HOST RANGE AND MULTITROPHIC INTERACTIONS BETWEEN THE PARASITOID COTESIA VANESSAE (HYMENOPTERA: BRACONIDAE) AND NOCTUIDAE (LEPIDOPTERA) HOSTS IN NORTH AMERICA VINCENT ALAIN DANIEL HERVET Diplôme Universitaire et Technologique Génie Biologique, Spécialité Agronomie, Université de Picardie Jules Verne, Amiens, France, 2007 Diplôme d’Ingénieur Agriculture, Spécialité Production Végétale, Institut Polytechnique LaSalle Beauvais, Beauvais, France, 2010 A Thesis Submitted to the School of Graduate Studies of the University of Lethbridge in Partial Fulfillment of the Requirements for the Degree DOCTOR OF PHILOSOPHY (BIOSYSTEMS AND BIODIVERSITY) (Evolutionary Biology) Department of Biological Sciences University of Lethbridge LETHBRIDGE, ALBERTA, CANADA © Vincent Alain Daniel Hervet, 2017 HOST RANGE AND MULTITROPHIC INTERACTIONS BETWEEN THE PARASITOID COTESIA VANESSAE (HYMENOPTERA: BRACONIDAE) AND NOCTUIDAE (LEPIDOPTERA) HOSTS IN NORTH AMERICA VINCENT ALAIN DANIEL HERVET Date of Defense: November 22, 2016 Dr. Robert Laird Associate Professor Ph.D. Co-supervisor Dr. Kevin Floate Research Scientist Ph.D. Co-supervisor Dr. Theresa Burg Associate Professor Ph.D. Thesis Examination Committee Member Dr. Matthew Letts Associate Professor Ph.D. Thesis Examination Committee Member Associate Dean of Arts and Science Dr. Rob Bourchier Research Scientist Ph.D. Internal Examiner Adjunct Professor Dr. Jeff Harvey Research Scientist Ph.D. External Examiner Netherlands Institute of Ecology Wageningen, Netherlands Dr. Brent Selinger Professor Ph.D. Chair, Thesis Examination Committee ii To my parents and my wife, for having always supported my passion for entomology iii ABSTRACT Caterpillars in the family Noctuidae are important crop pests. After discovering the parasitoid Cotesia vanessae in the Nearctic region, which was previously only known from the Palearctic and Afrotropic regions, I investigated its fundamental host range on North American Lepidoptera and found that it could parasitize a large number of species of Noctuidae. Its fitness varied among host species, and Plusiinae species appeared to be the best hosts. Nearly all hosts in this experiment were reared on McMorran diet, which appeared to have varying suitability among Lepidoptera species. Through a second experiment, I showed that the fitness of the parasitoid and that of its host varied according to host food quality. Thus, parasitoid fitness on different hosts is partly conditioned by diet quality. This study shows that C. vanessae can parasitize a large number of Noctuidae and can be easily mass-reared, which could make it an ideal biological control agent. iv ACKNOWLEDGEMENTS I would like to thank my advisors Rob Laird and Kevin Floate for their outstanding support and advice during this research. I also thank Paul Coghlin, Bob Byers, and Héctor Cárcamo and his lab for various technical assistance; John Dedes and the staff at IPS for providing custom orders of McMorran diet; José Fernández-Triana and Mark Shaw for help in identifying parasitoids, collaboration, and advice; Kayla Johnson, Jayden Dyck, Briana Smith, Braidon Schaufert, Marko Mićović, and Abbey Brusky for technical support and their great enthusiasm; Henry Murillo, Michaela Schmitke, Shelley Barkley, Melissa Fry, and Jessica Kwon for providing insect species; and Stephanie Harris for providing pictures of parasitoids. This research was supported by the Canola Council of Canada (Canola Agronomic Research Program Project 2012–1, to K.D.F.), Alberta Conservation Association (Grants in Biodiversity, to V.A.D.H.), and the Natural Science and Engineering Research Council of Canada (Discovery Grant, to R.A.L.). v TABLE OF CONTENTS Abstract ...................................................................................................................................... iv Acknowledgements ..................................................................................................................... v Table of contents ........................................................................................................................ vi List of Tables ............................................................................................................................. viii List of Figures .............................................................................................................................. ix List of Abbreviations ................................................................................................................... xi Chapter 1: General introduction ................................................................................................ 1 1.1. Context ............................................................................................................................ 1 1.2. Goals of the study ............................................................................................................ 5 Chapter 2: Identification and biology of Cotesia vanessae. First report of this species in North America ..................................................................................................................................... 18 2.1. Abstract ......................................................................................................................... 18 2.2. Introduction ................................................................................................................... 18 2.3. Collections ..................................................................................................................... 20 2.3.1. From Ontario .......................................................................................................... 20 2.3.2. From Alberta ........................................................................................................... 21 2.4. Current known range ..................................................................................................... 22 2.5. Conclusion ..................................................................................................................... 23 Chapter 3: Evaluation of the host range and fitness of the parasitoid Cotesia vanessae on North American Lepidoptera, particularly pestiferous Noctuidae ........................................... 29 3.1. Abstract ......................................................................................................................... 29 3.2. Introduction ................................................................................................................... 30 3.3. Materials and Methods ................................................................................................. 33 3.3.1. Insect collection ...................................................................................................... 33 3.3.2. Insect identification ................................................................................................ 34 3.3.3. Colony rearing ........................................................................................................ 34 3.3.4. Host-range and host-fitness testing ....................................................................... 36 3.3.5. Phylogeny of lepidopteran species ......................................................................... 40 3.3.6. Investigating parasitized caterpillars that did not produce parasitoids ................. 40 3.3.7. Data analyses .......................................................................................................... 42 3.4. Results ........................................................................................................................... 44 3.4.1. Fitness of C. vanessae in relation to host species .................................................. 