bedding, 1982, and Steinernema carpocapsae (Weiser, 1955) Wouts, MráÏcek, Gerdin and Bedding, 1982 have been reported to effectively control T. absoluta as foliar applications. The Agricultural Pest Act 36 of South Africa, prohibits importation of exotic species without a full impact study and permit. A search for native biological control agents for T. absoluta is therefore warranted. The aim of this study was to evaluate the efficacy of four native EPN species, viz. Steinernema jeffreyense Malan, Knoetze and Tiedt, 2015, Steinernema yirgalemense Nguyen, Tesfamariam, Gözel, Gaugler and Adams, 2005, Heterorhabditis baujardi Phan, Subbotin, Nguyen and Moens, 2003 and Heterorhabditis noenieputensis Malan, Knoetze and Tiedt et al., 2014 against T. absoluta in South Africa. Fourth instar T. absoluta larvae and pupae were exposed to IJs of the four EPN species in vitro. All four EPNs were found to be highly effective in controlling the larvae, with 100% larval morality caused, but pupae were less susceptible. Following the successful in vitro assays using the EPNs against T. absoluta larvae, greenhouse trials were conducted. Efficacy of S. jeffreyense and S. yirgalemense applied to the foliage of tomato seedlings for control of third and fourth instar T. absoluta larvae, was evaluated at four concentrations, viz. 250, 500, 1 000 and 2 000 IJs mL-1 distilled water containing 0.05% adjuvant (Nu-Film-P). High mortality rates of T. absoluta larvae in tomato leaves were recorded with both species at application rates of 1 000 and 2 000 IJs mL-1. Results from this study identified S. jeffreyense and S. yirgalemense as promising biocontrol agents of T. absoluta under greenhouse tomato production in South Africa, which could be included in IPM of this pest. By applying an IPM system and not relying on chemical control only, resistance to insecticides in South Africa, may be prevented or delayed. Key words: Biological control, biopesticide, EPNs, Integrated Pest Management, tomato leafminer TABLE OF CONTENTS ACKNOWLEDGEMENTS ...................................................................................................... i DECLARATION BY THE CANDIDATE ................................................................................ ii ABSTRACT ......................................................................................................................... iii Chapter 1: Introduction and Literature review .................................................................. 1 1.1. General introduction ....................................................................................................... 1 1.2. Tuta absoluta (Meyrick) (Lepidoptera: Gelechiidae) ....................................................... 3 1.2.1. Biology of Tuta absoluta and damage symptoms ..................................................... 6 1.3. Control strategies for Tuta absoluta................................................................................ 8 1.3.1. Chemical control ...................................................................................................... 8 1.3.2. Cultural control ...................................................................................................... 10 1.3.3. Host plant resistance ............................................................................................. 11 1.3.4. Biological control ................................................................................................... 12 1.4. Entomopathogenic nematodes ..................................................................................... 14 1.4.1. Entomopathogenic nematodes as biological control agents ................................... 17 1.4.2. Entomopathogenic nematodes used for the control of Tuta absoluta ..................... 22 1.4.3. Entomopathogenic nematodes in South Africa ...................................................... 22 1.5. Aim and Objectives ...................................................................................................... 27 1.6. References................................................................................................................... 27 Chapter 2: Efficacy of indigenous entomopathogenic nematodes (Rhabditida: Heterorhabditidae and Steinernematidae) against Tuta absoluta (Lepidoptera: Gelechiidae) in laboratory bioassays .............................................................................. 39 2.1. Introduction .................................................................................................................. 39 2.2. Material and methods ................................................................................................... 43 2.3. Results ......................................................................................................................... 45 2.4. Discussion.................................................................................................................... 48 2.5. References................................................................................................................... 50 Chapter 3: Efficacy of Steinernema yirgalemense and Steinernema jeffreyense applied as foliar applications for control of Tuta absoluta on tomato under greenhouse conditions in South Africa ................................................................................................ 58 3.1. Introduction .................................................................................................................. 58 3.2. Materials and Methods ................................................................................................. 60 2 3.3. Results ......................................................................................................................... 62 3.4. Discussion.................................................................................................................... 64 3.5. References................................................................................................................... 66 Chapter 4: Conclusion and recommendations ............................................................... 70 4.1. References................................................................................................................... 73 Appendix A ........................................................................................................................ 79 Appendix B ........................................................................................................................ 80 Appendix C ........................................................................................................................ 81 1 Chapter 1: Introduction and Literature review 1.1. General introduction Tomato (Solanum lycopersicon L.) is one of the most widely cultivated and economically important crops worldwide (Tropea Garzia et al., 2012). It has been cultivated on more than 5.7 million hectares, with a global production of approximately 233 million tons reported in 2018 (FAOSTAT, 2018). The total tomato production in the United States of America (USA) during this period was 14 million tons and the South African production, 580 000 tons (FAOSTAT, 2018). Tomato fruit is rich in lycopene (Falara et al., 2011; Fernández-Ruiz et al., 2011), a carotenoid with several important health benefits for humans (Clinton, 1998). It is a powerful antioxidant that acts as an anti-carcinogen (Kelley and Boyhan, 2006). Tomato is also rich in vitamins and minerals and one medium ripe tomato can provide up to 40% of the recommended daily allowance (RDA) of Vitamin C and 20% of RDA Vitamin A (Kelley and Boyhan, 2006). The B vitamins, potassium, iron and calcium needed in the human diet, are also found in tomato (Kelley and Boyhan, 2006). The commercial value of the fruit includes processed foods as well as selling it on the fresh market (Kelley and Boyhan, 2006). Weeds are serious competitors for environmental resources and if left unchecked can reduce crop yield and quality considerably (COPR, 1983; Ozores-Hampton et al., 2001). Fungal and bacterial diseases, viruses and mycoplasma-like diseases also contribute to lower yields and production of tomato crops (COPR, 1983). Various ecto- and endoparasitic nematodes, Phylum Nematoda (Rudolphi, 1808) Lankester, 1877, are economically important pests of tomato, causing excessive damage to crops worldwide (Greco and Di Vito, 2011; Seid et al. 2015; Jones et al., 2017). Ectoparasites listed as major pests include species of stubby-root (Paratrichodorus Siddiqi, 1974) and dagger (Xiphinema Cobb, 1913) nematodes as well as genera and species of the stunt nematodes (Dolichodoridae Chitwood, 1950) (Greco and Di Vito, 2011). By contrast the migratory, endoparasitic lesion nematodes Pratylenchus Filipjev, 1936 and Ditylenchus dipsaci (Kühn, 1857) Filipjev, 1936, sedentary counterpart species of cyst (Globodera Skarbilovich, 1959), root-knot (Meloidogyne Göldi, 1887) nematodes, the false root-knot (Nacobbus aberrans (Thorne, 1935) Thorne and Allen, 1944) and reniform (Rotylenchulus reniformis Linford and Oliveira, 1940) nematodes are major pests of tomato (Greco and Di Vito, 2011). Root-knot nematodes are, however, regarded as the most damaging genus that infect roots of tomato crops. The four species generally known to cause damage in tropical and subtropical areas being Meloidogyne arenaria (Neal, 1889) Chitwood, 1949, Meloidogyne enterolobii Yang and Eisenback, 1983, Meloidogyne incognita (Kofoid and White, 1919) Chitwood, 1949 and Meloidogyne javanica (Treub, 1885) Chitwood, 1949. In 2 temperate parts of the world Meloidogyne chitwoodi Golden, O'Bannon, Santo and Finley, 1980, Meloidogyne fallax Karssen, 1996 and Meloidogyne hapla Chitwood, 1949 cause damage to the crop (Greco and Di Vito, 2011; Seid et al. 2015). There are between 100 and 200 insect pest species globally recorded on tomato (Attwa et al., 2015). In South Africa, more than 45 insects were listed as pests on tomato (Table 1.1) (Visser, 2015). Table 1.1: Most common tomato insect pests in South Africa from Visser (2015). Order Pest species common name Scientific name Coleoptera Family Scarabaeidae Fruit and root chafer beetles Anoplocheilus friguratus Boheamn Dischista cincta (DeGeer) Pachnoda sinuata (Fabricius) Pedinorrhina trivittata (Schaum) Porphyronata Hebreae (Olivier) Tephraea dichroa (Schaum) Tephraea leucomelona (Gory and Percheron) Family Coccinellidae Potato ladybird Epilachna dregei Mulsant Solanum ladybird Henosepilachna hirta (Thunberg) Diptera Family Agromyzidae Potato leaf miner Liriomyza huidobrensis (Blanchard) Amperican leaf miner Liriomyza trifolii (Brugess) Hemiptera Family Aphididae Black bean aphids Aphis fabae Scopoli Potato aphid Macrosiphum euphorbiae (Thomas) Green peach aphid Myzus persicae (Sulzer) Family Petatomidae Green vegetable bug Nezara viridula (Linnaeus) Family Lygaeidae Milkweed bugs Spilostethus pandurus (Scopoli) Family Coreidae Large black tip wilter Anoplocnemis curvipes (Fabricius) Common tip wilter Elasmopoda valga (Linnaeus) Family Aphididae Sweet potato whitefly Bemisia tabaci (Gennadius) Greenhouse whitefly Trialeurodes vaporiorum (Westwood) Lepidoptera Family Noctuidae African bollworm Helicoverpa armigera (Hübner) Black cutworm Agrotis ipsilon (Hufnagel) Brown cutworm Agrotis longidentifera (Hampson) 3 Common cutworm Agrotis segetum (Denis and Grey cutworm Schiffermüller) Agrotis subalba Walker Lesser armyworm Spodoptera exigua (Hübner) Potato tuber moth Phthorimaea operculella (Zeller) Scrobipalpa aptatella (Walker) Tomato semi-looper Chrysodeixis acuta (Walker) Plusia semi-looper Thysanoplusia orichalcea (Fabricius) Tomato moth Spodoptera littoralis (de Boisduval) Thysanoptera Family Thripidae Western flower thrips Frankliniella occidentalis (Pergande) Kromnek thrips Frankliniella schultzei (Trybom) Onion thrips Thrips tabaci Lindeman Since the list was compiled by Visser in 2015, the tomato leafminer, Tuta absoluta (Meyrick) (Lepidoptera: Gelechiidae) invaded South Africa. 1.2. Tuta absoluta (Meyrick) (Lepidoptera: Gelechiidae) Tuta absoluta was first detected in South Africa in August 2016 (Visser et al., 2017). The pest originates from South America (Desneux et al., 2010) and has initially been introduced into Spain late in 2006 after which it spread quickly in the European countries (Loni et al., 2011). It also spread throughout the Mediterranean Basin, parts of Northern Africa and the Middle East (Urbaneja et al., 2012) and downwards into southern Africa (Biondi et al., 2018). It has been reported as one of the most devastating pests on tomato in South America (Desneux et al., 2010), Europe (Loni et al., 2011), the Middle East and Africa (Abbes et al., 2016). A map showing the distribution of T. absoluta in 2018 is provided in Figure 1.1. The distribution of T. absoluta now extends to latitudes that are well above its original habitat (South America) (Fig. 1.1), indicating the ability of this pest to adapt and acclimatise. For a species to establish in any newly invaded area, it needs climatic conditions suitable for its survival, availability of food sources as well as the ability to survive other stress related factors that occurs with transmission to a new environment (Renault et al., 2018). Tuta absoluta is considered a typical invasive species, due to its capacity to develop very quickly in suitable agro-ecological conditions, its ability to spread rapidly and to cause economic damage in new areas (Desneux et al., 2010; Tropea Garzia et al., 2012). Since its invasion, it has caused serious crop losses and has become an agricultural threat to crops in these areas (Desneux et al., 2010). 4 Tuta absoluta is an oligophagous insect and feeds almost exclusively on plants belonging to the Solanaceae family (Siqueira et al., 2000; Sannino and Espinosa, 2010a). The Solanaceae includes important crops such as potato (Solanum tuberosum L.), eggplant (Solanum melongena L.) and pepper (Capsicum annuum L.) (Tropea Garzia et al., 2012; Cocco et al., 2015, Hernández-Ruiz and Arnao, 2016). Tuta absoluta feeds mainly on tomato plants but other cultivated solanaceous species can also act as hosts for this pest (Cocco et al., 2015). The widespread cultivation of tomato, potato and other naturally occurring solanaceous crops around the world together with the mild European to subtropical African climates, allowed T. absoluta moths to be introduced and established in new areas (Sannino and Espinosa, 2010b). Figure 1.1: Geographic distribution of Tuta absoluta in 2018 (From Bondi et al., 2018). Despite its preference for Solanaceous vegetables, some weeds have also been reported to act as hosts for this insect pest in several countries (Unlu, 2012). Reported host plants of T. absoluta are provided in Table 1.2. 