44 3.4.2. Oviposition testing .................................................................................................. 48 3.4.3. Testing if C. vanessae causes premature death of Lepidoptera larvae .................. 49 3.5. Discussion ...................................................................................................................... 50 Chapter 4: A review of the McMorran diet for rearing Lepidoptera species with addition of a further 39 species ..................................................................................................................... 77 4.1. Abstract ......................................................................................................................... 77 4.2. Introduction ................................................................................................................... 77 4.3. Materials and Methods ................................................................................................. 79 4.4. Results and Discussion ................................................................................................... 82 Chapter 5: Multi-trophic effects of protein-restricted diet on the development and fitness of the cabbage looper, Trichoplusia ni (Lepidoptera: Noctuidae), and its parasitoid Cotesia vanessae (Hymenoptera: Braconidae) ..................................................................................... 96 5.1. Abstract ......................................................................................................................... 96 vi 5.2. Introduction ................................................................................................................... 97 5.3. Materials and methods ............................................................................................... 100 5.3.1. Studied insects ...................................................................................................... 100 5.3.2. Diet preparation ................................................................................................... 101 5.3.3. Insect handling...................................................................................................... 102 5.3.4. Measurements ...................................................................................................... 103 5.3.4.1. Data inclusion / exclusion ............................................................................. 103 5.3.4.2. Measurements for non-parasitized T. ni ...................................................... 104 5.3.4.3. Variables measured for C. vanessae ............................................................. 104 5.3.4.4. Measurement of food eaten by T. ni ............................................................ 105 5.3.4.5. Data analyses ................................................................................................ 106 5.4. Results ......................................................................................................................... 107 5.4.1. Trichoplusia ni ....................................................................................................... 107 5.4.2. Trichoplusia ni rates of survival and successful parasitism .................................. 108 5.4.3. Trichoplusia ni food consumption ........................................................................ 108 5.4.4. Cotesia vanessae development time .................................................................... 109 5.4.5. Cotesia vanessae numbers and mass ................................................................... 110 5.4.6. Cotesia vanessae F2 development, numbers, and mass ...................................... 111 5.5. Discussion .................................................................................................................... 111 Chapter 6: General discussion ................................................................................................ 133 References .............................................................................................................................. 137 Appendix 1: Some observations of Cotesia vanessae natural history .................................... 158 A.1. Engaging an unlikely host ............................................................................................ 158 A.2. Ovipositor attachment ................................................................................................ 158 A.3. Climbing prior to parasitoid egression ........................................................................ 158 A.4. Parasitoid larvae egression ......................................................................................... 158 A.5. Larvae color according to host taxa ............................................................................ 159 vii LIST OF TABLES Table 1.1. Parasitoids of Euxoa auxiliaris, Euxoa messoria, and Euxoa ochrogaster ................. 7 Table 1.2. Predators of Euxoa auxiliaris, Euxoa messoria, and Euxoa ochrogaster ................. 14 Table 1.3. Pathogens of Euxoa auxiliaris, Euxoa messoria, and Euxoa ochrogaster ............... 17 Table 3.1. Identity of the species studied ................................................................................ 65 Table 3.2. Species that didn’t produce Cotesia vanessae ........................................................ 68 Table 4.1. Species of Lepidoptera reared on McMorran diet .................................................. 84 Table 4.2. Lepidoptera species that did not successfully develop on McMorran diet ............ 92 Table 5.1. Diet manipulations ................................................................................................ 117 Table 5.2. Summary of the different treatments at the start of the experiment .................. 118 Table 5.3. Outcome of Trichoplusia ni individuals at the end of the experiment .................. 119 viii LIST OF FIGURES Figure 2.1. Parasitism (%) of C. chalcites and T. ni by C. vanessae collected from Ontario ..... 24 Figure 2.2. Neighbour-joining tree for CO1 sequences for Cotesia species ............................. 25 Figure 2.3. Cotesia vanessae adults ......................................................................................... 26 Figure 2.4. Cotesia vanessae immatures .................................................................................. 27 Figure 2.5. Distribution of Cotesia vanessae ............................................................................ 28 Figure 3.1. Cotesia vanessae parasitizing an early 4th instar Trichoplusia ni. .......................... 67 Figure 3.2. Fate of Lepidoptera larvae of species that produced C. vanessae ......................... 69 Figure 3.3. Neighbour-joining tree for CO1 sequences of known hosts of C. vanessae and other tested species within Noctuoidea .......................................................................... 70 Figure 3.4. Cotesia vanessae total development time on each host species........................... 71 Figure 3.5. Cotesia vanessae brood sizes on each host species ............................................... 72 Figure 3.6. Cotesia vanessae average individual mass per brood on each host species. ........ 73 Figure 3.7. Cotesia vanessae estimated brood mass per host ................................................. 74 Figure 3.8. Immature Cotesia vanessae ................................................................................... 75 Figure 3.9. Effect of parasitism on Spodoptera frugiperda ...................................................... 76 Figure 4.1. Set up used to rear caterpillars in cages on the McMorran diet ........................... 93 Figure 4.2. Set up for rearing cannibalistic species in cages .................................................... 94 Figure 4.3. Polar Plastic Ltd. cup (240 ml) used to rear individual caterpillars ........................ 95 Figure 5.1. Trichoplusia ni pupae ........................................................................................... 120 Figure 5.2. Trichoplusia ni larvae development time on diets containing different amounts of protein ............................................................................................................................ 121 Figure 5.3. Trichoplusia ni mass of pupae from larvae that developed on diets containing different amounts of protein .......................................................................................... 122 Figure 5.4. Estimated dry mass of food consumed by T. ni larvae developing on diets containing 43 to 8.1 g/L of protein ................................................................................. 123 Figure 5.5. Cotesia vanessae development time within T. ni larvae that developed on diets containing different amounts of protein ........................................................................ 124 Figure 5.6. Cotesia vanessae development time within their cocoons from T. ni hosts that developed on diets containing different amounts of protein ........................................ 125 Figure 5.7. Cotesia vanessae total development time in T. ni hosts that developed on diets containing different amounts of protein ........................................................................ 126 Figure 5.8. Scatterplot with trend lines showing Cotesia vanessae development time inside their host in relation to their development time inside cocoons, on each diet type .... 127 Figure 5.9. Cotesia vanessae brood size per type of diet from T. ni hosts that developed on diets containing different amounts of protein ............................................................... 128 Figure 5.10. Average dry mass of individual adult Cotesia vanessae per brood from T. ni hosts that developed on diets containing different amounts of protein ................................ 129 Figure 5.11. Scatterplot with trend lines showing C. vanessae brood size in relation to average dry mass of individuals from T. ni fed diets containing different amounts of protein ............................................................................................................................ 130 Figure 5.12. Scatterplot with trend lines showing number of C. vanessae adult per brood in relation to estimated total wasp mass of these broods, from T. ni fed diets containing different amounts of protein .......................................................................................... 131 ix Figure 5.13. F2 C. vanessae .................................................................................................... 132 Figure A.1. Last instar Trichoplusia ni larva that died following egression of Cotesia vanessae from its body .................................................................................................................. 160 Figure A.2. Same T. ni larva as in Fig. A.1 dissected dorsally ................................................. 161 Figure A.3. Top view of a Cotesia vanessae cocoon mass ..................................................... 162 Figure A.4. Cotesia vanessae cocoon mass lateral side ......................................................... 163 x LIST OF ABBREVIATIONS AAFC: Agriculture and Agri-Food Canada NC: North Carolina AB: Alberta ND: North Dakota AK: Alaska NE: Nebraska AL: Alabama NH: New Hampshire ANCOVA: Analysis of Covariance NJ: New Jersey ANOVA: Analysis of variance NL: Newfoundland and Labrador AR: Arkansas NM: New Mexico AZ: Arizona NS: Nova Scotia BC: British Columbia NT: Northwest Territories BOLD: Barcode of Life Data Systems NU: Nunavut CA: California NY: New York CAD: Canadian Dollar OH: Ohio CO: Colorado OK: Oklahoma CO1: Cytochrome c oxidase subunit 1 ON: Ontario CT: Connecticut OR: Oregon DC: District of Columbia PA: Pennsylvania DNA: Desoxyribonucleic Acid PE: Prince Edward Island FL: Florida Q1: 1st quartile GA: Georgia Q3: 3rd quartile HI: Hawaii QC: Quebec HSD: Honest Significant Difference RI: Rhode Island IA: Iowa SC: South Carolina ID: Idaho SD: South Dakota IL: Illinois SE: Standard error of the mean IN: Indiana SK: Saskatchewan IPS: Insect Production Services Sp.: species (singular) IQR: Interquartile Range Spp.: species (plural) K2P: Kimura 2-Parameter distance TN: Tennessee KOH: Potassium hydroxide TX: Texas KS: Kansas US: United States KY: Kentucky USA: United States of America L:D: Light:Dark cycle USD: American Dollar LA: Louisiana USDA: United States Department of Agriculture MA: Massachusetts UT: Utah MB: Manitoba UV: Ultraviolet MD: Maryland VA: Virginia ME: Maine VT: Vermont MI: Michigan WCCP: Western Committee on Crop Pests MN: Minnesota WA: Washington MO: Missouri WI: Wisconsin MS: Mississippi WV: West Virginia MT: Montana YT: Yukon NB: New Brunswick xi Chapter 1 CHAPTER 1: GENERAL INTRODUCTION 1.1. Context Larvae of many species of moths in the family Noctuidae are important pests of cultivated plants in North America and elsewhere in the world. For example, the pale western cutworm, Agrotis orthogonia Morrison, caused an average annual loss of over 2- million USD in 1920 and 1921 in Montana (Mabee, 1929), $4.7-million (likely CAD) annually between 1929 and 1932 in the prairies of Canada (McMillan, 1935), and 17-million USD annually between 1955 to 1975 in eight US states combined (Idaho, Montana, South Dakota, Wyoming, Nebraska, Colorado, Kansas, and Oklahoma). From about 1955 to 1975, the army cutworm, Euxoa auxiliaris (Grote), caused an annual loss of nearly 8-million USD in four US states combined (Montana, South Dakota, Colorado, and Oklahoma) (USDA, 1977); and in 1990 in southern Alberta, over 10,000 ha had to be either sprayed with insecticide or reseeded due to damage from this pest (Jones et al., 1990). In recent years, the cotton bollworm, Helicoverpa armigera (Hübner), caused an estimated annual worldwide loss of 3- billion USD (Sharma et al., 2014). Cutworms have been an enduring problem on the Canadian prairies for over a century (Jacobson, 1969, 1971; Jacobson et al., 1961; Strickland, 1916, 1923). In recent years, particularly damaging outbreaks have renewed interest in these pests (WCCP, 2012, 2013), which was the stimulus for initiating this thesis research. Larvae of these pests often go by common names such as cutworms, armyworms and loopers, according to the life history of the larva, and dart moth as adults. While armyworms and loopers are usually a problem on mature plants, as they feed on developing fruiting bodies, cutworms are mainly a problem in the spring when crops start to grow. Cutworms can be classified as subterranean, above-ground, or climbing species (Walkden, 1950). Subterranean species feed exclusively on the small part of the stem below the ground; thus, a single cutworm can destroy numerous individual plants. Above-ground species are mainly 1 Chapter 1 foliage feeders, feeding primarily at dawn and dusk, and hide in the soil and under debris during the day, and late instars can cut the stems of seedlings. Climbing cutworms, armyworms, and loopers usually remain on the crown of plants where they feed primarily on the foliage and fruiting bodies. Armyworms have a tendency to congregate and move together “as an army” from one area to a next as plants become consumed (Capinera, 2001; Walkden, 1950). Loopers move forward by drawing their posterior end toward their anterior end, thus forming a partial loop. Loopers are also called “semi-loopers” to avoid confusion with caterpillars of the family Geometridae that make a full loop with their body. In this thesis, Geometridae are not dealt with and for practicality the term “looper” is used when referring to noctuid species in the subfamily Plusiinae. The abundance of cutworms and their natural