5 Table 1.2: Host plants reported for Tuta absoluta. Plant family Host plants Common name Reference Crop host plants Solanaceae Lycopersicon esculentum Tomato Park et al. (2004) Linnaeus Desneux et al. (2010) Hernández-Ruiz and Arnao (2016) Solanum tuberosum Potato Megido et al. (2013) Linnaeus Solanum melongena Eggplant Megido et al. (2013) Linnaeus Nicotiana tabacum Tobacco Megido et al. (2013) Linnaeus Physalis peruviana Cape gooseberry USDA-APHIS (2011). Linnaeus Capsicum annuum Pepper USDA-APHIS (2011) Linnaeus Solanum muricatum Sweet pepper Desneux et al. (2010) Linnaeus Alternative host plants Fabaceae Phaseolus vulgaris Common beans Mohamed et al. (2015) Linnaeus Vicia faba Linnaeus Broad bean Mohamed et al. (2015) Vigna unguiculata Cowpea Mohamed et al. (2015) (Linnaeus) Walpers Medicago sativa Linnaeus Alfalfa Mohamed et al. (2015) Solanaceae Lycium barbarum Linnaeus Goji berry Mohamed et al. (2015) Brassicaceae Raphanus raphanistrum Wild radish Mohamed et al. (2015) Linnaeus Wild host plants Solanaceae Solanum nigrum Linnaeus Night shade Desneux et al. (2010) Solanum elaeagnifolium Silverleaf Desneux et al. (2010) Cavanilles nightshade Mekki (2007) Solanum bonariense Desneux et al. (2010) Linnaeus Solanum sisymbriifolium Desneux et al. (2010) Lamarck Solanum saponaceum Desneux et al. (2010) Welwitsch Bayram et al. (2015) Solanum hirtum Vahl Cocco et al. (2015) 6 Datura stramonium Jimson weed Desneux et al. (2010), Linnaeus Mohamed et al. (2015) Lycium chilense Miers ex Coralillo Mohamed et al. (2015) Bertero Varela et al. (2018) Datura ferox Linnaeus Long spined Desneux et al. (2010) thorn apple Nicotiana glauca Graham Tree tobacco Mohamed et al. (2015) Amaranthaceae Chenopodium album Lambs-quarters Mohamed et al. (2015) Linnaeus Convolvulaceae Convolvulus arvensis Bindweed Mohamed et al. (2015) Linnaeus Alternative host plants maintain this insect pest in the absence of tomato crops. For successful integrated pest management (IPM), the presence and control of these alternative host plants should be taken into consideration (Tropea Garzia et al., 2012). 1.2.1. Biology of Tuta absoluta and damage symptoms Tuta absoluta is a holometabolic insect with four development stages namely eggs, larvae, pupae and moths (Fig. 1.2) (Desneux et al., 2010). Spreading of T. absoluta is attributed to the uncontrolled trading of infested plants and fruit, but spreading by moths also contribute to the repeated and continued infestation in more enclosed areas (Sannino and Espinosa, 2010b). Active flight or passive spreading through wind currents are also means of dispersal (Desneux et al., 2010). The high population growth potential of T. absoluta whereby many generations are produced per season as well as greenhouse crops that create conditions for continuous feeding and reproducing, also contribute to the rapid distribution of the pest. Uncontrolled outbreaks of an introduced pest such as T. absoluta can also be as a result of the lack of natural enemies in newly infested countries as well as the development of resistance to commonly used insecticides (Sannino and Espinosa, 2010b). 7 Larva Egg Pupa Moth Figure 1.2: Life cycle of Tuta absoluta (Photo’s: Odette Coleman, NWU). Tuta absoluta completes 10-12 generations per year in South America and overwinters in all its life stages (Cocco et al., 2015). The life cycle of T. absoluta is completed in approximately 24 days at 27 °C but the development time can vary between 13-65 days depending on temperature (Desneux et al., 2010). Moths are 6-7 mm in length with silver to grey scales and the antennae are filiform (Fig. 1.3A) (Desneux et al., 2010). Female moths live for 10-15 days and males for 6-7 days, but moths can live for up to 40 days at 10 C (Cuthbertson et al., 2013). The females can mate once a day for 4-5 h, and up to six times during their life span (Desneux et al., 2010; Tropea Garzia et al., 2012). The longer life span of female moths allows them to become sexually mature before the males emerge (Tropea Garzia et al., 2012). On tomato, eggs are laid on leaves, usually underneath, or on the stems (Desneux et al., 2010). In cases of severe infestation, eggs are laid on the fruit as well (Cocco et al., 2015). Most eggs are laid in the seven days following the first mating, with 70% of the total oviposition occurring during this period (Tropea Garzia et al., 2012). A female can oviposit up to 260 eggs during her lifespan (Tropea Garzia et al., 2012). Larval development time at 27 °C is completed in 11-13 days (Sannino and Espinosa, 2010c). Fully developed, fourth instar larvae (Fig. 1.3B) drop to the soil or pupate in a cocoon on leaves of the host plant (Desneux et al., 2010). Pupae are 5-6 mm in length, cylindrical shaped, greenish after pupation and become darker in colour with time (Cocco et al., 2015). The larvae 8 have a very distinctive black mark behind the head capsule (Sannino and Espinosa, 2010a), are endophytic and briefly move around before mining into the plant (Cocco et al., 2015). Tuta absoluta attacks any stage of tomato plants, from seedlings to mature plants. They feed on the mesophyll tissue of leaves (Fig. 1.3C), flowers, stems and fruit (Fig. 1.3D) (Ubaneja et al., 2012). Mines in leaf stalks and stems can cause plant death (Sannino and Espinosa, 2010c). Larvae tunnel into the leaves. These tunnels expand rapidly and become large chambers leaving the two outer membranes (Fig. 1.3C) (Sannino and Espinosa, 2010c). This damage reduces the photosynthetic capacity of the plant, resulting in fewer fruit and a lower yield (Pereyra and Sánchez, 2006; Ubaneja et al., 2012). A B C D Figure 1.3: (A) Tuta absoluta moth, (B) fourth instar larva, (C) damage caused by larvae to leaves and (D) larval damage to fruit (Photo’s: Odette Coleman, NWU). 1.3. Control strategies for Tuta absoluta The best option for control of T. absoluta is with an IPM strategy (González-Cabrera et al., 2011). It can consist of a single method or a combination of methods to produce an effective management strategy (Kogan, 1998), in order to maintain pest populations below the economic injury level (Dent, 2000). These methods include chemical, biological and cultural control, and host plant resistance (Dent, 2000). Although IPM strategies include chemical control in tomato fields, the aim should be to reduce the overall usage and still maintain effective control (Miranda et al., 2005). More environmentally friendly control measures are also needed to reduce the potential harmful effects that might disrupt existing IPM programs (Urbaneja et al., 2012). 1.3.1. Chemical control The production cost of tomato has increased substantially in countries where crops are infested with T. absoluta due to additional pest control and monitoring strategies (Tropea Garzia et al., 2012). For example, production costs more than tripled in the main tomato production areas of South America due to the increased use of insecticides to control T. absoluta (Guedes and Picanço, 2012). The number of insecticide applications increased from 9 10-12 to over 30 per cultivation period, with approximately four to six insecticide applications per week in commercial fields (Guedes and Picanço, 2012). The year-round cultivation of tomato crops and the short T. absoluta life cycle of 26-38 days in fields, provide for multiple overlapping generations and a challenge to effectively control the pest. It subsequently results in increased levels of infestation and higher losses (Guedes and Picanço, 2012). Tuta absoluta is mainly controlled with insecticides (Loni et al., 2011). Effective chemical control is problematic due to the endophytic feeding of this pest that takes place between the two lamellas of a leaf (Gözel and Kasap, 2015). The efficacy of insecticides applied to tomato crops is compromised when an insect population is already well established before the reproductive stage (Guedes and Picanço, 2012). The main concern with extensive use of insecticides is the quick development of resistance (Guedes and Picanço, 2012; Urbaneja et al., 2012). The high reproduction capacity, short generation cycle of T. absoluta as well as the intensive use of insecticides, enhance and account for the higher risk of resistance development to insecticides (Gözel and Kasap, 2015). Insecticides from four groups with different modes of action, namely diamides, avermectins, spinosyns and oxadiazines are the most extensively used in Europe (Roditakis et al., 2018). Excessive use of insecticides results in selection pressure where resistant genotypes are favoured reducing the efficacy of insecticides (Roditakis et al., 2018). The use of insecticides is therefore not a sustainable management strategy as shown by the efficacy of organophosphates to control T. absoluta which has gradually decreased in many countries since the 1980s (Desneux et al., 2010). The only insecticides that were initially available for T. absoluta control in Argentina were organophosphates, but they were replaced by pyrethoids in 1970 (Lietti et al., 2005). In 1980, as an alternative to pyrethoids, cartap and thiocyclam that were highly effective, were introduced (Lietti et al., 2005). Only a limited number of products within these groups, were available to control T. absoluta before 1990 increasing the risk of resistance development. Resistance to organophosphates and pyrethroids was reported in Chile and Brazil (Desneux et al., 2010). Abamectin, cartap and permethrin were subsequently introduced followed by chitin synthesis inhibitors which was then relied on by producers. This resulted in extreme resistance development against chitin synthesis inhibitors, indoxacarb and spinosad (Guedes and Picanço, 2012). Tuta absoluta resistance development and susceptibility to insecticides in different tomato production localities are not universal and the loss in effectiveness does not occur in all regions (Lietti et al., 2005). Resistance to deltamethrin and abamectin was reported in T. absoluta populations in open fields and greenhouses in Argentina in 2010 (Desneux et al., 2010). Roditakis et al. (2018) conducted a follow up baseline susceptibility study on European T. absoluta and the efficacy of key insecticides. In his study, resistance to chlorantraniliprole was 10 reported from Italy, Greece and Israel. In a few cases resistance to oxadiazines, avermectins and spinosyns was also detected (Roditakis et al., 2018). Registered agrochemicals that can be used against T. absoluta in South Africa are spinetoram, indoxacarb, spinosad, cypermethrin as well as the mixtures, emamectin benzoate and lufenuron and chlorantraniliprole and lambda-cyhalothrin (DAFF, 2017). No resistance to insecticides by T. absoluta has been reported in South Africa to date. Insecticide resistance management (IRM) is applied to delay or prevent resistance development to insecticides or to regain susceptibility to insecticides in an insect pest population (Spark and Nauen, 2015). To reduce selection pressure to an insecticide, methods such as rotation or alteration of insecticides, mixtures and mosaics can be implemented (Spark and Nauen, 2015). Following a basic IPM strategy and implementing IRM can reduce selection pressure which will reduce resistance development enabling products to maintain field efficacy (Roditakis et al., 2018). The use of chemical insecticides is not a sustainable management strategy due to the proven development of resistance by T. absoluta, but also due to the potential effect it has on non-target organisms occurring in fields where tomato crops are planted. Moreover, its adverse effects on animals, humans and the environment lead to numerous pesticides being banned from world markets (PAN, 2019). 1.3.2. Cultural control Cultural control is a strategy that aims to manipulate the environment to render it unfavourable for pests to live or breed in. This method interferes with the ability of the pests to colonise, reproduce or survive (Dent, 2000). Cultural control is the management of an agro-ecosystem to favour crop production and to avoid or reduce pest populations (Bajwa and Kogan, 2004). The main focus of cultural control is therefore to enhance the processes that limit pest invasion and population growth in agro-ecosystems and also limits the potential of an organism to reach pest status (Bajwa and Kogan, 2004). Preventive measures that could be implemented are the destruction of crop remains and the removal of naturally occurring alternative hosts in and around the tomato production area. Such actions, however, increase labour and can become costly (Sannino and Espinosa, 2010d; Guedes and Picanço, 2012). Crop rotation should also be applied and consists of seasonal alteration of one or more crops within a field with the aim to disrupt the biology of the pest that depends on feeding on one of those crops (All, 1999). Overwintering insect pests will then have to colonise a non-host crop which will reduce colonisation of the pest in a field (Dent, 2000). Crop rotation with non-host plants is effective in interrupting the pest cycle and lower pest numbers. Tilling practices can destroy hibernating insect individuals (Sannino and Espinosa, 2010d). Continuous cultivation of tomato crops in tomato production areas throughout the year reduces the effectiveness of 11 this control method because of the continuous availability of hosts (Guedes and Picanço, 2012). Other cultural control management strategies include methods aimed at preventing the pest from mating. Development of a population will be affected and high pest numbers will be prevented. For these methods, either synthetic pheromones for male annihilation and mating disruption are used, or the sterile insect technique (SIT) is applied (Sannino and Espinosa, 2010d; Guedes and Picanço, 2012; Megido et al., 2012). By monitoring insect pests to detect when increases in pest populations occur, an informed decision for application of control measures can be made. Pheromone traps baited with female sex pheromones are very effective in monitoring the increase in population size of T. absoluta and can be used to decide on the best time for control applications in the field (Sannino and Espinosa, 2010d). Correct timing of application is important because early application, before the major outbreak, would not be effective (Sannino and Espinosa, 2010d). The traps are also used for mass capturing to lower numbers of the insect pest (Sannino and Espinosa, 2010d). An important factor with sex pheromone based management and SIT is that the control is aimed at target insect pests which reproduce sexually (Megido et al., 2013). As soon as asexual or parthenogenetic reproduction occur, the efficacy of these control methods are reduced which is the case with T. absoluta since females can reproduce asexually (Megido et al., 2012; 2013). 1.3.3. Host plant resistance Host plant resistance to T. absoluta has not been successfully implemented because tomato yield is lower in resistant cultivars (Guedes and Picanço, 2012). Cultivated tomato is highly susceptible to T. absoluta and even though there are genetic sources of resistance located in germplasm banks, accessions of S. lycopersicum (wild tomato) are the most promising sources of resistance (Bondi et al., 2018). Density of granular trichomes on the leaves and the insecticidal compounds produced by these trichomes fend off T. absoluta. Allelochemicals are mainly focussed on to impair egg laying as well as feeding by larvae (Guedes and Picanço, 2012; Bondi et al., 2018). Tomato cultivars with induced resistance mechanisms activated by larval feeding is also considered because feeding on the plant could activate plant defences against various other insect pests (Bondi et al., 2018). Breeding tomato cultivars resistant to this pest remains a highly considered control management strategy but the development of such varieties is still in progress (Guedes and Picanço, 2012; Bondi et al., 2018). Research on breeding lines with acyl sugars have been conducted and it was found that a high content of these chemicals are the main provider for host plant resistance (Dias et al., 2013; Bondi et al., 2018). Acyl sugars are easily introduced into elite tomato lines with a simple inheritance that are essentially monogenic, containing modifier genes with additive effect (Dias et al., 2013). 