enemies, both in time and in space, is largely conditioned by biotic and abiotic conditions. Cutworms can generally be found at low densities throughout their range, but during outbreaks they can completely destroy a crop (personal observation). Specific conditions during oviposition time, mainly of soil texture, structure, humidity, and cover, drive cutworm moths to oviposit in certain fields rather than others; e.g., Euxoa spp., Agrotis orthogonia, and Agrotis segetum (Denis & Schiffermüller) prefer to lay eggs in dry loose sandy or dusty soil; thus fields tilled in late summer to early fall are of higher risk of cutworm outbreaks in the spring, and outbreaks are often composed of multiple species (Esbjerg et al., 2009; Jacobson, 1971; Jacobson et al., 1961; Wagner et al., 2012). In some species, spring rains contribute to the mortality of cutworm larvae; e.g., wet conditions increase mortality rates of A. segetum and A. orthogonia (Cook, 1926; Esbjerg et al., 2009; Seamans, 1935). Cutworm outbreaks will in turn induce a build-up of the populations of their natural enemies that will then slowly reduce pest populations (Jacobson, 1971). Moist spring conditions have been reported to be detrimental to cutworms by increasing incidence of parasitism and disease (Jacobson, 1971; Seamans, 1935). Prolonged severe winds also may be detrimental to parasitoids (Strickland, 1923). These conditions 2 Chapter 1 generally drive important outbreaks to occur episodically. Cutworm outbreaks can be prevented, reduced, or stopped by agronomic practices, chemical control, and naturally occurring biological control agents (Jacobson, 1971; Strickland, 1916). The latter, also referred to as “natural enemies” include pathogens, predators, and parasitoids (Capinera, 2001; Jacobson, 1971). Many species of natural enemies have been reported to kill cutworm pest species in North America, including pests of particular economic importance such as the redbacked cutworm, Euxoa ochrogaster (Guenée), darksided cutworm, Euxoa messoria (Harris), and army cutworm, Euxoa auxiliaris (Grote) (Lepidoptera: Noctuidae) (Tables 1.1, 1.2, 1.3) (Gavloski et al., 2013). Parasitoids have long been identified as important naturally occurring biological control agents of cutworms in the prairies of Canada (Jacobson, 1971; Strickland, 1923). The term “parasitoid” was coined in 1913 in German by O. M. Reuter and in 1914 in English by W. M. Wheeler (Wheeler, 1914) to define parasitic insects that, unlike parasites, only use a single host throughout their larval stage, that always end up killing the host, and whose adults are free living. The parasitoid lifestyle has evolved independently multiple times in the insect orders Hymenoptera, Diptera, Coleoptera, Neuroptera, Lepidoptera, and Trichoptera (Godfray, 1994). Most hymenopteran parasitoids use their ovipositor to insert eggs within the body of their host. During oviposition, parasitoids may inject other substances into the host, such as venom, calyx fluid, and viruses, which can interfere with the development of the host and combat its immune system to aid the development of their offspring (Lanzrein et al., 2012). Knowledge of host-parasitoid interactions can guide use of parasitoids in different types of biocontrol programs targeting pest species. Common biological control strategies include innundative (flooding the environment with biological control agents; a strategy mainly used in greenhouses), classical (introducing an exotic biological control agent to control an exotic invasive pest), neoclassical (introducing a foreign biological control agent to control a native pest), and conservation (preservation or 3 Chapter 1 modification of the environment to enhance populations of biological control agents) programs (Bellows et al., 1999; van den Bosch et al., 1982). Multitrophic interactions (in this case between plant-host and host-parasitoids) drive host suitability and fitness of parasitoids. To locate suitable hosts, parasitoids rely on cues produced by the environment that their hosts live in, and many studies have found that parasitoids of herbivorous insects are attracted to herbivore-induced plant volatiles. For example, Cotesia rubecula (Marshall) and Cotesia glomerata (L.) (Hymenoptera: Braconidae) are attracted to volatiles produced by damaged leaves of cabbage, Brassica oleracea L. (Brassicaceae) (Agelopoulos et al., 1994; Mattiacci et al., 1994, 1995; Steinberg et al., 1993), and Cotesia marginiventris (Cresson) is attracted to corn, Zea mays L. (Poaceae), damaged by Spodoptera exigua (Hübner) larvae (Turlings et al., 1990). In addition, parasitoid fitness can be affected when hosts feed on poor quality diet, either because of the presence of plant defensive secondary plant compounds such as tannins, terpenes, flavones, phenols, and glucosinolates (Agrawal et al., 2015; Barbosa, et al. 1986; Gols et al., 2008a, 2008b, 2008c; Harvey 2005; Harvey et al., 2007, 2011; Ode 2006; Rhoades et al., 1976; Soler et al., 2005), or because the diet is nutritionally unbalanced (Bloem et al., 1990; Pimentel, 1966). The fitness of parasitoids can be assessed by measuring different parameters. These can include development time, survival rate, adult longevity, fertility, fecundity, body size, body mass, efficiency in locating hosts, host patch-exploitation strategy, offspring sex ratio, and offspring performance (Boivin et al., 2004; Harvey, 2005; Roitberg et al., 2001; Vos et al., 2003). Because of limited time and resources, normally only a subset of these parameters can be estimated experimentally. Body size and mass are the most commonly used proxies of fitness because their increase is often positively correlated with longevity and fecundity (Harvey, 2005; Roitberg et al., 2001). However, a too large increase in one parameter can result in a decrease of other parameters of fitness; e.g., too large body size can result in 4 Chapter 1 increased predation (West et al., 1996). Hence, multiple parameters should be measured to increase the assessment accuracy of the fitness of individuals (Roitberg et al., 2001). 1.2. Goals of the study The current research was initiated following the discovery in the Nearctic region of the parasitoid Cotesia vanessae (Reinhard) (Braconidae) (Chapter 2). This discovery arose from recovery of C. vanessae from the pest species tomato looper, Chrysodeixis chalcites (Esper), and cabbage looper, Trichoplusia ni (Hübner), collected in greenhouses and fields in southern Ontario, Canada (Hervet et al., 2014). This parasitoid species was previously only known from the Palearctic and parts of the Afrotropic regions where it has been reared from a number of Noctuidae species. In fact, it has been reported from 33 species in seven families (Yu et al., 2016), but some of these reports are considered doubtful (Hervet et al., 2014). Preliminary laboratory observations showed that this species was able to parasitize North American noctuid pests, which provided rationale for the study of C. vanessae as a potential control agent for cutworms (Chapter 3), and its mass-rearing in the lab for potential future redistribution programs. For this study, nearly all Lepidoptera species were reared on an artificial diet known as McMorran diet (McMorran, 1965), using the recipe modified by Grisdale (1973). This artificial diet has been used by previous authors to rear a number of Lepidoptera species but it has not previously been used for the rearing of most of the species reared in the Chapter 3 study. Finding a suitable diet for the rearing of insects in the laboratory can greatly ease and enhance investigations of a species. For this reason, I report the species that I was able to rear on McMorran diet and species that are mentioned to have been reared on this diet in the literature (Chapter 4). The McMorran diet may not be equally suitable for all species, even