12 1.3.4. Biological control Biological control is the use of living organisms that include invertebrates and a wide variety of microbial pathogens, including fungi, bacteria and viruses, as pest control agents to maintain low pest population density. It may operate alone or as a component of integrated pest management (Greathead and Waage, 1983). It is an environmentally safe measure to manage T. absoluta in the countries where it is present. It also includes natural occurring predators and parasitoids that attack this pest (González-Cabrera et al., 2011). Applied by humans, biological control is the deliberate exploitation of natural enemies to control and regulate insect pest populations. It is therefore a human activity to control pests in a more environmentally friendly manner (Hagler, 2000). It is actions of parasites, predators or pathogens that maintain the population density of a pest organism at a density lower than what it would have been in their absence (Ruberson et al., 1999). It can also be defined as the manipulation of natural enemies to control pests in crop systems (Dent, 2000). These enemies can occur naturally or they can be introduced into the cropping system. The strategies used for biological control are the introduction, inundation, augmentation and inoculation of natural enemies as well as their conservation (Greathead and Waage, 1983; Dent, 2000). A biological control strategy could be a long term sustainable strategy to manage T. absoluta (Desneux et al., 2010). Classical biological control is the introduction of exotic natural enemies to control a newly invaded exotic insect pest (Lacey et al., 2001). It presents a permanent control solution with few risks and provides a highly cost effective approach. Introduction of exotic natural enemies of insect pests may take two to three years since it requires intensive research on the target pest and its native complex of natural enemies, as well as selection, collection, rearing and release of ideal candidates (Dent, 2000). This is a long process and not an immediate solution, but when effectively introduced, it would provide long term control of a newly introduced insect pest (Dent, 2000). The introductions of exotic pathogens are not as popular as the introduction of predators and parasitoids (Lacey et al., 2001). Inoculation and augmentation are used in cases where the natural enemies are absent or where the population densities are too low to be effective. The numbers of natural enemies may be increased with the release of laboratory reared insects (Dent, 2000). Augmentation is, however, only a temporary approach and provides only temporary control to keep the population below an economic threshold level. An inoculation biological control approach is a seasonal application which lasts for the duration of the crop, making it the preferred approach over augmentation (Dent, 2000). Inundation is the mass release of the biological control agent, which kills the host fast, but it is usually not persistent (Dent, 2000). Biological control agents used for inundation are pathogens, 13 consisting of viruses, bacteria, fungi as well as EPNs which are formulated into a biopesticide that can be used as an alternative to chemical insecticides (Dent, 2000). Many natural enemies had been described and successfully implemented to control T. absoluta in regions where this pest naturally occurs (Urbaneja et al., 2012). Biological control agents were reported to be successful in effectively suppressing T. absoluta populations (Van Damme et al., 2016). Both predators and parasitoids attack T. absoluta in European and North African countries (Zappalà et al., 2013). Parasitoids attack the egg, larval and pupal stages of this pest (Desneux et al., 2010). Predators are mainly species from the Miridae family and are used as part of IPM strategies (Zappalà et al., 2013). These predators prey mostly on the eggs and to a limited extend on the larvae of the leaf miner (Van Damme et al., 2016). The most commonly used predators against T. absoluta in European greenhouses are Macrolophus pygmaeus (Rambur) (Hemiptera: Miridae) and Nesidiocoris tenuis (Reuter) (Hemiptera: Miridae) (Van Damme et al., 2016). More than 70 species of generalist natural enemies have been reported to attack T. absoluta in the western Palaearctic region (Zappala et al., 2013). The efficacy of entomopathogens for control of T. absoluta is not well documented in South America (Desneux et al., 2010), but Lacey et al. (2001) reported entomopathogens to provide good control of insect pests compared to other biological control agents. The entomopathogen, Bacillus thuringiensis has been reported to have great potential for controlling T. absoluta in tomato greenhouses and fields in the Mediterranean Basin (González-Cabrera et al., 2011). Bacillus thuringiensis is applied with other biological agents to control the early instars of T. absoluta, for example with the predators, M. pygmaeus and N. tenuis (Van Damme et al., 2016). In these instances, B. thuringiensis acts as a complimentary biological control agent. Another group of biological control agents with potential to target the larval stages of T. absoluta, is EPNs (Van Damme et al., 2016). Entomopathogenic nematodes are already applied successfully against weevils, for example the black vine weevil Otiorhynchus sulcatus (Fabricius) (Coleoptera: Curculionidae), the root weevil Diaprepes abbreviatus Linnaeus (Coleoptera: Curculionidae), and the sweet potato weevil Cylas formicarius (Fabricius) (Coleoptera: Brentidae), fungus gnats, Bradysia spp. Winnertz (Diptera: Sciaridae), Lycoriella spp. Frey (Diptera: Sciaridae), grubs (the japanese beetle Popillia japonica Newman (Coleoptera: Scarabaeidae) and the garden chafer Phylloperta horticola (Linnaeus) (Coleoptera: Scarabaeidae) (Půža, 2015). It is also applied against pest organisms that reside in the soil or spend part of their life cycles in the soil (Půža, 2015; Van Damme et al., 2016). Although EPNs are soil-dwelling organisms, they do have the potential to control pests that attack the foliar parts of plants (Van Damme et al., 2016). Their efficacy increased when they are applied in areas that are protected from adverse environmental conditions, e.g. 14 greenhouses, or if the pest lives in a cryptic habitat such as within a leaf (Van Damme et al., 2016). The success rate of EPNs is lower with foliar application because of the exposure to abiotic factors such as UV radiation, desiccation and extreme temperatures. The use of adjuvants increases leaf coverage and persistence of the infective juveniles (IJs), which in turn enhances the use of EPNs against these foliar pests (Zolfagharian et al., 2016; Kamali et al., 2018; Platt et al., 2019a). Nematode species from the families Steinernematidae Travassos, 1927 and Heterorhabditidae Poinar, 1976 are considered as possible control agents for T. absoluta (Gözel and Kasap, 2015). 1.4. Entomopathogenic nematodes Entomopathogenic nematodes occur naturally in soil and act as obligate parasites by attacking different life stages of insects. They therefore have the benefit of not being harmful to animals and plants (Shapiro-Ilan et al., 2006; Zolfagharian et al., 2016; Devi and Nath, 2017). This is a diverse group of organisms that can be found almost anywhere in the world, in any biome such as cultivated fields, forests, grasslands, deserts and even on beaches or in the ocean (Devi and Nath, 2017). These nematodes are safe to use in the environment and are exempted from registration in European countries such as Austria and Germany, North America including the USA and the United Kingdom (UK) (Van Zyl and Malan, 2014; Devi and Nath, 2017; Malan and Ferreira, 2017). Many countries are, however, required to undergo a registration prosess (Ehlers, 2005). The introduction of non-endemic EPNs in South Africa is also not exempted from registration as in most of the European countries, North America or the UK (Van Zyl and Malan, 2014). Malan and Ferreira (2017) stated that: ”according to the amended Act 18 of 1989 (South African Agricultural Pests Act No. 36 of 1947), the introduction of exotic animals such as non-endemic EPN is only allowed under permit, which has to be accompanied by a full impact study”. Research to determine the efficacy of native EPN species is therefore important. Nearly 40 nematode families are associated with insects (Gaugler and Kaya, 1990), but not all cause host mortality and only 23 of these families contain species described as EPNs (Lacey and Georgis, 2012). Research is focused on species which have the potential to act as biological control agents, and these are found in the families Mermithidae Braun, 1883, Tetradonematidae Cobb 1919, Allantonematidae Pereira, 1931, Phaenopsitylenchidae Blinova and Korenchenko, 1986, Sphaerulariidae Lubbock, 1861, Steinernematidae and Heterorhabditidae (Goodey, 1960; Lacey et al., 2001; Poinar et al., 2002; Stock and Hunt, 2005). Heterorhabditis spp. Poinar, 1976 and Steinernema spp. Travassos, 1927 are the best known EPNs that are widely available and used for biological control of many insect pests worldwide (Griffin, 2012). In the family Steinernematidae, the genus Steinernema contains at least 88 identified species (Abate et al., 2017), while the genus Neosteinernema has only one identified species (Devi 15 and Nath, 2017). The family Heterorhabditidae has only one genus, namely Heterorhabditis with 19 identified species (Stock, 2015; Abate et al., 2017). The use of these EPNs against crop pests has shown promise (Griffin, 2012). Steinernema spp. and Heterorhabditis spp. have both parasitoid and pathogenic attributes. Chemoreceptors, a parasitoid characteristic, enables them to actively search for their host while the association with mutualistic bacteria is a pathogenic characteristic (Lacey et al., 2001). The mutualistic bacteria contained within EPNs is the main reason for these nematodes to be such attractive biopesticides in management of insect pests (Akhurts, 1993). The bacteria associated with Steinernema spp. are Xenorhabdus spp., that are motile, gram-negative enterobacteria, and γ-proteobacteria, while Photorhabdus spp. are gram-negative rods that occur in Heterorhabditis spp. (Chaston et al., 2011, Poinar and Grewal, 2012). Although each nematode species is associated with a specific bacterium, the bacterium can be associated with more than one nematode species (Lacey et al., 2001). Steinernema spp. are associated with 20 Xenorhabdus spp., while four Photorhabdus spp. are described to be associated with Heterorhabditis spp. (Devi and Nath, 2017). These associations are synergistic, with Steinernema spp. carrying the bacteria in a specialized vesicle in their intestine (Bird and Akhurst, 1983), whereas in Heterorhabditis spp. the bacteria occur in the foregut and midgut region of the non-feeding IJs (Boermare et al., 1996). Infective juveniles, the only free living stage of Heterorhabditis and Steinernema, is adapted to survive outside the insect host (Mahmoud, 2016) using stored reserves of energy to search for and enter the host (Griffin et al., 2005). They enter their insect hosts through natural openings such as the mouth, anus and spiracles (Kaya and Gaugler 1993; Griffin et al., 2005). Once the IJs have entered the haemocoel of an insect, the symbiotic bacteria are released and suppress the insect’s defence systems. The bacterium multiply and produces a diverse group of components namely bacteriocins, antibiotics, antimicrobials and a scavenger deterrent compound (Chaston et al., 2011). This compound suppresses the growth of antagonistic microorganisms to provide a safe niche, and breaks down the host tissue which causes death of the insect host usually within 24-48 h after infection, and ultimately produces food (Chaston et al., 2011; Devi and Nath, 2017). The EPNs act by feeding on the bacteria and the degrading host tissue, they multiply, develop and reproduce (Griffin, 2012; Mahmoud, 2016). Depending on the size of the host, the nematode completes one generation or more. In the case of Steinernema, the first generation develops into males and females, while in Heterorhabditis the first generation nematodes are hermaphrodites (Hazir et al., 2003; Griffin et al., 2005). When nutrients are depleted, a high nematode population density induces the development of new non-feeding IJs. They therefore leave the host and move to the soil to search for a new host, continuing the life cycle (Chaston et al., 2011; Mahmoud, 2016). The 16 bacterium-nematode complex is highly virulent and kills its hosts quickly (Lacey et al., 2001). The general life cycle of EPNs is illustrated in Figure 1.4 and the advantages and disadvantages associated with the use of EPNs as biocontrol agents are provided in Table 1.3. Figure 1.4: An illustration of the typical life cycle of entomopathogenic nematodes (Hannes Visagie, North-West University, South Africa; photo’s supplied by Antoinette Malan, University of Stellenbosch; illustration cited by Malan and Ferreira, 2017). 17 Table 1.3: Advantages and disadvantages of EPNs used commercially to control insect pests. Advantages References Easily massed produced in vivo and in vitro and formulated as a Lacey et al. (2001) biopesticide. Gözel and Gozel (2016) Exempted from registration in many countries. Gözel and Gozel (2016) Compatible with many pesticides. Kaya and Gaugler (1993) Amendable to genetic selection. Kaya and Gaugler (1993) A broad host range of insect pests. Gözel and Gozel (2016) Able to seek or ambush an insect host and rapidly kill it. Gözel and Gozel (2016) Can be applied with conventional application equipment. Lacey et al. (2001) Gözel and Gozel (2016) Increase biodiversity in managed ecosystems, resulting in increase Lacey et al. (2001) of other natural enemy activities. Not harmful to humans or higher organisms. Lacey et al. (2001) Gözel and Gozel (2016) Pesticide residues in food is reduced. Lacey et al. (2001) Disadvantages References Formulation and quality control is challenging. Gözel and Gozel (2016) Limited shelf life and refrigerated storage required. Gözel and Gozel (2016) Environmental limitations for survival and infestation are factors like Gözel and Gozel (2016) adequate moisture, temperatures, harsh environmental conditions. Has a broad host range. Lacey et al. (2001) 1.4.1. Entomopathogenic nematodes as biological control agents The potential use of EPNs as biopesticides is an environmentally friendly control option. Various EPN species are globally used against a wide range of insect pest species and success has been achieved against insect pests occurring in different habitats (Gözel and Gozel, 2016). The success of EPNs for control of insects varied according to the host species as well as the prevailing environmental conditions where they were applied. Commercially available EPNs applied against insect pest worldwide are listed in Table 1.4. The commercialised EPN species listed in Table 1.4 are those that are currently the most commonly used to successfully control target insect pests. Their wide geographic distribution enables them to be applied where law restriction only permits naturally occurring species to be used for biological control (Půža et al. 2016; Abate et al., 2017). More than a decade ago, Kaya et al. (2006) listed many insect pests controlled by EPNs specifically in Asia and Central and South America, which included species not listed above, such as Steinernema arenarium (Artyukhovsky, 1967) Wouts, MráÏcek, Gerdin and Bedding, 1982, Steinernema bicornutum Tallosi, Peters and Ehlers, 1995, Steinernema longicaudum Shen and Wang, 1992, 18 Steinernema monticolum Stock, Choo and Kaya, 1997 and Steinernema thermophilum Ganguly and Singh, 2000, Steinernema kushidai Mamiya, 1988, Steinernema scarabaei Stock and Koppenhöfer, 2003, Heterorhabditis downesi Stock, Griffin and Brunell, 2002, Heterorhabditis indica Poinar, Karunakar and David, 1992, and Heterorhabditis marelata Liu and Berry, 1996. Heterorhabditis zealandica Poinar, 1990 can also be considered for economic use having effectively controlled insects in bioassays (Gözel and Gozel, 2016). However, EPN’s have the potential to control many more insect pests as proved in laboratory and field studies (Grewal et al., 2001). 19 Table 1.4: Entomopathogenic nematode species commercially available, according to Abate et al. (2017), with their associated symbiotic bacteria, the geographic distribution and a list of insect pests successfully controlled (Gözel and Gozel, 2016). EPN species Symbiotic bacteria Geographic distribution Targeted insect pest (Gözel and Gozel, 2016) Steinernema carpocapsae Xenorhabdus nematophila Worldwide Agrotis ipsilon (Hufnagel) (Lepidoptera: Noctuidae) (Weiser, 1955) Wouts, (Půža et al., 2016) (Půža et al., 2016) Amyelois transitella (Walker) (Lepidoptera: Pyralidae) MráÏcek, Gerdin and Chrysoteuchia topiaria (Zeller) (Lepidoptera: Crambidae) Bedding, 1982 Cosmopolites sordidus (Germar) (Coleoptera: Curculionidae) Ctenocephalides felis (Bouché) (Siphonaptera: Pulicidae) Cylas formicarius (Fabricius) (Coleoptera: Brentidae) Cydia pomonella (Linnaeus) (Lepidoptera: Tortricidae) Diabrotica spp. Chevrolet (Coleoptera: Chrysomelidae) Helicoverpa zea (Boddie) (Lepidoptera: Noctuidae) Hylobius abietis Linnaeus (Coleoptera: Curculionidae) Liriomyza spp. Mik (Diptera: Agromyzidae) Macronoctua onusta Grote (Lepidoptera: Noctuidae) Opogona sacchari (Bojer) (Lepidoptera: Tineidae) Otiorhynchus sulcatus (Fabricius) (Coleoptera: Curculionidae) Platyptilia carduidactyla (Riley) (Lepidoptera: Pterophoridae) Spodoptera spp. Guenée (Lepidoptera: Noctuidae) Scapteriscus spp. (Scudder) (Orthoptera: Gryllotalpidae) Scatella spp Robineau-Desvoidy (Diptera: Ephydridae) Sphenophorus spp. Schönherr (Coleoptera: Curculionidae) Synanthedon spp. Hübner (Lepidoptera: Sesiidae) Tipula spp Linnaeus (Diptera: Tipulidae) Turf and ornamental scarub grubs (Coleoptera: Scarabaeidae) 20 Steinernema feltiae (Filipjev, Xenorhabdus bovienii Worldwide Bradysia spp. Winnertz (Diptera: Sciaridae) 1934) Wouts, MráÏcek, Gerdin (Půža et al., 2016) (Půža et al., 2016) C. sordidus and Bedding, C. pomonella 1982 C. formicarius, Liriomyza spp H. zea Scatella spp Spodoptera spp. Synanthedon spp Steinernema glaseri Steiner, Xenorhabdus poinarii Europe, Russia, USA, C. sordidus 1929 (Půža et al., 2016) Argentina, Azores, O. sulcatus China, Korea, Spain Turf and ornamental scarub grubs (Scarabaeidae), (Půža et al., 2016) Steinernema kraussei (Steiner, Xenorhabdus bovienii Europe, Russia, 1923) Travassos, (Půža et al., 2016) Canada 1927 (Půža et al., 2016) Steinernema riobrave Xenorhabdus cabanillasii North America Spodoptera spp. Aethina tumida Murray (Coleoptera: Cabanillas, Poinar and (Půža et al., 2016) (Půža et al., 2016) Nitidulidae), Conotrachelus nenuphar Herbst (Coleoptera: Raulston, 1994 Curculionidae) Diaprepes abbreviatus (Linnaeus) (Coleoptera: Curculionidae) Pachnaeus spp Schönherr (Coleoptera: Curculionidae) H. zea Scapteriscus spp. 21 Steinernema scapterisci Xenorhabdus innexi South America Scapteriscus spp. Nguyen and Smart, 1992 (Půža et al., 2016) (Hominick et al., 1996) Heterorhabditis bacteriophora Photorhabdus luminescens, Worldwide Bradysia spp. Poinar, 1976 Photorhabdus temperata (Půža et al., 2016) C. formicarius (Půža et al., 2016) Diabrotica spp. D. abbreviatus Macronoctua onusta Grote (Lepidoptera: Noctuidae) O. sacchari O. sulcatus Pachnaeus spp. Sphenophorus spp Synanthedon spp. Vitacea polistiformis (Harris) (Lepidoptera: Sesiidae) Turf and ornamental scarab grubs (Scarabaeidae) Heterorhabditis megidis Photorhabdus temperata North America, Asia, Otiorhynchus ovatus (Linnaeus) (Coleoptera: Curculionidae) Poinar, Jackson and Klein, (Půža et al., 2016) Europe O. sulcatus 1988 (Půža et al., 2016) 22 1.4.2. Entomopathogenic nematodes used for the control of Tuta absoluta Three nematode species indigenous to Spain has been evaluated for control of T. absoluta larvae and pupae under laboratory and greenhouse conditions (Batalla-Carrera et al., 2010). These species were effective in controlling the larvae (78.6-100% mortality), but not the pupae (<10% pupa mortality). Steinernema carpocapsae (Weiser, 1955) Wouts, MráÏcek, Gerdin and Bedding, 1982, Steinernema feltiae (Filipjev, 1934) Wouts, MráÏcek, Gerdin and Bedding, 1982, and Heterorhabditis bacteriophora Poinar, 1976 were also evaluated in laboratory bioassays for the control of T. absoluta larvae, pupae and adults and were highly effective as biocontrol agents of this pest in Spain (Garcia-del-Pino et al., 2013). Soil application of S. carpocapsae, S. feltiae and H. bacteriophora for the control of final instar larvae that enter the soil to pupate, was evaluated by catching emerging adults with first flight in a trapping adhesive for T. absoluta. High mortality of larvae was reported, but no pupal mortality. A high percentage adults emerged where S. carpocapsae was applied, while the emergence rate was very low where S. feltiae was applied. Four species of nematodes native to Turkey, viz. Steinernema affine Bovien, 1937 (isolate 46), S. carpocapsae (isolate 1133), S. feltiae (isolate 879) and H. bacteriophora (isolate 1144) were evaluated for control of T. absoluta larvae in a field in Turkey (Gözel and Kasap, 2015). Steinernema feltiae proved to be the most effective with high levels of control, but the other three species provided little control of T. absoluta larvae in tomato fields (Gözel and Kasap, 2015). 1.4.3. Entomopathogenic nematodes in South Africa The discovery of EPNs in South Africa took place during 1953 (Harington, 1953). Entomopathogenic nematodes were found in all life stages, except in the eggs of the black maize beetle, Heteronychus arator Fabricius (Coleoptera: Scarabidae), near Grahamstown, Eastern Cape Province in a maize field (Harington, 1953). The first survey of EPNs took place 33 years (1988) after the initial discovery in South Africa (Malan and Ferreira, 2017). The susceptibility of crop pests to native EPN species is continuously investigated in South Africa (Malan and Ferreira, 2017). All EPN species isolated, and the insect pest species used for susceptibility testing in South Africa, are provided in Table 1.5. There are currently 19 identified EPN species in South Africa, viz. 13 Steinernema spp. and 7 Heterorhabditis spp. The most recent studies were conducted on grapevine (Platt et al., 2019b), avocado, litchi, macadamia (Steyn et al., 2019) and blueberries (Dlamini et al., 2020). 23 Table 1.5: Entomopathogenic nematodes isolated, associated bacteria and insect pests that has been tested for susceptibility. EPN spp. identified in South Africa Associated bacteria Insect pests (Malan and Ferreira, 2017) Steinernema beitlechemi Çimen, Puža, Nermuť, Hatting, Xenorhabdus khoisanae Ramakuwela, Faktorová and Hazir, 2016 (Çimen, Půža, Nermuť, (Çimen, Půža, Nermuť, Hatting, Ramakuwela, Faktorová et Hatting, Ramakuwela, al., 2016; Abate et al., 2018) Faktorová et al., 2016; Abate et al., 2018) Steinernema bertusi Katumanyane, Malan, Tiedt and Hurley Unknown 2019 (Katumanyane et al., 2019) Steinernema biddulphi Çimen, Půža, Nermuť, Hatting, New Xenorhabdus spp. still Ramakuwela and Hazir, 2016 being identified (Çimen, Půža, Nermuť, Hatting, Ramakuwela and Hazir, (Çimen, Půža, Nermuť, 2016) Hatting, Ramakuwela and Hazir, 2016) Steinernema citrae Stokwe, Malan, Nguyen and Tiedt, 2011 Xenorhabdus bovienii Cydia pomonella (Linnaeus) Codling moth (Lepidoptera: Tortricidae) (Malan et al., 2014; Malan and Ferreira, 2017; Abate et al., (Abate et al., 2018) Phlyctinus callosus (Schönherr) Banded fruit weevil (Coleoptera: 2018) Curculionidae) Planococcus citri (Risso) Citrus mealybug (Hemiptera: Pseudococcidae) Planococcus ficus (Signoret) Vine mealybug (Hemiptera: Pseudococcidae) Thaumatotibia leucotreta (Meyrick) False codling moth (Lepidoptera: Tortricidae) 24 Steinernema fabii Abate, Malan, Tiedt, Wingfield, Slippers Xenorhabdus khoisanae and Hurley, 2016 (Abate et al., 2018) (Abate et al., 2016; Malan and Ferreira, 2017) Steinernema innovationi Çimen, Lee, Hatting, Hazir and Unknown Stock, 2014 (Abate et al., 2018) (Çimen et al., 2015; Malan and Ferreira, 2016; Abate et al., 2018) Steinernema jeffreyense Malan, Knoetze and Tiedt, 2015 Unknown T. leucotreta (Malan et al., 2016a; Malan and Ferreira, 2017) Steinernema khoisanae Nguyen, Malan and Gozel, 2006 Xenorhabdus khoisanae Ceratitis capitata (Wiedemann) Mediterranean fruit fly (Diptera: (Malan et al. 2014) (Malan and Ferreira, 2017) Tephritidae) Ceratitis rosa Karsch Natal fruit fly (Diptera: Tephritidae) C. pomonella P. callosus P. citri P. ficus T. leucotreta Steinernema litchii Steyn, Knoetze, Tiedt and Malan, 2017 Unknown (Steyn, Knoetze et al., 2017) Steinernema nguyeni Malan, Knoetze and Tiedt, 2016 Unknown (Malan et al., 2016b; Malan and Ferreira, 2017) Steinernema sacchari Nthenga, Knoetze, Berry, Tiedt and Xenorhabdus khoisanae Malan, 2014 (Abate et al., 2018) (Nthenga et al., 2014; Malan and Ferreira, 2017; Abate et al., 2018) 25 Steinernema tophus Çimen, Lee, Hatting, Hazir and Stock, Unknown 2014 (Abate et al., 2018) (Çimen et al., 2014; Malan and Ferreira, 2017; Abate et al., 2018) Steinernema yirgalemense Nguyen, Tesfamariam, Gözel, Xenorhabdus indica C. capitata Gaugler and Adams, 2005 (Malan and Ferreira, 2017) C. rosa Cryptolaemus montrouzieri (Mulsant) mealybug ladybird (Malan et al., 2014) (Coleoptera: Coccinellidae) C. pomonella P. callosus P. citri P. ficus Pseudococcus viburni (Signoret) Obscure mealybug (Hemiptera: Pseudococcidae) T. leucotreta Heterorhabditis bacteriophora Poinar, 1976 Photorhabdus luminescens C. capitata (Malan et al., 2014; Malan and Ferreira, 2017; Abate et al., subsp. Laumondii C. rosa 2018) (Abate et al., 2018) C. pomonella P. callosus P. citr P. ficus P. viburni T. leucotreta Heterorhabditis baujardi Phan, Subbotin, Nguyen, Photorhabdus luminescens and Moens, 2003 subsp. luminescens (Steyn, Malan et al., 2017) (Abate et al., 2018) 26 Heterorhabditis indica Poinar, Karunakar and David, 1992 Unknown (James et al., 2018) Heterorhabditis noenieputensis Malan, Knoetze and Tiedt, Photorhabdus C. pomonella 2014 luminescence subsp. P. callosus (Malan et al., 2014; Malan and Ferreira, 2017) noenieputensis P. citri (Malan and Ferreira, 2017) P. ficus Heterorhabditis safricana Malan, Nguyen, de Waal Photorhabdus luminescens P. citri and Tiedt, 2008 subsp. Laumondii (Malan and Ferreira, 2017; Abate et al., 2018; Hatting et al., (Abate et al., 2018) 2019) Heterorhabditis taysaerae Shamseldean, Abou-El-Sooud, Unknown Ab-El-Gawad and Saleh, 1996 (Abate et al., 2018) (Steyn, Malan et al., 2017; Abate et al., 2018) Heterorhabditis zealandica Poinar, 1990 Photorhabdus zealandica C. montrouzieri (Steyn, Malan et al., 2017) (Malan and Ferreira, 2017) C. pomonella P. callosus P. citri P. ficus P. viburni T. leucotreta 27 Heterorhabditis bacteriophora is an EPN commonly isolated from South African soils (Malan and Ferreira, 2017). Steinernema feltiae is an exotic nematode in Africa and has only once being reported as naturally occurring on the African continent, in Algeria (Malan and Ferreira, 2017). In South Africa, only one EPN species, H. bacteriophora, is registered for use and is commercially available as Cryptonem® (Hatting et al., 2019). This product is registered against the False codling moth, Thaumatotibia leucotreta (Meyrick) (Lepidoptera: Tortricidae), Codling moth, Cydia pomonella (Linnaeus) (Lepidoptera: Tortricidae), weevils and gnats (Hatting et al., 2019). No research on the control of T. absoluta with native South African EPN species has been done to date. 1.5. Aim and Objectives The aim of this study was to determine the efficacy of selected EPN species native to South Africa for the control of T. absoluta. The objectives of this study were: 1) To evaluate the efficacy of four native EPN species, viz. Steinernema Jeffreyense Malan, Knoetze and Tiedt, 2015, Steinernema yirgalemense Nguyen, Tesfamariam, Gözel, Gaugler and Adams, 2005, Heterorhabditis baujardi Phan, Subbotin, Nguyen and Moens, 2003 and Heterotrhabditis noenieputensis Malan, Knoetze and Tiedt, 2014 against T. absoluta in South Africa. 2) To evaluate the efficacy of two native EPN species, viz. 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Seasonal changes in morphophysiological traits of two native Patagonian shrubs from Argentina with different drought resistance strategies. Plant Physiology and Biochemistry 127:506-515. VISSER, D. 2015. Tomato, brinjal and peppers. pp 66-73. In: Insects of cultivated plants and natural pastures in Southern Africa. (Eds. Prinsloo, G.L. and Uys, V.M.). Entomological: Society of Southern Africa: Hatfield. VISSER, D., UYS, V.M., NIEUWENHUIS, R.J. and PIETERSE, W. 2017. First records of the tomato leaf miner Tuta absoluta (Meyrick, 1917) (Lepidoptera: Gelechiidae) in South Africa. BioInvasions Records 6(4):301-305. 38 ZAPPALÀ, L., BIONDI, A., ALMA, A., AL-JBOORY, I.J., ARNO`, J., BAYRAM, A., CHAILLEUX, A., EL-ARNAOUTY, A., GERLING, D., GUENAOUI, Y., SHALTIEL-HARPAZ, L., SISCARO, G., STAVRINIDES, M., TAVELLA, L., AZNAR, R.V., URBANEJA, A. and DESNEUX, N. 2013. Natural enemies of the South American moth, Tuta absoluta, in Europe, North Africa and Middle East, and their potential use in pest control strategies. Journal of Pest Science 86:635-647. ZOLFAGHARIAN, M., SAEEDIZADEH, A. and ABBASIPOUR, H. 2016. Efficacy of two entomopathogenic nematode species as potential biocontrol agents against the diamondback moth, Plutella xylostella (L.). Journal of Biological Control 30(2):78-83. 