if they complete development; e.g., some individuals had deformed wings, which can be indicative of a 5 Chapter 1 nutritionally unbalanced diet (Morris, 1967). Unbalanced diet for a caterpillar can affect the fitness of its parasitoid (Bloem et al., 1990; Pimentel, 1966). Thus, the diet used may confound parasitoid performance on different host species. One of the most important and often limiting elements in the diet of phytophagous insects is nitrogen (Harrison et al., 2012). To investigate how an unbalanced diet affects the fitness of a caterpillar and that of its parasitoid, in Chapter 5 I investigated the fitness of a caterpillar, Trichoplusia ni, and that of its parasitoid, C. vanessae, when the host was reared on artificial diets containing different concentrations of protein. Except for the General Introduction and General Discussion (Chapters 1 & 6), each chapter is written as a stand-alone manuscript intended to be published, or already published, as individual articles. The tables in Chapter 1 are published in Gavloski et al. (2013), Chapter 2 is published as Hervet et al. (2014), and Chapter 4 is published as Hervet et al. (2016b). An appendix (Appendix 1) summarizes some observations of the life history of C. vanessae that have not previously been reported. 6 Table 1.1. Parasitoids of army cutworm, Euxoa auxiliaris, darksided cutworm, Euxoa messoria, and redbacked cutworm, Euxoa ochrogaster (published in Gavloski et al., 2013). Parasitoid species Known distribution in North Euxoa auxiliaris Euxoa messoria Euxoa ochrogaster America INSECTA Diptera Bombyliidae Poecilanthrax halcyon (Say) Canada: SK (King et al., 1928) Poecilanthrax willistoni (Coquillet) Canada: SK (King et al., 1928) Villa alternata (Say) Canada: AB, ON, SK (Brooks, 1952) Villa fulviana (Say) Canada: SK (King et al., 1928) Villa lateralis (Say) Canada: SK (King et al., 1928) Muscidae Muscina stabulans (Fallén) Worldwide (Cheng, 1977) Sarcophagidae Sarcophaga cimbicis Townsend Canada: AB, MB, NS, ON, QC, (Crumb, 1929; SK; USA: Throughout Dahlem et al., 1996) Tachinidae Aphria ocypterata Townsend Canada: AB, BC, MB, NB, ON, (Arnaud, 1978; QC, SK, YT; USA: WA to ME, Cheng, 1977, 1981; South to CA, NM and VA O'Hara et al., 2004) Bonnetia comta (Fallén) Canada: AB, BC, MB, NB, NS, (Arnaud, 1978; Capinera, (Arnaud, 1978; (Arnaud, 1978; King et al., NT, ON, PE, QC, SK, YT; 2001; O'Hara et al., 2004) Cheng, 1977, 1981) 1928; O'Hara et al., 2004) Mexico: Throughout; USA: AK, throughout continental USA Chapter 1 7 Chetogena claripennis (Macquart) Canada: AK to ON; but (Arnaud, 1978; Capinera, probably a mixture with 2001; O'Hara et al., 2004) edwardsii, and QC; Throughout USA and Mexico Chetogena edwardsii (Williston) Distribution confused with (Arnaud, 1978; (Arnaud, 1978; O'Hara et (probably) that of C. claripennis O'Hara et al., 2004) al., 2004) Gonia aldrichi Tothill Canada: BC to NB; USA: (O'Hara et al., (Arnaud, 1978; King et al., South to CA and VA 2004) 1928) Gonia breviforceps Tothill Canada: BC to QC; USA: MT, (Arnaud, 1978; O'Hara et South to CA and AZ, also MI al., 2004) and KS Gonia fuscicollis Tothill Canada: MB, SK; USA: South (Arnaud, 1978; King et al., to NE and TN, also MD and 1928; O'Hara et al., 2004) IN Gonia spp. (Arnaud, 1978) Panzeria spp. (Arnaud, 1978; Capinera, 2001) Peleteria rubescens Robineau-Desvoidy (Arnaud, 1978) Peleteria texensis Curran USA only (Arnaud, 1978; O'Hara et al., 2004) Periscepsia cinerosa (Coquillett) Canada: BC to MT, also AB (Arnaud, 1978; Capinera, and YT; USA: South to CA, 2001) AZ, CO Periscepsia helymus (Walker) Canada: BC to NS, also NU (Arnaud, 1978; Capinera, (Arnaud, 1978; O'Hara et and YT; USA: South to CA, 2001; O'Hara et al., 2004) al., 2004) NM, KS, CT and AK Periscepsia laevigata (Wulp) Canada: AK to NU; also BC, (Arnaud, 1978; O'Hara et NB, NL, NS, and QC; Mexico ; al., 2004) USA: South to CA, TX and VA; Chapter 1 8 Tachina algens Wiedemann Canada: BC; NT to NL, NS (Arnaud, 1978; and QC; USA: South to CA, O'Hara et al., 2004) AZ, NM, MA and AK Winthemia deilephilae (Osten Sacken) Canada: ON (Cheng, 1977) Winthemia rufopicta (Bigot) Canada: ON (Cheng, 1977) Winthemia quadripustulata Fabricius Canada: NT, YT; BC to NS; (Arnaud, 1978; O'Hara et USA: South to CA, AZ, KS and al., 2004) NJ, also WV and VA Hymenoptera Braconidae Bracon erucarum (Cushman) USA: AZ, CA, CO, ID, MT, OK, (Yu et al., 2016) OR, UT, WY Chelonus insularis Cresson Mexico; USA: AR, AZ, CA, CO, (Capinera, 2001; Krombein FL, GA, HI, IL, KS, LA, MO, et al., 1979) MS, NM, OK, SC, TX, UT Cotesia acronyctae (Riley) Canada: AB, ON, SK; USA: CA, (Yu et al., 2016) CO, CT, IA, IL, IN, MA, MD, ME, MO, NH, NJ, OH Cotesia griffini (Viereck) Canada: AB, NB, QC; USA: (Krombein et al., 1979; (Schaaf, 1972; Yu et al., AR, FL, KS, MA, NY, OK, SC, Soteres et al., 1984; Yu et 2016) SD, TX, WA al., 2016) Cotesia laeviceps (Ashmead) Canada: AB, BC, MB, NB, ON, (Krombein et al., 1979; (Cheng, 1977, (Krombein et al., 1979; QC, SK; USA: CA, CO, CT, GA, Strickland, 1923; Yu et al., 1981; Yu et al., Strickland, 1923; Yu et al., IA, IL, MO, NM, NY, UT 2016) 2016) 2016) Cotesia vanessae (Reinhard) Canada: AB, ON This study (see Chapter 3) This study (see This study (see Chapter 3) Chapter 3) Glyptapanteles alticola (Ashmead) Canada: BC, NB; USA: AK, CA, CO, ID, ME, NH, OR, UT Chapter 1 9 Apatia truncator (Say) Canada: AB, BC, MB, NB, NS, (Capinera, 2001; Krombein ON, QC, SK; Mexico; USA: et al., 1979) Throughout Canada: AB, SK; North of (Capinera, 2001; Krombein Macrocentrus incompletus Muesebeck Mexico; USA: AZ, CA, CO, KS, et al., 1979; Yu et al., 2016) MT, NE, NM, OK, OR, SD, UT, WY Meteorus dimidiatus (Cresson) Canada: AB, BC, MB, NS, ON; (Strickland, 1923; Yu et al., (Strickland, 1923) USA: AK, AZ, CA, CO, DC, IL, 2016) KS, MA, ME, MI, MN, MO, NH, NJ, NY, OH, OR, PA, UT, VT, VA, WA Meteorus pendulus (Müller) Canada: BC, NB, NS, ON, QC; (Cheng, 1977, USA: AK, CA, CO, CT, DC, IL, 1981; Krombein et KS, MA, ME, MI, MO, NC, al., 1979; Yu et al., NH, NJ, NY, OR, PA, RI, TX, 2016) VA, VT, WV Meteorus rubens (Nees) Canada: AB, BC, MB, NB, NT, (Cheng, 1977; Krombein et (Cheng, 1977; (Schaaf, 1972; Yu et al., NS, ON, QC, SK; Mexico; al., 1979; Soteres et al., Krombein et al., 2016) USA: Throughout 1984; Yu et al., 2016) 1979; Yu et al., 2016) Microplitis feltiae Muesebeck USA: AL, AZ, CA, CO, ID, IL, (Capinera, 2001; Krombein IN, KS, LA, MO, ND, OK, SC, et al., 1979); (Yu et al., TN, TX, WA 2012)1 Microplitis kewleyi Muesebeck Canada: AB, MB, NB, NL, NS, (Schaaf, 1972; Yu et al., ON, PE, QC; USA: CA, DC, IA, 2016) MD, ME, MI, NJ, NY, WI Microplitis melianae Viereck Canada: AB, ON; USA: IA, IL, (Capinera, 2001; Krombein KS, MI, MN, NY, OH, OK, TN, et al., 1979) TX Chapter 1 10 Protapanteles militaris (Walsh) Canada: MB, NB, ON, QC; (Krombein et al., 1979; Yu (Cheng, 1977; Yu Mexico; USA: AR, AZ, CA, CT, et al., 2016) et al., 2016) DC, FL, HI, IA, IL, IN, KS, LA, MA, MD, MI, MN, MO, NJ, NM, NY, OK, TN, TX, VA Rogas sp. (Capinera, 2001) Encyrtidae Copidosoma bakeri (Howard) Canada: AB, NB, ON, SK; (Byers et al., 1993; Noyes, (Cheng, 1977; (Cheng, 1977; Noyes, USA: AZ, CO, KS, MA, MT, 2011; Yu et al., 2016) Noyes, 2011; Peck, 2011; Peck, 1963; Schaaf, NM, UT 1963; Yu et al., 1972; Yu et al., 2016) 2016) Ichneumonidae Arenetra canadensis Cresson Canada: AB, NB, ON, SK; (Yu et al., 2016) USA: CA, CO, IA, ID, IL, KS, MI, MT, NM, OR, SD, UT, WA, WI, WY Arenetra fumipennis Townes Canada: AB; USA: CO, MT, (Yu et al., 2016) ND Arenetra rufipes Cresson Canada: AB, BC, MB, NL, NS, (Cheng, 1977; Yu (Yu et al., 2016) ON, QC, SK; USA: AK, AZ, CA, et al., 2016) CO, ID, KS, MA, ME, MI, MN, MT, ND, NH, NM, NY, OR, PA, SD, WA, WY Campoletis atkinsoni (Viereck) Canada: AB, MB, SK; USA: ID, (Schaaf, 1972; Yu et al., OR, WA 2016) Campoletis australis (Viereck) Canada: AB; USA: NM (Schaaf, 1972; Yu et al., 2016) Campoletis