39 Chapter 2: Efficacy of indigenous entomopathogenic nematodes (Rhabditida: Heterorhabditidae and Steinernematidae) against Tuta absoluta (Lepidoptera: Gelechiidae) in laboratory bioassays Abstract Tuta absoluta (Meyrick) (Lepidoptera: Gelechiidae) is a new invasive pest of tomato and other Solanaceous crops in South Africa. Tuta absoluta developed resistance to various insecticide groups in Asia, Europe, South and North America which emphasizes the need for an integrated pest management (IPM) approach to control this pest. Laboratory bioassays were conducted to determine the potential of entomopathogenic nematodes (EPNs) as a possible biological control agent for T. absoluta. The infective juveniles (IJs) of four indigenous nematode species were screened in vitro for their efficacy in killing fourth instar larvae and pupae of T. absoluta. Steinernema Jeffreyense Malan, Knoetze and Tiedt, 2015, Steinernema yirgalemense Nguyen, Tesfamariam, Gözel, Gaugler and Adams, 2005, Heterorhabditis baujardi Phan, Subbotin, Nguyen and Moens, 2003 and Heterotrhabditis noenieputensis Malan, Knoetze and Tiedt, 2014 were all highly effective and killed 100% of the T. absoluta larvae. The EPNs were, however, less effective in killing T. absoluta pupae. The highest mortality was caused by S. yirgalemense (41.7%), with less than 30% mortality caused by S. jeffreyense, H. baujardi and H. noenieputensis. The highest percentage infection was by S. yirgalemense (38%), while infection by all the other species was lower than 20%. The number of IJs that penetrated T. absoluta larvae and pupae was also low. All four EPN species tested have potential for use in an IPM system for to control of T. absoluta larvae. Key words: biocontrol, EPNs, Heterorhabditis spp., laboratory bioassays, native, Steinernema spp., tomato leafminer 2.1. Introduction Biological control is the most important component of an integrated pest management (IPM) program (Hoodle, 2006) and an alternative to chemical control of pests (De Bach and Rosen, 1991). The growing need to develop biological control as an alternative method to control insect pests coincided with the rate at which entomopathogenic nematode (EPN) species was discovered and described (Adams and Nguyen, 2002). Most of the EPN species originally used as biopesticides were described before 1990. One or a few isolates from these species, were initially collected from insects that were naturally parasitized (Bedding, 2006). Little research has been done on EPN species before 1960 when the use of insecticides was affordable and effective (Adams and Nguyen, 2002). The negative environmental effects of pesticides were recognized during 1960 (Adams and Nguyen, 2002). When the use of pesticides became more restricted and costly, and their efficacy decreased gradually, 40 alternative control methods became more attractive (Adams and Nguyen, 2002). Biological control received renewed attention from scientists with research on EPNs that rapidly expanded during the 1980’s. The search for new species that could provide effective control and be commercialised was prioritised (Adams and Nguyen, 2002). The first investigations of the potential of EPNs as biological control agents against insect pests, specifically Steinernema glaseri Steiner, 1929 for control of the Japanese beetle, was done in the early 1930s (Poinar and Grewal, 2012). This EPN species effectively controlled the Japanese beetle in field experiments (Glaser et al., 1935). Although Steinernema kraussei (Steiner, 1923) Travassos, 1927 was the first EPN to be discovered, S. glaseri was the first EPN species investigated for its potential as a biological control agent (Poinar and Grewal, 2012). Heterorhabditis bacteriophora Poinar, 1976 (Heterorhabditidae) was discovered and described in 1975 by Poinar (1975). It has since been used in various virulence testings (Chyzik et al., 1996; Han and Ehlers, 2000; Ehlers et al., 2003; Anbesse et al., 2008, Batalla- Carrera et al., 2010; Van Damme et al., 2016; Kamali et al., 2018). Entomopathogenic nematodes have thus been marketed and already proved to be effective bio-pesticides in the early years of discovery (Gaugler et al., 1997). Steinernema spp., as well as Heterorhabditis spp. are now commercially available (Gözel and Gozel, 2016; Abate et al., 2017) and commercially produced (Georgis et al., 2006, Keshari et al., 2019) for control of various insect pests worldwide. Six Steinernema spp., viz. Steinernema carpocapsae (Weiser, 1955) Wouts, MráÏcek, Gerdin and Bedding, 1982; Steinernema feltiae (Filipjev, 1934) Wouts, MráÏcek, Gerdin and Bedding, 1982; S. glaseri; Steinernema kraussei, Steinernema riobrave Cabanillas, Poinar and Raulston, 1994; Steinernema scapterisci Nguyen and Smart, 1992; and two Heterorhabditis sp., viz. H. bacteriophora and Heterorhabditis megidis Poinar, Jackson and Klein, 1988 are the most commonly produced species. These nematode species are produced in a liquid culture and are successfully applied in the industry. Other species commercially available are Steinernema kushidai Mamiya, 1988; Steinernema scarabaei Stock and Koppenhöfer, 2003; Heterorhabditis indica Poinar, Karunakar and David, 1992; Heterorhabditis marelata Liu and Berry, 1996 and Heterorhabditis downesi Stock, Griffin and Brunell, 2002 (Kaya et al., 2006; Georgis et al., 2006; Lacey et al., 2015; Abate et al., 2017). Entomopathogenic nematodes have a broad host spectrum and can be cultivated in vivo or in vitro on a large scale (Mahmoud, 2016). They do not pose a threat towards any plants, vertebrates and many invertebrates (Boemare et al., 1996). The short life cycle, ability to kill hosts fast and effective mass culturing of EPNs contribute to their status as good potential biological control agents (Kaya and Gaugler, 1993; Griffin, 2012). 41 In the family Steinernematidae, at least 88 species have been identified in the genus Steinernema (Abate et al., 2017) and only one species in the genus Neosteinernema Nguyen and Smart, 1994 (Hunt and Subbotin, 2016). The family Heterorhabditidae has only one genus, namely Heterorhabditis with 19 identified species (Abate et al., 2017). Steinernema carpocapsae and S. feltiae occur worldwide as well as H. indica and H. bacteriophora, with H. bacteriophora being the most widespread EPN in the world (Hominick, 2002). The application of EPNs in South Africa is not exempted from registration unlike most of the European countries, North America or the United Kingdom (Van Zyl and Malan, 2014). South Africa is subjected to regulations by the Department of Agriculture, Forestry and Fisheries (DAFF) for the import of EPNs, in terms of importation of exotic species for use in biological control (Van Zyl and Malan, 2014). This is enforced by the South African Agricultural Pests Act 36 of 1983, which prohibits importation of exotic organisms, including foreign EPNs, without a permit and a full-impact study provided by DAFF under the Agricultural Pest Act (DAFF, 2020; Klein et al., 2011). This is needed since the introduction of exotic EPNs could reduce and possibly displace native nematodes, and it could also have unknown effects on non-target organisms (Malan et al., 2011). It is therefore important to search for local EPNs that could be used for possible biological control. Currently South Africa has 19 identified EPN species consisting of 12 Steinernema spp. and 7 Heterorhabditis spp. The Steinernema spp. are Steinernema beitlechemi Çimen, Puža, Nermuť, Hatting, Ramakuwela, Faktorová and Hazir, 2016 (Çimen, Půža, Nermuť, Hatting, Ramakuwela, Faktorová et al., 2016), Steinernema biddulphi Çimen, Půža, Nermuť, Hatting, Ramakuwela and Hazir, 2016 (Çimen, Půža, Nermuť, Hatting, Ramakuwela and Hazir, 2016), Steinernema citrae Stokwe, Malan, Nguyen and Tiedt, 2011 (Malan et al., 2014), Steinernema fabii Abate, Malan, Tiedt , Wingfield, Slippers and Hurley, 2016 (Abate et al., 2016), Steinernema innovationi Çimen, Lee, Hatting, Hazir and Stock, 2014 (Çimen et al., 2015), Steinernema jeffreyense Malan, Knoetze and Tiedt, 2015 (Malan et al., 2016a), Steinernema khoisanae Nguyen, Malan and Gozel, 2006 (Malan et al., 2014), Steinernema litchii Steyn, Knoetze, Tiedt and Malan, 2017 (Steyn, Knoetze et al., 2017), Steinernema nguyeni Malan Knoetze and Tiedt, 2016 (Malan et al., 2016b), Steinernema sacchari Nthenga, Knoetze, Berry, Tiedt and Malan, 2014 (Nthenga et al., 2014), Steinernema tophus Çimen, Lee, Hatting, Hazir and Stock, 2014 (Çimen et al., 2014), Steinernema yirgalemense Nguyen, Tesfamariam, Gozel, Gaugler and Adams, 2005 (Malan et al., 2014). The Heterorhabditis spp. include H. bacteriophora (Malan et al., 2014), Heterorhabditis baujardi Phan, Subbotin, Nguyen and Moens, 2003 (Steyn, Malan et al., 2017), H. indica (James et al., 2018), Heterorhabditis noenieputensis Malan, Knoetze and Tiedt, 2014 (Malan et al., 2014), Heterorhabditis safricana Malan, Nguyen, de Waal and Tiedt, 2008 (Hatting et al., 2018), 42 Heterorhabditis taysaerae Shamseldean, Abou-El-Sooud, Ab-El-Gawad and Saleh, 1996 (Steyn, Malan et al., 2017) and Heterorhabditis zealandica Poinar, 1990 (Steyn, Malan et al., 2017). Tuta absoluta (Meyrick) (Lepidoptera: Gelechiidae) is an invasive insect pest which poses a threat to Solanaceous crop production in South Africa since 2016 (Visser et al., 2017). This pest has also been reported to be resistant to insecticides from groups that have various modes of action from regions in Europe and South America (Guedes et al., 2019). It caused increased tomato production costs and crop losses, in newly invaded areas (Sannino and Espinosa, 2010). It is therefore important to use alternative control methods for effective control of this pest and also to reduce selection pressure for insecticide resistant development (Roditakis et al., 2018). Biological agents can be considered as an alternative control method for T. absoluta (Desneux et al., 2010). Entomopathogens such as fungi, bacteria and nematodes are used to control a variety of insect pests (Lacey et al., 2001). Studies on the efficacy of EPNs against insect pests have been conducted with promising results, but it is yet to be implemented for commercial use in South Africa. These include studies conducted in South Africa with indigenous EPNs against key lepidopteran pests, such as Cydia pomonella (Linnaeus) (Lepidoptera: Tortricidae) (De Waal et al., 2011), Thaumatotibia leucotreta (Meyrick) (Lepidoptera: Tortricidae) (Steyn, Malan et al., 2017), Helicoverpa armigera (Lepidoptera: Noctuidae) (Jankielsohn and Hatting, 2005) and Busseola fusca (Lepidoptera: Noctuidae) (Ramakuwela et al., 2011). A more recent study conducted on Phlyctinus callosus (Schönherr) (Coleoptera: Curculionidae), the banded fruit weevil also illustrated that various EPN species have the potential to control this pest, with S. yirgalemense and H. indica that provided effective control. These species were also highly effective in controlling the stages of the banded fruit weevil that occurred in the soil in field experiments (Dlamini et al., 2019). Planococcus ficus (Signoret) (Hemiptera: Pseudococcidae) the vine mealybug, another key grapevine pest in South Africa, was also found to be successfully controlled by S. yirgalemense, which is highly pathogenic to the adult females (Le Vieux and Malan, 2013). Since one EPN species is pathogenic to more than one insect species, only one EPN species could therefore be applied in a field to target more than one important pest (Dlamini et al., 2019). Only one EPN species, H. bacteriophora, is currently registered for use in South Africa. It is available as Cryptonem® for control of the false codling moth, codling moth, weevils and gnats (Hatting et al., 2018). According to the above-mentioned successful trial results, potential for more EPN species to be registered for control of various insect species in South Africa exists. 43 The objective of this study was to evaluate the efficacy of four native EPN species, viz. S. jeffreyense, S. yirgalemense, H. baujardi and H. noenieputensis against T. absoluta in South Africa. 2.2. Material and methods Tuta absoluta stock colony A rearing colony was established with T. absoluta larvae collected from commercially produced tomatoes at Tarlton (26º03’29.07’’S 27º40’44.00’’E; Gauteng Province), Ventersdorp (26º24’24.07’’S 26º47’36.00’’E; North-West Province) and Polokwane (23°46’48.03”S 29°28’00.02”E; Limpopo Province). The larvae were released onto potted tomato plants (cvs Money maker and Monica) that were placed in a 4 x 3 m netted enclosure at Potchefstroom (26°40'26.65"S 27°06'23.23"E; North-West Province) under environmental conditions. Uninfested plants were regularly introduced into the cage serving as food for the larvae. Larvae from this stock colony were used in all bioassays. Entomopathogenic nematodes Infective juveniles (IJs) of four EPN species, viz. S. jeffreyense (J194), S. yirgalemense (157- C), H. baujardi (MT17) and H. noenieputensis (SF669) were obtained from the Department of Conservation Ecology and Entomology, Stellenbosch University, Western Cape Province, South Africa. The nematodes were kept in vented culture flasks in 150 ml filtered water, in a dark incubator at 15±1 °C and the flasks were shaken weekly to improve aeration and nematode survival (Platt et al., 2019). Bioassay protocol Six milliliters of the base EPN solution, containing IJs, reared in vivo for inoculation purposes were pipetted into a 50 ml capacity plastic tube containing two small magnetic stirrers. The tube was nested upright into a 200 ml capacity glass beaker, which was placed onto a magnetic stirrer. The content was stirred at 2 000 rpm for 2 min. Eight, 10 µL drops were then removed from the stirring nematode solution using a micropipette. Each drop was placed individually on a De Grisse nematode counting dish (De Grisse, 1963). The number of IJs in each drop was counted. This was repeated six times (8 drops of 10 µL each x 6 replicates) and the mean number of IJs was hence determined per 10 µL drop (number of IJs counted in the 10 x 8 µL drops / 8). This enabled calculation of the volume to be extracted, from the base solution, to obtain the required number of IJs (100) for inoculation on filter paper discs for the respective experiments. The following equation was used to determine how many microliters were needed to obtain 100 IJs. Firstly, the number of IJs present in the 6 mL (6 000 µL extracted from the base solution were calculated as follows: 44 Volume extracted from base solution (6 000 µL) in which IJs was suspendedin = x mean number of IJs in 8 individual 10 µL drops (6 replicates) volume of individual drops (10 µL) IJs were suspended in for counting = number of IJs in base suspension in (6 000 µL). The next step entailed the calculation of the volume to be extracted from the 6 000 µL base solution to obtain 100 IJs for inoculation purposes, which was done as follows: number of IJs needed (namely 100IJ) = x volume of base solution, namely 6 000 µL total number of IJs suspended in the base suspension of 6 000µL = volume (µL) needed to obtain 100IJs Every second well of a 24 multi-well plate (Cellstar® Cat.-No. 662 102) was lined with a circular filter paper disc (13 mm diameter). The IJs were inoculated at a rate of 100 IJs/50 µL water into each of these wells. Leaf disks (10 mm diameter) were cut from tomato leaves and inoculated with one, fourth instar T. absoluta larva per disk. One of these leaf disks containing a larva that had already mined into the disk, was transferred to each of the EPN inoculated wells (Fig. 2.1A). Two-day old pupae were also collected from the stock colony and bioassays were conducted by placing one T. absoluta pupa on a paper disk in every second well (Fig 2.1B). For the respective experiments with either larvae or pupae, each plate, containing 12 individuals, served as a replicate and there were five replicates per experiment. The bioassay plates were sealed with lids, placed into plastic containers lined with moist paper towels and closed with a lid to maintain high humidity. The treatment and control multi-well plates were kept in separate containers. These containers were kept at 25 ± 1 °C in an incubator in darkness. Mortality of T. absoluta larvae or pupae (depending on the experiment) was determined by means of gentle prodding with a thin, soft brush 72 h after inoculation. Larvae and pupae from the respective experiments were frozen after 72 h and were later dissected. The number of nematodes inside each individual was counted to determine the penetration success. 45 A B A Figure 2.1: Multi-well plates containing, (A) filter paper disc and a leaf disc with one Tuta absoluta larva and (B) pupae placed on filter paper discs. Filter paper discs were inoculated with 100 infective juveniles (IJs) of entomopathogenic nematode species per 50 µL stock solution before insect larvae or pupae were transferred (Photo’s: Odette Coleman, NWU, Potchefstroom). Data analysis Larval and pupal mortality and infection caused by the IJs of the respective EPN species were compared against the control treatment by means of binomial distribution tests. Each experiment for each EPN species used, larvae and pupae, were repeated once. Data from both experiments were combined and one-way ANOVAs were used to compare the number of IJs of the respective EPN species that penetrated T. absoluta larvae and pupae, respectively. Means were separated by Tukey’s HSD test (P<0.05). All analyses were done with Statistica Version 13.3 (TIBCO Software Inc., 2017). 2.3. Results Infective juvenile efficacy of four entomopathogenic nematode species against Tuta absoluta larvae and pupae Infected juveniles of all four EPN species, viz. S. jeffreyense S. yirgalemense, H. baujardi and H. noenieputensis successfully penetrated fourth instar T. absoluta larvae (Table 2.1). All T. absoluta larvae died after infestation with the respective EPN species in both bioassays conducted. Mortality of T. absoluta larvae in the control treatments of all bioassays was 13% or less, except in one bioassay with S. jeffreyense where 20% of larvae in the control treatment died (Table 2.1). Larval mortality caused by all four EPN species differed significantly from larval mortality in the respective control treatments (Table 2.1). 46 Table 2.1: Mean percentage mortality and infection of Tuta absoluta larvae with infective juveniles of Steinernema jeffreyense, Steinernema yirgalemense, Heterorhabditis baujardi and Heterorhabditis noenieputensis, respectively, after 72 hours exposure. Bioassay 1 Bioassay 2 Parameter (± Treatments Treatments SE) P- P-value Steinernema Control value Steinernema Control jeffreyense (water) jeffreyense (water) Mortality (%) 100 11.67±6.24 <0.001 100 20.00±6.24 <0.001 Infected T. absoluta larvae 100 0 <0.001 100 0 <0.001 (%) Steinernema Control Steinernema Control yirgalemense (water) yirgalemense (water) Mortality (%) 100 0 <0.001 100 11.67±3.33 <0.001 Infected T. absoluta larvae 100 0 <0.001 96.67±3.33 0 <0.001 (%) Heterorhabditis Control Heterorhabditis Control baujardi (water) baujardi (water) Mortality (%) 100 8.33±2.64 <0.001 100 10.00±6.24 <0.001 Infected T. absoluta larvae 100 0 <0.001 100 0 <0.001 (%) Heterorhabditis Control Heterorhabditis Control noenieputensis (water) noenieputensis (water) Mortality (%) 100 13.00±5.00 <0.001 100 11.67±6.24 <0.001 Infected T. absoluta larva 98.33±1.67 0 <0.001 100 0 <0.001 (%) P-value determined with the binomial distribution test; Standard Error (SE). Although a significantly higher percentage pupae was infected by S. jeffreyense IJs in the first bioassy compared to the pupae not exposed to the EPN in the control (P<0.001), only 23% was infected. There was, however, no significant difference in the percentage pupae infected between the treated and control pupae that were not exposed to any IJs (P>0.05) (Table 2.2). Percentage T. absoluta pupae penetrated by S. yirgalemense was low (38% and 8% in the two bioassays, respectively), despite significantly more T. absoluta pupae penetrated by IJs compared to untreated control pupae (P<0.001 and P<0.02). The percentage penetration of T. absoluta larvae with H. baujardi and H. noenieputensis differed significantly from the 47 uninfected pupae in the respective control treatments of one of the two bioassays. The percentage penetration was, however, below 20% in both these bioassays (Table 2.2). Table 2.2: Mean percentage mortality and infection of Tuta absoluta pupae with infective juveniles of Steinernema jeffreyense, Steinernema yirgalemense, Heterorhabditis baujardi and Heterorhabditis noenieputensis, respectively, after 72 hours exposure. Bioassay 1 Bioassay 2 Parameter Treatments Treatments (±SE) P- P- Steinernema Control value Steinernema Control value jeffreyense (water) jeffreyense (water) Mortality (%) 23.33±6.12 8.33±2.64 0.027 6.67±3.12 3.33±2.04 0.402 Infected Tuta absoluta 36.67±8.96 0 <0.001 1.67±1.67 0 0.315 pupae (%) Steinernema Control Steinernema Control yirgalemense (water) yirgalemense (water) Mortality (%) 41.67±3.73 11.67±2.04 0.002 18.33±4.86 3.33±2.04 0.008 Infected Tuta absoluta 38.33±4.25 0 <0.001 8.33±4.56 0 0.022 pupae (%) Heterorhabditis Control Heterorhabditis Control baujardi (water) baujardi (water) Mortality (%) 16.67±3.73 13.33±4.25 0.540 10±4.08 6.67±1.67 0.509 Infected Tuta absoluta 13.33±3.73 0 0.004 1.67±1.67 0 0.315 pupae (%) Heterorhabditis Control Heterorhabditis Control noenieputensis (water) noenieputensis (water) Mortality (%) 28.33±3.33 5.00±3.33 0.006 16.67±2.64 15.00±6.12 0.803 Infected Tuta absoluta 18.33±5.53 0 0.005 0 0 1.000 pupae (%) P-value determined with the binomial distribution test. The number of IJs that penetrated T. absoluta larvae or pupae in both bioassays conducted with the respective EPN species, was combined and is provided as the mean number IJ per T. absoluta larva or pupa per EPN species (Table 2.3). There were significant differences in the number of IJs that penetrated larvae of the respective species (F3, 476 = 31.62), but not in the number that penetrated pupae of the respective species (F3,47 = 0.72). Significantly fewer S. yirgalemense IJs penetrated T. absoluta larvae compared to all three other species. The number of H. noenieputensis IJs that penetrated T. absoluta was also significantly lower 48 compared to S. jeffreyense, but did not differ significantly from H. baujardi (Table 2.3). Heterorhabditis baujardi and S. jeffreyense had the highest number of IJs that penetrated T. absoluta larvae. The number of IJs that penetrated T. absoluta pupae, were however low, with no significant difference in the mean number of IJs among the four species (Table 2.3). Table 2.3: Mean number of Steinernema jeffreyense, Steinernema yirgalemense, Heterorhabditis baujardi and Heterorhabditis noenieputensis infective juveniles that penetrated Tuta absoluta larvae and pupae. EPN species Mean IJ/larva Mean IJ/pupa Steinernema jeffreyense 15.44±1.05c 6.50±3.50a Steinernema yirgalemense 6.71±0.53a 6.45±0.88a Heterorhabditis baujardi 12.00±0.63bc 5.00±0.93a Heterorhabditis noenieputensis 9.84±0.60b 4.55±0.87a Means within a column followed by the same lowercase letter are not significantly different at P<0.05 (Tukey). 2.4. Discussion The potential for control of T. absoluta with EPNs has been investigated extensively in Europe (Batalla-Carrera et al., 2010; Garcia-del-Pino et al., 2013; Van Damme et al., 2016). The present study was, however, according to our knowledge, the first one conducted in South Africa where native EPN species was evaluated for control of T. absoluta larvae and pupae. All four EPN species evaluated, viz. S. jeffreyense, S. yirgalemense, H. baujardi and H. noenieputensis were effective, with 100% control of fourth instar T. absoluta larvae under controlled laboratory conditions. Although fewer S. yirgalemense IJs infected fourth instar T. absoluta larvae compared to the other three species, 100% mortality was still achieved. Similar results with high levels of mortality at a dosage rate of 25 IJs/cm2 caused by IJs of S. feltiae (100%), S. carpocapsae (85.7%) and H. bacteriophora (78.6%) were reported by Batalla- Carrera et al. (2010) in Spain. The potential of S. feltiae, S. carpocapsae and H. bacteriophora to control T. absoluta larvae inside tomato leaf mines, was also confirmed by Van Damme et al. (2016) under laboratory conditions. High efficacy of H. bacteriophora and S. carpocapsae for control of T. absoluta larvae was recently also reported by Kamali et al. (2018) in Iran, with H. bacteriophora that caused 99.1 ± 0.03% mortality of last instar T. absoluta larvae at 20 and 50 IJs/cm2. The mean number of IJs that penetrated fourth instar T. absoluta larvae in the current study was below 20 although inoculation was done with 100 IJs per larva or pupa. Invasion of larvae does, however, decrease with a decrease in insect size as well as with an increase in 49 nematode species size (Bastidas et al., 2014). Smaller EPN species control smaller insect species more effectively compared to bigger EPNs (Steyn et al., 2019). Size of the EPN species applied to small insect pests can therefore jeopardize good control and persistence of EPNs (Bastidas et al., 2014; Steyn et al., 2019). Steyn et al. (2019) determined the efficacy of seven EPN species collected in South Africa, viz. H. bacteriophora (SF351), H. baujardi (MT19), H. indica (SGS), H. zealandica (MJ2C), H. noenieputensis (SF669), S. jeffreyense (J194) and S. yirgalemense (157C) against Holocacista capensis Van Nieukerken and Geertsema (Lepidoptera: Heliozelidae). High larval mortality (>83%) was recorded for H. indica, H. noenieputensis and H. baujardi which was ascribed to their foraging ability to locate and penetrate leaf mining galleries (Steyn et al., 2019). He found a low percentage larval mortality (26% - 50%) for the other species evaluated. The T. absoluta pupal infection rate by S. jeffreyense, H. baujardi and H. noenieputensis in the first bioassay of this study, was higher than what was reported by Batalla-Carrera et al. (2010) for S. feltiae, S. carpocapsae and H. bacteriophora, while results from the second experiment corresponded with results from these authors. The EPN species in the study of Batalla-Carrera et al. (2010) as well as those in the present study, were not effective in controlling the pupae. The only EPN species found to significantly infect and kill T. absoluta pupae, when compared to pupae of the untreated control, was S. yirgalemense. The number of S. yirgalemense IJs that infected T. absoluta pupae were, however, similar to IJ infection numbers of S. jeffreyense, H. baujardi and H. noenieputensis. Despite this finding, and according to the low infection rate of all EPN species evaluated in this study, none will be able to successfully control T. absoluta pupae. Pupae from different insect species can vary in susceptibility in terms of penetration by IJs of EPN species. Henneberry et al. (1995) ascribed the inability of EPNs to infect pink bollworm, Pectinophora gossypiella Saunders (Lepidoptera: Gelechiidae) pupae, to a lack of punctures in the integument. Yan et al. (2019) tested 15 EPN isolates and reported the same tendency with less penetration and lower efficacy of EPNs against Spodoptera litura Fabricius (Lepidoptera: Noctuidae) pupae, and as a result, lower mortality, compared to efficacy of control of the larval stage. A possible reason for the low infection rate of T. absoluta pupae in this study may also be a lack of entry route since two-day-old T. absoluta pupae of which the cuticle has already hardened, were used in this study. The age of pupae affects their susceptibility, but it may vary between species (Kaya and Hara, 1981). Almost no penetration by EPNs was reported for three-day old, and older, S. litura pupae, while the pre-pupa and soft one-day-old pupae were infected when exposed to IJs (Kondo and Ishibashi, 1986). Should pupae survive after unsuccessful penetration by EPNs, it may still hold benefits in 50 terms of control, since Kondo and Ishibashi (1986) reported that moths from such pupae were infected upon emergence and were killed a short while afterwards. Foraging behaviours had no definitive effect in this study. Steinernema jeffreyense is an ambusher (Dlamini et al., 2020) that wait for hosts (Campbell and Gaugler, 1997). Ambush foragers scan for cues while they wait and are more effective in finding resources with higher mobility (Lewis et al., 2006). It is therefore doubtful that S. jeffreyense IJs will be successful in finding and infecting T. absoluta pupae. The low T. absoluta pupal infestation by S. yirgalemense recorded in this study could, however, not be explained by its foraging behaviour as an intermediate forager. Intermediate foragers act as both ambushers and as cruisers (Dlamini et al., 2020). It is therefore expected that the cruiser behaviour of S. yirgalemense should assist this species in its ability to find host pupae also. Repelling cues of a host could also inhibit further penetration and prevent over population of resources, as demonstrated with Steinernema spp. (S. carpocapae, S. riobrave and S. feltiae) by Glazer (1997). More nematodes invade susceptible hosts in which they can develop into the adult stage, reproduce and produce offspring (Bastidas et al., 2014). A suitable host does not only provide food, but also a safe environment for mating and where competition can be avoided (Lewis et al., 2006). A small host is a limited food source and can therefore sustain a lower number of individuals in contrast to larger hosts that can sustain a higher number of nematodes (Bastidas et al., 2014). The competition in smaller hosts is higher and a limited number of individuals could enter these hosts. 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African Entomology 22(2):235-249. VISSER, D., UYS, V.M., NIEUWENHUIS, R.J. and PIETERSE, W. 2017. First records of the tomato leaf miner Tuta absoluta (Meyrick, 1917) (Lepidoptera: Gelechiidae) in South Africa. BioInvasions Records 6(4):301-305. YAN, X., ARAIN, M.S., LIN, Y., GU, X., ZHANG, L., LI, J. and HAN, R. 2019. Efficacy of entomopathogenic nematodes against the tobacco cutworm, Spodoptera litura (Lepidoptera: Noctuidae). Journal of Economic Entomology 113(1):64-72. 58 Chapter 3: Efficacy of Steinernema yirgalemense and Steinernema jeffreyense applied as foliar applications for control of Tuta absoluta on tomato under greenhouse conditions in South Africa Abstract Entomopathogenic nematodes (EPNs) were mainly used as biological control agents of soil- inhabiting insect pests, but interest to use it in above-ground applications increased with restrictions on the use of insecticides and increased availability of products containing EPNs. Greenhouse bioassays were conducted to determine the efficacy of two indigenous nematode species, Steinernema Jeffreyense Malan, Knoetze and Tiedt, 2015 and Steinernema yirgalemense Nguyen, Tesfamariam, Gözel, Gaugler and Adams, 2005 against third to fourth instar Tuta absoluta (Meyrick) (Lepidoptera: Gelechiidae) larvae on tomato seedlings. The nematodes were applied at four concentrations, viz. 250, 500, 1 000 and 2 000 IJs mL-1 distilled water to T. absoluta infested tomato seedlings. Leaves with T. absoluta larvae were collected 24 h after application of the nematodes and placed in petri dishes on moist filter paper, and kept in a growth chamber at 25±1 oC. Mortality of T. absoluta larvae were recorded after a further 48 h. The larvae were dissected and the number of IJs that penetrated each larva was recorded. Despite a low number of IJs of both species found in the host larvae, high mortality rates of more than 80% were achieved at high application rates of 1 000 and 2 000 IJs mL-1 water. Steinernema jeffreyense and S. yirgalemense are therefore promising biocontrol agents of T. absoluta larvae infesting tomato cultivated under greenhouse conditions in South Africa. Key words: EPNs, foliar application, Steinernema yirgalemense, Steinernema, jeffreyense, greenhouse, tomato leafminer. 