flavicincta (Ashmead) Canada: AB, BC, NB, ON; (Capinera, 2001) (Cheng, 1977; Yu Mexico; USA: AR, CA, CO, DE, et al., 2016) FL, GA, IL, IN, KS, NC, OH, OK, SC, TN, VA Chapter 1 11 Campoletis sonorensis (Cameron) Canada: AB, BC, ON; Mexico; (Capinera, 2001; Murillo, This study2 This study2 USA: AL, AZ, CA, FL, GA, KY, 2008); (Yu et al., 2012)1 MS, NC, OK, SC, TN, TX, WA Diphyus apiculatus (Walkley) Canada: AB; USA: CO, ID, (Schaaf, 1972; Yu et al., MT, OR, WY 2016) Diphyus euxoae Heinrich Canada: AB, BC; USA: CA, CO, (Yu et al., 2016) (Cheng, 1977, (Schaaf, 1972; Yu et al., ME, NJ, NM 1981; Yu et al., 2016) 2016) Diphyus nuncius (Cresson) Canada: AB, BC; USA: CA, NJ, (Capinera, 2001; Yu et al., NY, UT 2016) Diphyus subfuscus (Cresson) Canada: AB, BC; USA: CA, CO, (Strickland, 1923; Yu et al., (Strickland, 1923; Yu et NV, NJ, NY 2016) al., 2016) Enicospilus sp. Canada: ON (Cheng, 1977) Erigorgus ambiguus (Norton) Canada: AB, MB, NB, ON, QC, (Schaaf, 1972; Yu et al., SK; USA: AR, AZ, CA, CO, KS, 2016) KY, MA, ME, MI, MN, MO, MS, MT, NC, ND, NH, NM, NY, OH, PA, SD, TX Eutanyacra suturalis (Say) Canada: AB, BC, MN, NB, NL, (Cheng, 1977; Yu (Schaaf, 1972; Yu et al., NU, ON, QC, SK; Mexico; et al., 2016) 2016) USA: CO, DE, GA, IL, KS, MA, NH, NJ, NM, NY, PA, UT, VA Exetastes lasius Cushman USA: CA, KS, NE, WY (Capinera, 2001; Yu et al., 2016) Exetastes obscurus Cresson Canada: AB, MB; Mexico; (Schaaf, 1972; Yu et al., USA: AL, CO, CT, DC, GA, ID, 2016) IL, KS, MD, MA, MI, MO, NC, NH, NJ, NY, OH, OR, PA, SC, TX, VA, WV Ichneumon longulus Cresson Canada: AB, BC; USA: AZ, CA, (Yu et al., 2016) CO, GA, IL, KS, NJ, NM, UT Chapter 1 12 Netelia sp. Canada: AB (Schaaf, 1972) Spilichneumon inconstans (Cresson) Canada: AB, BC, NL, NT, QC, (Yu et al., 2016) YT; USA: CA, CO, CT, IL, MA, ME, MI, NJ, NY, VA Spilichneumon superbus (Provancher) Canada: AB, BC, MB, NL, NS, (Capinera, 2001; Yu et al., (Cheng, 1977, (Krombein et al., 1979; NT, ON, QC, SK; USA: CO, HI, 2016) 1981; Krombein et Schaaf, 1972; Yu et al., TX al., 1979; Yu et al., 2016) 2016) ADENOPHOREA Mermithida Mermithidae Agamermis sp. (probably) Canada: AB (Schaaf, 1972) 1 Information previously appeared in Yu et al. (2012), but not anymore in Yu et al. (2016). 2 Campoletis sonorensis (GenBank accession number: KF640230) originally reared from Chrysodeixis includens and Trichoplusia ni collected in southwestern Ontario and maintained at the University of Windsor before being transferred to us. The three hosts reported here were exposed to C. sonorensis using a protocol identical to that described in Chapter 3. Chapter 1 13 Table 1.2. Predators of army cutworm, Euxoa auxiliaris, darksided cutworm, Euxoa messoria, and redbacked cutworm, Euxoa ochrogaster (published in Gavloski et al., 2013). Predator species Known distribution Euxoa auxiliaris Euxoa messoria Euxoa ochrogaster in North America ARACHNIDA Opiliones Phalangiidae (kill moths) Leiobunum vittatum (Say) Canada (Cheng, 1984) Phalangium opilio (Linnaeus) Canada (Cheng, 1984) Odiellus pictus (Wood) Canada (Cheng, 1984) Sclerosomatidae Hadrobunus maculosus (Wood) Canada (Cheng, 1984) AVES (kill karvae) Charadriiformes Laridae Leucophaeus pipixcan (Wagler) Throughout North (King et al., 1928) America Passeriformes Icteridae Quiscalus quiscalus (L.) Canada: ON (Cheng, 1984) Sturnidae Sturnis vulgaris L. Canada: ON (Cheng, 1984) INSECTA Coleoptera Carabidae (kill larvae and eggs) Agonum cupreum Dejean Canada, USA (Frank, 1971; Larochelle, 1990) Agonum placidum (Say) Canada, USA (Frank, 1971; Larochelle, 1990) Amara apricaria (Paykull) Canada, USA (Frank, 1971; Larochelle, 1990) Amara avida (Say) Canada (Frank, 1971; Larochelle, 1990) Amara ellipsis (Casey) Canada (Frank, 1971; Larochelle, 1990) Chapter 1 14 Amara latior (Kirby) Canada, USA (Frank, 1971; Larochelle, 1990) Amara patruelis Dejean Canada (Frank, 1971; Larochelle, 1990) Amara quenseli (Schönherr) Canada (Frank, 1971; Larochelle, 1990) Amara torrida (Panzer) Canada (Frank, 1971; Larochelle, 1990) Bembidion bimaculatum (Kirby) Canada (Frank, 1971; Larochelle, 1990) Bembidion canadianum Casey Canada (Frank, 1971; Larochelle, 1990) Bembidion mutatum Gemminger & Harold Canada (Frank, 1971; Larochelle, 1990) Bembidion nitidum (Kirby) Canada (Frank, 1971; Larochelle, 1990) Bembidion obscurellum (Motschulsky) Canada, USA (Frank, 1971; Larochelle, 1990) Bembidion quadrimaculatum oppositum Say Canada, USA (Frank, 1971; Larochelle, 1990) Bembidion rupicola (Kirby) Canada (Frank, 1971; Larochelle, 1990) Bembidion versicolor (LeConte) Canada (Frank, 1971; Larochelle, 1990) Calosoma calidum (Fabricius) Canada, USA (King et al., 1928; (King et al., 1928; Larochelle, Larochelle, 1990) 1990) Carabus serratus Say Canada, USA (Frank, 1971; Larochelle, 1990) Carabus taedatus Fabricius Canada, USA (Frank, 1971; Larochelle, 1990) Harpalus amputatus Say Canada, USA (Frank, 1971; Larochelle, 1990) Harpalus caliginosus (Fabricius) Canada, USA (Cheng, 1984; Larochelle, 1990) Harpalus funerarius Csiki Canada (Frank, 1971; Larochelle, 1990) Poecilus lucublandus (Say) Canada, USA (Frank, 1971; Larochelle, 1990) Pterostichus adstrictus Eschscholtz Canada (Frank, 1971; Larochelle, 1990) Trichocellus cognatus (Gyllenhal) Canada (Frank, 1971; Larochelle, 1990) Staphylinidae (kill eggs) Leptacinus batychrus (Gyllenhal) Canada (Frank, 1971; Larochelle, 1990) Philonthus occidentalis Horn Canada (Frank, 1971; Larochelle, 1990) Tachyporus sp. Canada (Frank, 1971; Larochelle, 1990) Hymenoptera Formicidae (kill eggs) Lasius niger neoniger Emery Temperate region (King et al., 1928; Wang et al., of North Americ a 1995) Chapter 1 15 MAMMALIA Carnivora Ursidae (kill moths) Ursus arctos horribilis Ord USA: WY, ID, MT (French et al., 1994) (Yellowstone National Park area) Chapter 1 16 Chapter 1 Table 1.3. Pathogens of army cutworm, Euxoa auxiliaris, darksided cutworm, Euxoa messoria, and redbacked cutworm, Euxoa ochrogaster (published in Gavloski et al., 2013). Pathogen species Euxoa auxiliaris Euxoa Euxoa messoria ochrogaster (BACTERIA) BETAPROTEOBACTERIA Burkholderiales Alcaligenaceae Achromobacter spp.2 (Cheng, 1984) BACILLI Bacillales Bacillaceae Bacillus cereus Frankland & (Cheng, 1984) Frankland (Cheng, Bacillus sphaericus Neide2,3 1984)3 Streptococcus faecalis (Andrewes (Cheng, 1984) & Horder)2 GAMMAPROTEOBACTERIA Enterobacteriales Enterobacteriaceae Enterobacter cloacae (Jordan)2 (Cheng, 1984) Enterobacter aerogenes (Kruse)1 (Cheng, 1984) Klebsiella pneumonia (Schroeter)2 (Cheng, 1984) Pseudomonadales Pseudomonadaceae Pseudomonas fluorescens (Cheng, 1984) (Migula)1 (FUNGI) DIHAPLOPHASEA Dissociodihaplophasida Nosematidae Nosema sp. (Cheng, 1984) SORDARIOMYCETES Hypocreales Sorosporella uvella (Krass.) (Cheng, 1984) (VIRUSES) Baculoviridae Granulosis virus (Jackson et al., (Cheng, 1984) (Cheng, 1985) 1984)3 Nuclear polyhedrosis virus (Jackson et al., (Cheng, 1984) (Cheng, 1985; McCarthy 1984)3 et al., 1975) Nonoccluded nuclear (Sutter, 1973) polyhedrosis virus Poxviridae Entomopoxvirus (Sutter, 1973) 1 Detected in haemocoel, potential pathogens 2 Detected in haemocoel, unlikely pathogens 3 Information that does not appear in Gavloski et al. (2013) 17 17 Chapter 2 CHAPTER 2: IDENTIFICATION AND BIOLOGY OF COTESIA VANESSAE. FIRST REPORT OF THIS SPECIES IN NORTH AMERICA (Published as: Hervet, V. A. D., Murillo, H., Fernández-Triana, J., Shaw, M. R., Laird, R. A., & Floate, K. D. (2014). First Report of Cotesia vanessae (Hymenoptera: Braconidae) in North America. The Canadian Entomologist, 146(5), 560-566) 2.1. Abstract This chapter reports for the first time the occurrence of the well-known Eurasian and north African parasitoid Cotesia vanessae (Reinhard) (Hymenoptera: Braconidae) in North America. Specimens were reared from Chrysodeixis chalcites (Esper) and Trichoplusia ni (Hübner) (Lepidoptera: Noctuidae) recovered from several locations in southwestern Ontario, and detected by DNA sequencing from one Autographa californica (Speyer) (Lepidoptera: Noctuidae) in southern Alberta, Canada. 