3.1. Introduction Entomopathogenic nematodes (EPNs) are generally applied as an inundative biological control strategy (Beck et al., 2013). The early uses of EPNs as biological control agents were mainly aimed to control soil-inhabiting insect pests, primarily in high valued crops (Kaya et al., 2006). The interest was partially due to a lack of other means to effectively control soil- inhabiting pests in an environmentally friendly manner (Klein, 1990). The application of EPNs to foliage is limited due to the sensitivity of these nematodes to diverse biotic and abiotic factors (Smits, 1996; Noosidum et al., 2016). The main limiting environmental factors are insufficient moisture (desiccation), exposure to extreme temperatures and ultraviolet radiation (Kaya and Gaugler, 1993; Laznik and Trdan, 2012). Entomopathogenic nematodes are soil inhabiting organisms and are therefore protected from adverse environmental conditions (Kaya and Gaugler, 1993). Above-ground applications expose EPNs to environmental 59 conditions, which reduces their efficacy (Kaya and Gaugler, 1993). Most nematode species do not infect hosts when temperatures exceed 32 C (Grewal, 2002), and infective juveniles (IJs) exposed to dry conditions desiccate and die quickly (Moore, 1965). The tolerance levels of IJs are therefore taken to the extremes with above-ground application and desiccation is imminent during warm days. Infective juveniles can survive on leaf surfaces during the evening for up to 12 h (Moore, 1965). Application of EPNs should therefore be done from dusk to evenings (Lello et al., 1996), or during dawn and cloudy days (Smith-Fiola et al., 1996). Restrictions on the use of insecticides and increased availability of products containing EPNs renewed the interest in use in above-ground applications (Arthurs et al., 2004). Protection from environmental conditions is provided to IJs by cryptic habitats. Entomopathogenic nematodes have therefore been tested in these habitats, which include holes and galleries in stems or wood, leaf mines, curled leaves, reproductive plant structures and on leaf surfaces (Arthurs et al., 2004). For EPNs to be effective biopesticides, application should enable the IJs to find and infect a host (Shapiro-Ilan et al., 2006). The application of nematodes in both glasshouses and fields can be done by means of conventional liquid application (Grewal, 2002). An aqueous nematode mix can be applied with general agrochemical or horticultural ground application equipment including hand-held pressurized sprayers, mist blowers, electrostatic or spinning disc systems, aircraft-mounted atomizer sprayers and irrigation systems such as tickle, centre- pivot or furrow irrigation (Georgis, 1990; Arthurs et al., 2004). The same methods are used for soil as well as foliar applications (Grewal, 2002). A constraint for application of liquid formulations of nematodes is sedimentation in the sprayer tanks (Grewal, 2002; Schroer et al., 2005; Beck et al., 2013). Conventional hydraulic nozzles, viz. standard fan and full cone provide a wide range of droplet sizes. These droplets are, however, too small to carry IJs (Lello et al., 1996). Higher output hydraulic nozzles deposit nematodes onto foliar parts more effectively and, up to 98% mortality of Plutella xylostella (Linnaues) (Lepidoptera: Plutellidae) on Chinese cabbage was achieved when using these nozzles for application (Lello et al., 1996). With the spraying of nematodes, droplets can bounce or roll from leaves. The larger and coarser the droplets, the higher the risk becomes for bouncing and rolling from leaves (Crease et al.,1991), which increases the loss of EPNs. The size of the EPN species determines if the droplet should be larger or whether it can be smaller (Beck et al., 2013). Surfactants can alter the surface tension of the droplet preventing spray droplets from bouncing and rolling of leaf surfaces which can increase the deposit of EPNs (Beck et al., 2013). The addition of an adjuvant increases the deposit of IJs on foliage, but can also have 60 an anti-desiccation action (Mason et al., 1998; Laznik and Trdan, 2012). Nematode application at a pressure lower than 2070 kPa is recommended, since nematodes can be damaged with high pressure and extensive recycling through pumping systems (Grewal, 2002). Hydraulic pumps that develop high internal pressure, can shred nematodes. Most pumping systems do, however, make use of membrane or roller pumps and high pressure or force that could damage the IJs, is not generated by these pumps (Grewal, 2002). Oxygen deprivation in tanks and hoses can also inactivate nematodes. It is aggravated when equipment is exposed to direct sunlight which increases the temperature and oxygen demand (Grewal, 2002). Application of nematodes should be avoided when the temperature inside tanks, hoses or nozzles of application equipment, exceeds 30 C (Grewal, 2002). Cool temperatures reduce activity and infectivity while warmer temperatures reduce survival (Grewal, 2002). The equipment used should always be adjusted to enhance motility, survival and deliverance of IJs during spray applications (Brusselman et al., 2012). Steinernema jeffreyense Nguyen, Tesfamariam, Gozel, Gaugler and Adams, 2005 (J194) and Steinernema yirgalemense Malan, Knoetze and Tiedt, 2015 (157-C) (Rhabditida: Steinernematidae) were highly effective in controlling T. absoluta larvae in laboratory bioassays (Chapter 2). Both these species were readily available from the Department of Conservation Ecology and Entomology, Stellenbosch University and were therefore used in this greenhouse study. The objective of this study was to evaluate the efficacy of S. jeffreyense and S. yirgalemense applied as foliar sprays for the control of T. absoluta larvae in tomato plants under controlled greenhouse conditions. 3.2. Materials and Methods Entomopathogenic nematodes Steinernema jeffreyense and S. yirgalemense IJs were obtained from the Department of Conservation Ecology and Entomology, Stellenbosch University, Western Cape Province, South Africa to test their efficacy against third and fourth instar T. absoluta larvae in tomato plants, in a greenhouse trial. The nematodes were kept in vented culture flasks in 150 ml filtered water, in a dark incubator at 15 ± 1 °C. The flasks were shaken weekly to improve aeration and nematode survival following Platt et al. (2019). Bioassay protocol Two 4-week-old tomato seedlings, cv. Monica were planted in 4-L capacity pots in sandy loam soil consisting of percentages of sand:silt:clay = 90.8:2.2:7.0, pH = 6.83 and organic matter = 1.78 % by weight. Third and fourth instar T. absoluta larvae were inoculated onto leaves of seedlings. Each seedling was inoculated with five larvae that tunneled into the leaves and 61 developed well-formed tunnels. The infested plants were kept in a greenhouse with temperature and relative humidity (RH) set at 24 ± 0.5 °C and 80 ± 2%, respectively. Temperature and RH were recorded at 60-min intervals using iButtons® from ColdChain Thermo Dynamics (Fairbridge Technologies). Only pots containing seedlings with five larvae that tunnelled into the leaves and formed tunnels (10 larvae per pot) were used. Seedlings were sprayed with water prior to the application of nematode treatments. The treatments consisted of S. jeffreyense and S. yirgalemense IJs applied at four concentrations, viz. 250, 500, 1 000, and 2 000 IJs mL-1 distilled water. Calculation of the respective concentrations was done according to the method described in chapter 2 (See bioassay protocol).For application of the respective nematodes at each concentration, a 200 mL spray mixture was prepared, which consisted of the EPNs in distilled water and 0.05 mL Nu-Film-P adjuvant (Miller Chemical & Fertilizer CO, Hygrotech). There were eight replicates (pots) for each treatment per EPN species. The untreated control also consisted of eight pots sprayed with a water and Nu-Film adjuvant mixture only. The respective treatments were applied, one day after T. absoluta inoculation, with a spray bottle onto the leaves of the potted tomato seedlings until run off. Leaves with T. absoluta larvae were collected 24 h after application of the nematodes and placed in petri dishes on moist filter paper. The petri dishes were kept in a growth chamber at 25±1 oC for a further 48 h where after the larvae were lightly probed to elicit a response and noted as dead when no movement was detected. The T. absoluta larvae were dissected to confirm infection by nematodes and the number of IJs that penetrated each larvae was recorded. The experiment was repeated twice. Data analysis Results from each experiment conducted with S. jeffreyense and S. yirgalemense on different test dates, were analysed separately. Abbott’s formula was used to correct the data for control mortality (Abbott, 1925). One way ANOVAs were used to compare the corrected percentage mortality, followed by Tukey’s HSD test (P<0.05). The mean number of S. jeffreyense and S. yirgalemense IJs per T. absoluta larva after exposure to the two nematode species at different concentrations, were tested for normality (Shapiro-Wilk test) and homogeneity of variance (Levene’s test). These assumptions were not met. The respective data sets were therefore analysed by means of Kruskal–Wallis test followed by Dunn’s multiple comparison post hoc test. All statistical analyses were performed using TIBCO Statistica™ 13.3 (TIBCO Software, Inc., 2017). 62 3.3. Results Mortality of T. absoluta larvae was high in the control treatment of one experiment where S. jeffreyense was applied. This experiment was, therefore omitted from further analyses. The corrected percentage mortality for T. absoluta larvae exposed to S. yirgalemense varied from 52% to 88% (Exp. 1) and 59 to 87% (Exp. 2) at the respective dosage rates applied and differed significantly between application rates (F3,28 = 6.75; P<0.01; Exp. 1) and (F3,28 = 4.30; P<0.05; Exp. 2). Significantly more T. absoluta larvae died at an application rate of 1 000 S. yirgalemense IJs mL-1 when compared to the other application rates in experiment 1 (Fig. 3.1). Mortality recorded after IJ application at 250, 500 and 2 000 IJs mL-1, did, however, not differ significantly. However, in experiment 2, significantly more larvae were killed by S. yirgalemense at an application rate of 2 000 IJs mL-1 compared to application rates of 250 and 500 IJs mL-1. Control of T. absoluta larvae at application rates of 1 000 and 2 000 IJs mL-1 did not differ significantly. 100 b B 90 Exp. 1 80 Exp. 2 AB 70 a A a A 60 a 50 40 30 20 10 0 250 500 1°000 2°000 Concentration (IJs mL⁻¹) Figure 3.1: Mean corrected percentage mortality ± SE of Tuta absoluta larvae on tomato seedlings caused by Steinernema yirgalemense applied at concentrations of 250 IJs, 500 IJs, 1 000 IJs and 2 000 IJs mL-1 in a (water + 0.05% Nu-Film-P) spray solution. Bars capped with the same lower or upper case letters are not significantly different at P<0.05 (Tukey). The dosage rate at which S. jeffreyense was applied to tomato foliage had a significant effect on the corrected percentage mortality of T. absoluta larvae in galleries (F3,28 = 6.16; P<0.01). The highest mortality was recorded with application rates of 1 000 and 2 000 IJs mL-1 (Fig. 3.2). Corrected percentage mortality ± SE 63 100 b 90 ab 80 70 a a 60 50 40 30 20 10 0 250 500 1°000 2°000 Concentration (IJs mL⁻¹) Figure 3.2: Mean corrected percentage mortality ± SE of Tuta absoluta larvae on tomato seedlings caused by Steinernema jeffreyense applied at concentrations of 250 IJs, 500 IJs, 1 000 IJs and 2 000 IJs mL-1 in a (water + 0.05% Nu-Film-P) spray solution. Bars capped with the same letter are not significantly different at P<0.05 (Tukey). Mean number of IJs that infected T. absoluta larvae was less than three per larva (Table 3.1). Significantly fewer S. yirgalemense IJs, applied at 500 IJs mL-1, penetrated T. absoluta larvae compared to the number that penetrated after application at 2 000 IJs mL-1((H (3, N = 186) = 18.47; P < 0.05). The number of IJs that penetrated after application at concentrations of 250, 1 000 and 2 000 IJs mL-1 did not differ significantly. The number of S. jeffreyense IJs that penetrated T. absoluta larvae did not differ significantly regardless of the application rates of 250, 500, 1 000 and 2 000 IJs mL-1 ((H (3, N = 221) = 9.24; P = 0.06). Table 3.1: Mean number of infected Steinernema jeffreyense and Steinernema yirgalemense juveniles (± SE) per Tuta absoluta larva at the respective concentrations applied to tomato seedlings in a greenhouse. Concentrations Steinernema jeffreyense Steinernema yirgalemense (IJs mL-1) Mean IJs±SE Mean IJs±SE 250 1.72±0.16a 1.84±0.18ab 500 1.58±0.11a 1.33±0.10a 1 000 1.66±0.11a 1.74±0.13ab 2 000 2.25±0.18a 2.51±0.23b Means within a column followed by the same lowercase letter are not significantly different at P<0.05 (Kruskall-Wallis). Corrected percentage mortality ± SE 64 3.4. Discussion Chemical control of pests and weeds affects non-target organisms, which are exposed to the toxins (Sanchez-Bayo, 2011). It is also detrimental to human health causing acute and chronic problems (Paoletti and Pimentel, 2000). Although pesticides benefit producers economically (Pimentel and Ali, 1998), health risks and the development of resistance to insecticides necessitate a change in management strategies to better control insect pests (Ehlers, 1996). The use of EPNs as a biological control practice minimizes the impact on human and environmental health (Ehlers, 1996). Tuta absoluta larvae feed on parenchyma cells inside leaf galleries (Sannino and Espinosa, 2010) and foliar applications of EPNs will therefore be needed for its control. Foliar application does, however, expose EPNs to the harsh environmental conditions and limits the efficacy of EPNs applied against foliar pests (Lacey and Georgis, 2012). The effective control of pests on foliar parts as well as in stems and trunks with application of EPNs under field conditions was, however, reported by Tomalak et al. (2005). Another important consideration is the susceptibility of the target insect species to the EPN species under consideration. This was proven by Belair et al. (2003) who reported that S. carpocapsae applied as a foliar application, was not effective in controlling the cabbageworm, Artogeia rapae (Linneaus) (Lepidoptera: Pieridae), despite its efficacy against a wide range of insect pests. In this study, both S. jeffreyense and S. yirgalemense controlled T. absoluta larvae effectively after foliar application to infested tomato seedlings in a greenhouse. High mortality rates of more than 80% were achieved at high application rates of 1 000 and 2 000 IJs mL-1. These were lower than the mortality rates of the two EPNs in laboratory bioassays (Chapter 2). It was, however, in agreement with the results reported by Batalla-Carrera et al. (2010) who reported high efficacy of control of T. absoluta larvae (87 – 95%) with S. carpocapsae, Steinernema feltiae (Filipjev, 1934) Wouts, MráÏcek, Gerdin and Bedding, 1982 and Heterorhabditis bacteriophora Poinar, 1976 applied at 1 000 IJs mL-1 to tomato seedlings in a greenhouse experiment. Kamali et al. (2018) reported 50% efficacy of T. absoluta larvae of various instars by S. carpocapsae and H. bacteriophora at a dosage rate of 50 IJs cm-2 applied without an adjuvant in a greenhouse experiment. This difference in efficacy reported between the two studies could be explained by the difference in larval instars exposed as well as the addition of an adjuvant in the foliar application of Batalla-Carrera et al. (2010). Adjuvants increase the efficacy of control by EPNs (Kamali et al., 2018; Platt et al., 2019). For a 95% control of the leafminer, Liriomyza bryoniae (Kaltenbach) (Diptera: Agromizidae) with S. feltia, a high dosage rate of 10 000 IJs mL-1 was reported (Williams and Walters, 2000). 