2.2. Introduction Cotesia is a genus of braconid wasps that are parasitoids of Lepidoptera larvae. Several species are of interest as biological control agents of pest caterpillars. For example, Cotesia urabae Austin & Allen was released into New Zealand in 2011 for the biological control of the eucalyptus pest Uraba lugens Walker (Lepidoptera: Nolidae) (Avila et al., 2011). Cotesia flavipes Cameron has been studied for many years and has been introduced in many countries as a biological control agent for the sugarcane borer Diatraea saccharalis (Fabricius) (Lepidoptera: Crambidae) on cereal crops (Poaceae) (Jiang et al., 2004; Overholt et al., 1997). Cotesia rubecula (Marshall) has been released in the United States as a natural enemy of Pieris rapae (Linnaeus) (Lepidoptera: Pieridae) on cabbage, Brassica spp. (Brassicaceae) (van Driesche et al., 2002). 18 Chapter 2 In the most recent checklist of Cotesia spp. in Canada and Alaska, Fernández-Triana (2010) reported 55 species (including four undescribed species) with the expectation that many more species in the genus remained unreported. The current work adds to this list the first report of Cotesia vanessae (Reinhard) in Canada and, more broadly, North America (Whitfield, 1995). Cotesia vanessae is a gregarious endoparasitoid that primarily attacks species of Vanessini (Nymphalidae) and certain Noctuidae. It has also been reported from Notodontidae, Lasiocampidae, Crambidae, and Pterophoridae (Yu et al., 2012), but these reports are considered doubtful. Host records from the Old World here consider reliable include the vanessines Vanessa cardui (Linnaeus), Vanessa atalanta (Linnaeus) and Aglais urticae (Linnaeus), and the noctuids Actebia praecox (Linnaeus) and Mythimna litoralis (Curtis) (specimens in the British Museum of Natural History and the National Museum of Scotland). Nixon (1974) recorded Spodoptera exigua (Hübner) as a host, but in the present study C. vanessae couldn’t be reared on this species in laboratory experiments (see Chapter 3). In Britain, C. vanessae overwinters within overwintering noctuid host larvae because its vanessine hosts are available as larvae only in the summer and the parasitoid adults always emerge from the cocoons in the year of their formation (Nixon, 1974). Genetic analyses, as well as morphological and biological observations, show the existence of populations of C. vanessae in Ontario and Alberta, Canada. This parasitoid species has previously been reported throughout Europe and scattered locations in Asia and North Africa (Stefanescu et al., 2012; Yu et al., 2012). European populations of C. vanessae may be bisexual or, in the Mediterranean region, comprise only thelytokous females (Stefanescu et al., 2012). 19 Chapter 2 2.3. Collections 2.3.1. From Ontario Cotesia vanessae was first reared from the tomato looper, Chrysodeixis chalcites and cabbage looper, Trichoplusia ni, collected in 2009 in southwestern Ontario. C. chalcites has already been reported as a host of C. vanessae (Messelink, 2002), but T. ni is a new host record for C. vanessae. For Chrysodeixis chalcites, parasitized loopers were collected from tomato plants, Solanum lycopersicum Linnaeus (Solanaceae), in fields around the cities of Harrow and Leamington in the summers of 2009 and 2010 (42°05'07.25"N, 82°36'08.65"W; 42°02'11.54"N, 82°57'03.78"W; 42°02'36.45"N, 82°38'05.60"W; 42°05'00.78"N, 82°37'07.26"W; 42°02'00.82"N, 82°53'58.49"W). Parasitized C. chalcites were also collected from tomato, pepper, Capsicum annuum Linnaeus (Solanaceae), and cucumbers, Cucumis sativus Linnaeus (Cucurbitaceae), in greenhouses around Leamington and Chatham-Kent in the fall of 2009, and in the late summer and fall of 2010. For T. ni, a parasitized larva was collected from a broccoli plant in a field west of Leamington in August 2009 (42°02'36.45"N, 82°38'05.60"W) and from tomato plants in fields around Harrow and Chatham-Kent in the summer of 2010 (42°02'11.54"N, 82°57'03.78"W; 42°23'26.93"N, 82°09'27.13"W) (Fig. 2.1). Parasitoids from these original field collections initially were reared at the Sunrite Greenhouses Biocontrol laboratory, Ontario (by Henry Murillo, University of Windsor), and subsequently at the Lethbridge Research Centre, Alberta (by VH). Colonies were maintained on T. ni purchased from the Insect Production Services of the Great Lakes Forestry Centre, Sault Ste. Marie, Ontario, which were reared on McMorran diet (McMorran, 1965) with Grisdale’s modification (Grisdale, 1973). Only females have been observed in our laboratory colony, reproducing by thelytokous parthenogenesis over multiple generations. From a subsample of 26 T. ni (these included the ten T. ni previously mentioned here in Hervet et al., 2014), each exposed to a single C. vanessae for 48 hours, an average of 95 adult parasitoids 20 Chapter 2 emerged per host (range: 5-176) (see Fig. 3.5 in Chapter 3). From another subsample of 31 T. ni, each oviposited into only once by a C. vanessae, an average of 92 adult parasitoids emerged per host (range: 16-199) (see Fig. 5.9 in Chapter 5). These hosts were parasitized at their early 4th instar and reared on the diet previously mentioned. To identify the parasitoid species, CO1 sequences were obtained from laboratory colony specimens using primers and general methods as described in Hebert et al. (2003). All the sequences were identical, one of which (KF640231) was deposited in GenBank (http://www.ncbi.nlm.nih.gov/genbank/). This sequence was compared with existing sequences from GenBank, using phylogenetic analyses that identified these specimens as Cotesia vanessae (Fig. 2.2). To characterize the morphology of this species, photographs of adult and immature C. vanessae are provided in Figures 2.3 and 2.4. There are many similar Cotesia species, thus these pictures are not meant to be used to identify an unknown specimen. Reinhard’s descriptions of C. vanessae (Reinhard, 1880, 1881) and Nixon’s (1974) diagnosis correspond with our observations. However, the Ontario specimens tend to have lighter metafemur, metatibia, and metasoma (especially the laterotergites) compared to the specimens that were examined (by Mark R. Shaw) from northwestern Europe. Specimens from Libya and Ethiopia (perhaps really Eritrea) also have been reported to be lighter in color than European specimens (Nixon, 1974; Papp, 1987). Nixon (1974) also drew attention to minor differences, that he considered to be of little significance, between European series reared from vanessines and noctuids – presumably merely season- and/or host-related. 2.3.2. From Alberta A DNA sequence for C. vanessae, but not specimens, was recovered from a final instar larva of alfalfa looper, Autographa californica (Speyer) (Lepidoptera: Noctuidae), that had also produced a brood of a different species of Cotesia. The larva was one of about thirty 21 Chapter 2 A. californica collected for an unrelated study on 9 and 10 July 2012 from three canola fields; i.e., 12 km NE of Claresholm (50°04’45”N, 113°26’21”W), 17 km NE of Claresholm (50°07’23”N, 113°26’21”W), 30 km E of Calgary (50°57’27”N, 113°31’42”W). Larvae were held indoors in a Plexiglas cage (30 x 30 x 30 cm) at 22 °C and fed canola leaves, Brassica napus Linnaeus (Brassicaceae) for three days after which an unidentified gregarious species of Cotesia emerged from most individuals. Based on morphology and CO1 sequences (KF640233), the parasitoid was not C. vanessae. However, in sequencing parasitized caterpillars to verify their identity, a CO1 sequence (KF640229) of one of these hosts showed to be identical to that for C. vanessae from Ontario (Fig. 2.2). It is speculated that this A. californica was parasitized by both the unknown Cotesia sp. and by C. vanessae, but only the former emerged from the host. Cases of caterpillars being parasitized by two Cotesia species and progeny of both species successfully developing within the host have previously been documented (Ngi-Song et al., 2001). 