65 This difference in dosage rates necessary were ascribed to the differences in insect behaviour as well as the types of leaf mines created by respective insect species (Batalla-Carrera et al., 2010). Liriomyza bryoniae creates only one entry route during oviposition with no other entry made by the larva during its development inside the leaf mine. Tuta absoluta larvae create big tunnels which provides for entry by EPNs as well as protection against harsh environmental conditions. Insect larvae of this species also move between leaves which give EPNs effortless access to their host (Batalla-Carrera et al., 2010). A high percentage larval mortality, although infected with only a few nematodes was found in this study. It could be explained by the bacterial release by the IJs (Gaugler et al., 1994). Bacteria are released before the host immune response encapsulate and melanise invading IJs (Wang et al., 1994). Lethal bacteria are also carried on the nematode surface or enter insect host through wound that nematodes created on point of entry and could kill hosts on invasion (Poinar and Kaul, 1981). Insect mortality should be recorded to determine the insecticidal effect of EPNs, and penetration rate of insects by IJs can be used to compare infectivity of entomopathogenic nematode strains, but a lack of correlation between pathogenicity and levels of host invasion can still occur (Caroli et al., 1996). The low number of IJs of both species found in the host larvae, could possibly be explained by the size of the T. absoluta larvae and the size of the nematodes. A small size host affects the number of nematodes that infects the host (Bastidas et al., 2014). A higher number of IJs recorded in some larvae, did, however, indicate that development and reproduction of nematodes occurred after invasion also. The foraging traits of EPNs are important to take into account when different species are considered for foliar applications. The foraging behaviour of the two species did also not explain the low number of IJs that penetrated T. absoluta larvae. Steinernema yirgalemense is an intermediate forager, which acts as an ambusher (wait for its prey) as well as a cruiser that search for prey (Dlamini et al., 2020). Steinernema jeffreyense acts as an ambusher (Dlamini et al., 2020) that waits for hosts (Campbell and Gaugler, 1997). A similar number of S. yirgalemense and S. jeffreyense IJs penetrated T. absoluta larvae in these greenhouse experiments. Host seeking activity as explanation for their efficacy, was therefore excluded. 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Biocontrol Science and Technology 6(3):379-388. TIBCO SOFTWARE INC. 2017. Statistica (data analysis software system), version 13.3. www.tibco.com. TOMALAK, M., PIGGOTT, S. and JAGDALE, G.B. 2005. Glasshouse applications. pp 147- 166. In: Nematodes as biocontrol agents. (Eds. Grewal, P.S., Ehlers, R-U. and Shapiro-Ilan, D.I.). CAB International: Oxon. WANG, Y., GAUGLER, R. and CUI, L. 1994. Variations in immune response of Popillia japonica and Acheta domesticus to Heterorhabditis bacteriophora and Steinernema species. Journal of Nematology 26(1):11-18. WILLIAMS, E.C. and WALTERS, K.F.A. 2000. Foliar application of the entomopathogenic nematode Steinernema feltiae against leafminers on vegetables. Biocontrol Science and Technology 10(1):61-70. 70 Chapter 4: Conclusion and recommendations The South American tomato pinworm, Tuta absoluta Meyrick (Lepidoptera: Gelechiidae) invaded Africa in 2008, and continued to spread rapidly. It is now a key, devastating insect pest in the tomato-producing areas in Northern and sub-Saharan Africa (Mansour et al., 2018). It also became a threat to South African producers after its detection in the country in 2016 (Visser et al., 2017). Current management tactics for T. absoluta are mainly based on monitoring with sex pheromone traps and application of synthetic insecticides (Biondi et al., 2018; Mansour et al., 2018). Although application of insecticides was considered the most effective control method for T. absoluta (Desneux et al., 2010), the importance of using an integrated pest management (IPM) strategy for its control and to slow down insecticide resistant development, was earlier emphasized by Georghiou (1983). Resistance to insecticides did, however, develop (Siqueira et al., 2000; Lietti et al., 2005; Silva et al., 2011; Haddi et al., 2012; Roditakis et al., 2013; 2015; 2018). Tuta absoluta populations from Greece, Italy, Israel, Spain and UK in the Euro- Asian region, showed resistance to emamectin benzoate, indoxacarb and spinosad. Except for T. absoluta populations in Spain, resistance was also reported to chlorantraniliprole (Roditakis et al., 2018). The authors attributed this finding to an IPM program which included non-chemical measures and the rotation of different mode of action classes when insecticides were applied. Host plant resistance has been investigated with susceptibility testing of tomato cultivars to T. absoluta. Larval survival, development time, fecundity and fertility of the pest were used as parameters (Gharekhani and Salek-Ebrahimi, 2014a,b; Ghaderi et al., 2017; Krechemer and Foerster, 2017; Azadi et al., 2018). Results reported by these authors were similar to that of Oliveira et al. (2009) indicating that some tomato cultivars are less susceptible to T. absoluta, but no cultivar is entirely resistant to this pest. Although tomato cultivars containing Bt Cry proteins have been developed and proven to be successful in controlling T. absoluta, it has not been released for commercial use (Selale et al., 2017). The failure to control T. absoluta may have serious economic impacts, and strategies to control it with natural enemies, is therefore important (Bajonero et al., 2008). Over the years, many parasitoids in various families were reported in the area where T. absoluta originated from, as well as in Asia and Europe (Faria et al., 2008; Desneux et al., 2010; Doġanlar and Yiğit, 2011; Loni et al., 2011; Luna et al., 2012; 2015; Al-Jboory et al., 2012, Cabello et al., 2012; Ferracini et al., 2012; Zappalà et al., 2012; Biondi et al., 2013; Chailleux et al., 2013; El–Arnaouty et al., 2014; Gabarra et al., 2014; Bayram et al., 2016). To date, parasitoids from only four families have been reported in Africa, namely Braconidae (Boualem et al., 2012; Abbes et al., 2014; 71 Idriss et al., 2018), Eulophidae (Boualem et al., 2012; Abbes et al., 2014), Ichneumonidae (Boualem et al., 2012) and Pteromalidae (Idriss, 2019). Pesticides affects natural enemies and will consequently reduce biological control of this pest (Luna et al., 2007). It is therefore important to recommend approaches consisting of reduced-risk insecticide compounds, such as plant extracts or microbial-based selective insecticides, and/or biorational alternatives for effective control of T. absoluta in Africa and worldwide (Mansour et al., 2018). The correct use of insect-proof nets (impregnated with insecticides or not), the use of pheromone-based tools (especially for mass trapping) and the conservation and augmentation of indigenous biological control agents have also recommended (Mansour et al., 2018). Mirids, especially Nesidiocoris tenuis (Reuter) (Hemiptera: Miridae) used in IPM programs also provide effective control of T. absoluta (Mollá et al., 2011). A variety of insect pests are controlled with entomopathogens such as fungi, bacteria and nematodes (Lacey et al., 2001). Control of T. absoluta with Metarhizium anisopliae var. anisopliae (Metsch.) Soroki, Beauveria bassiana (Balsamo) Vuillemin (Ínanli et al., 2013) and Bacillus thuringiensis (Berliner), has also been reported (González-Cabrera et al., 2011). The increased rate of insecticide resistance development prompted the search for new, effective, environmentally safe control methods and increased the interest in entomopathogenic nematodes (EPNs) since their discovery (Abate et al., 2017; Guedes et al., 2019). Entomopathogenic nematodes are produced commercially to control insect pests globally (Abate et al., 2017). The EPNs, Heterorhabditis bacteriophora Poinar, 1976 Steinernema feltiae (Filipjev, 1934) Wouts, MráÏcek, Gerdin and Bedding, 1982 and Steinernema carpocapsae (Weiser, 1955) Wouts, MráÏcek, Gerdin and Bedding, 1982, were evaluated and found to effectively control T. absoluta (Batalla-Carrera et al., 2010; Van Damme et al., 2016; Kamali et al., 2018). Importation of exotic species without a permit and a full impact study is, however, not allowed in South Africa due to the regulations of the Department of Agriculture, Forestry and Fisheries (DAFF), under the Agricultural pest act 36 (Klein et al., 2011; DAFF, 2020). With these restrictions in place research for control of T. absoluta with native biological control agents is therefore warranted. In this study, four native EPN species viz. Steinernema jeffreyense Malan, Knoetze, Tiedt, 2015, Steinernema yirgalemense Nguyen, Tesfamariam, Gozel, Gaugler and Adams, 2005. Heterorhabditis baujardi Phan, Subbotin, Nguyen and Moens, 2003 and Heterorhabditis noenieputensis Malan, Knoetze and Tiedt, 2014, were found to be highly effective in killing fourth instar T. absoluta larvae, with 100% larvae killed in laboratory bioassays. The efficacy 72 of these EPNs was, however, lower against T. absoluta pupae. The highest pupal mortality was caused by S. yirgalemense (41.7%), and less than 30% mortality was recorded for S. jeffreyense, H. baujardi and H. noenieputensis (Chapter 2). The number of IJs that penetrated T. absoluta larvae and pupae in the laboratory bioassays was also low. Fewer S. yirgalemense IJs penetrated T. absoluta larvae compared to IJs of the other species used in this study. The number of IJs that penetrated T. absoluta pupae, was also fewer compared to the number of IJs that penetrated larvae. On the basis of these laboratory bioassays, all four EPN species tested have potential for use in an IPM system for control of T. absoluta larvae (Chapter 2). Subsequent greenhouse trials were conducted to investigate the potential of S. yirgalemense and S. jeffreyense for control of T. absoluta larvae in tomato cultivated in greenhouses (Chapter 3). High T. absoluta larval mortality rates were recorded with S. yirgalemense and S. jeffreyense applied at 1 000 IJs mL-1 to tomato foliage in a greenhouse infested with third and fourth instar larvae. It is in agreement with high efficacy of control of T. absoluta larvae reported with S. carpocapsae, Steinernema feltiae and Heterohabditis bacteriophora, also applied at 1 000 IJs mL-1 to tomato seedlings in a greenhouse (Batalla-Carrera et al., 2010). These studies indicated the efficacy of various EPN species to kill T. absoluta larvae and their potential to act as successful biological control agents of this pest. The importance of high dosage rates of 1 000 IJs mL-1 and higher, under moderate temperature and high humidity conditions was also confirmed in the current study. The size of the T. absoluta larvae to which EPNs are applied, is also important for successful control (Van Damme et al., 2016). Third and fourth instar larvae were reported to be the most susceptible. Control was lower when EPNs were applied to smaller (first and second instar) larvae (Bastidas et al., 2014). A decrease in host size and an increase in nematode size decrease invasion of the host by EPNs (Bastidas et al., 2014; Van Damme et al., 2016; Kamali et al., 2018). Third and fourth instar T. absoluta larvae should therefore be the stage to target when applying EPNs for control of this pest. Both H. baujardi and H. noenieputensis were also highly effective in killing T. absoluta larvae in laboratory bioassays of this study. It justifies evaluation of these species for control of T. absoluta in tomato under controlled greenhouse conditions. Future research could also include the evaluation of other EPN species isolated in South Africa for control of T. absoluta larvae. The susceptibility of T. absoluta larvae from different instars to all native EPNs found to be effective in preliminary studies, could also be investigated in future studies. It will be an added advantage to an IPM program if a species could be identified that is effective against the 73 smaller instar larvae. Furthermore, when effective control of T. absoluta in controlled greenhouse conditions could be confirmed for native EPNs, research on the efficacy of EPN field applications, or improvement of control with the addition of adjuvants in foliar applications, could be conducted. 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First records of the tomato leaf miner Tuta absoluta (Meyrick, 1917) (Lepidoptera: Gelechiidae) in South Africa. BioInvasions Records 6(4):301-305. ZAPPALÀ, L., BERNARDO, U., BIONDI, A., COCCO, A., DELIPRRI, S., DELRIO, G., GIORGINI, M., PEDATA, P.C., RAPISARDA, C., TROPEA GARZIA, G. and SISCARO, G. 2012. Recruitment of native parasitoids by the exotic pest Tuta absoluta in Southern Italy. Bulletin of Insectology 65(1):51-61. 79 Appendix A Declaration of language editing Language editing statement To whom this may concern, I, Prof. Hannalene du Plessis, hereby declare that the thesis titled: “Efficacy of entomopathogenic nematodes for control of Tuta absoluta in South Africa” by Odette Coleman has been edited for language correctness and spelling by the supervisors. No changes were made to the academic content or structure of this work. 16 May 2020 Prof. Hannalene du Plessis Date 80 Appendix B FNAS Ethics Committee Tel: +27 18 3892598 Fax: +27 18 3892052 Email lesetja.motadi@nwu.ac.za Internet http://www.nwu.ac.za Date: 22-May- 2019 Dear Researcher, Re: Ethics weaver This letter serves as confirmation that based on the scientific committee assessment of the research proposal concluded that: Student: Ms O Coleman (24093807) Title: Efficacy of entomopathogenic nematodes for control of Tuta absoluta in South Africa. Does not require ethical clearance as the study has no/low risk. Supervisor (s) Hannalene Du Plessis ---------------------------------------------- Signature: Chairperson of FNAS Ethics Committee Prof Lesetja Motadi 81 Appendix C