2.4. Current known range Cotesia vanessae is widely distributed throughout the Palearctic (Fig. 2.5). Nixon (1974) recorded it from Eritrea (Asmara), Libya (Benghazi), Turkey, Italy, and “northwest Europe”. Stefanescu et al. (2012) reported it from locations in central-western Morocco and north-east Spain. Specimens in the Natural Museum of Scotland also include records from: Israel (Negev Mountains), Bulgaria (East Rhodope Mountains), Greece (N. Peloponnese), Crete, Canary Islands (Lanzarote, Fuerteventura), Spain (Girona, Lleida, Barcelona, Córdoba, Aragon), France (Charente), Finland (Åland), and most of the counties in the southern half of England. Additional records (Fig. 2.5) are from Yu et al. (2012). 22 Chapter 2 2.5. Conclusion The first record of Cotesia vanessae in North America is here documented by morphological, biological and genetic evidence. Where, when and how this parasitoid may have been introduced to North America is unknown. It may have been introduced into North America while developing inside a host in a shipment of imported plants. Recovery of this parasitoid from three pest species thus far (Chrysodeixis chalcites, Trichoplusia ni and Autographa californica) warrants further study of C. vanessae as a potential biological control agent against noctuid crop pest caterpillars. However, this parasitoid is also reported to parasitize nymphalid butterflies such that non-pest species are likely to be at risk. Benzon et al. (Benson et al., 2003) showed that the deliberate introduction of Cotesia rubecula (Marshall) for the biological control of pestiferous Pieris butterflies had a negative impact on a native non-pest congener. Hence, monitoring the establishment and spread of C. vanessae in North America and its possible colonization of native vanessines and noctuids is advised, before any deliberate manipulations in the interests of crop protection are attempted. 23 Chapter 2 A B 5% 192 30% 104 4% 25% 31 20%3% 15% 2% 61 10% 1% 5% 71 16 39 9 48 73 0 11 4 0 0 0 15 0% 0% 69 0 72 0 176 0 0 0 June July August September October June July August September October CC.. cChhallciites - Tomaattooeess TT.. nii - Broccoolili Tomatoes Cucumbers Peppers C D 20% 5% 544 6 24 15% 4% 3% 10% 359 69 40 2% 5% 137 31 1% 585 4 229 457 0 7 0 0 0 279 0 257 0 203 18 0% 0% June July August September May June July August September C. Cchalciitess -- T Toommaatotoees s T.. ni - Tomattooeess C. Cchalcitess -- TToommaatotoees s C. cChalcites -- CCuuccuummbbeersr s T. nii - - C Cuuccuummbberesr s Figure 2.1. Parasitism (%) of Chrysodeixis chalcites and Trichoplusia ni by Cotesia vanessae collected from fields and commercial greenhouses in southwestern Ontario, Canada in 2009 and 2010. The numbers represent the total number of caterpillars collected each month on the indicated plants: A. 2009 in fields; B. 2009 in greenhouses; C. 2010 in fields; D. 2010 in greenhouses. 24 Chapter 2 C. pieridis (JF963155) C. glomerata (JQ240173) C. sibyllarum (JF963159) C. rubecula (JQ396720) C. urabae (JQ396721) C. salebrosa (JF963157) C. sesamiae (JQ396743) C. vestalis (AY934820)1 88 91 C. flavipes (JQ396717) 99 C. nonagriae (JQ396713) C. tetrica (JF963163) 78 C. bignellii (AY333892) 85 Cotesia sp. jft08 (JN639962) Cotesia sp. jft02 (JN639952) C. yakutatensis (GU141139) C. plathypenae (JQ849202) C. xylina (JN639978) C. vanessae (JF963165) 100 C. vanessae – Ontario (KF643231) 75 C . vanessae – Alberta (KF640229) 0 .01 1Although the name Cotesia plutellae (Kurdjumov) is used for this sequence in GenBank, the valid name for this species is Cotesia vestalis (Haliday) (see Shaw (2003)). Figure 2.2. Neighbour-joining tree, K2P distance model for CO1 sequences for Cotesia species generated using Mega 5.2. Sequences obtained from GenBank. Bootstrap values < 70 are not reported. 25 Chapter 2 A B 1 mm 1 mm C D 1 mm 1 mm E F 500 µm 250 µm Figure 2.3. Cotesia vanessae adults (females): A. live, dorsal side; B. live, ventral side; C. live, emerging from pupal exuvia, dorsal side; D. dead, pointed, left lateral side; E. dead, upper surface of right wings; F. dead, dorsal side of abdomen. 26 Chapter 2 A 3 mm B 1 mm D 1 mm E 1 mm C F 5 mm 1 mm Figure 2.4. Cotesia vanessae immature stages: A. larvae (prepupae) erupting from a T. ni larva and starting to spin cocoons; B. prepupa just emerged from a Noctuidae host; C. cocoon mass of about 80 cocoons around dead host; D. one-day-old pupa, left lateral side, with cast larval exuvia and meconium (black mass at posterior end); E. three-day-old pupa, dorsolateral side; F. five-day-old pupa, ventral side. 27 Chapter 2 Figure 2.5. Reported distribution of Cotesia vanessae. The North American reports are from this study. The Old World distribution includes unpublished data (specimens in the National Museums of Scotland). Figure generated using SimpleMappr. 28 Chapter 3 CHAPTER 3: EVALUATION OF THE HOST RANGE AND FITNESS OF THE PARASITOID COTESIA VANESSAE ON NORTH AMERICAN LEPIDOPTERA, PARTICULARLY PESTIFEROUS NOCTUIDAE “I have a friend who's an artist and has sometimes taken a view which I don't agree with very well. He'll hold up a flower and say "Look how beautiful it is," and I'll agree, I think. And he says– "you see, I as an artist can see how beautiful this is, but you as a scientist, oh, take this all apart and it becomes a dull thing." And I think that he's kind of nutty. First of all, the beauty that he sees is available to other people and to me, too, I believe, although I may not be quite as refined aesthetically as he is; but I can appreciate the beauty of a flower. At the same time I see much more about the flower than he sees. I can imagine the cells in there, the complicated actions inside which also have a beauty. I mean it's not just beauty at this dimension of one centimeter, there is also beauty at smaller dimensions, the inner structure. Also the processes, the fact that the colors in the flower evolved in order to attract insects to pollinate it is interesting–it means that insects can see the color. It adds a question: Does this aesthetic sense also exist in the lower forms? Why is it aesthetic? All kinds of interesting questions which a science knowledge only adds to the excitement and mystery and the awe of a flower.” Richard P. Feynman (1981). The pleasure of finding things out. 3.1. Abstract Noctuidae are major pests of cultivated plants worldwide, including in North America. Natural enemies can help reduce damage, both in greenhouses and in fields. The occurrence of a parthenogenetic population of the gregarious braconid parasitoid Cotesia vanessae in Canada is evidenced by Chapter 2. This species was already known from the Palearctic region where it is reported to use Noctuidae and Nymphalini as hosts. The potential North American host range of this species was assessed in a laboratory experiment by testing its ability to complete its development using Noctuidae, Nymphalidae, and Lepidoptera from other families. Of the 47 species tested, I found 30 Noctuidae and one Nymphalidae species not previously known as hosts, and confirmed three Noctuidae that were already known hosts of C. vanessae. For most of these host species, parasitoid fitness was investigated by measuring rate of successful parasitism, development time (in host, in cocoon, and total), brood size, individual parasitoid mass, and estimated brood mass. Noctuidae in the subfamily Plusiinae were found to generally be the best hosts. Species of similar host quality were usually, but 29 Chapter 3 not always, closely related. Thus, we can likely predict non-tested species from certain genera to be good hosts (e.g., Euxoa, Apamea) or non-hosts (e.g., Lacinipolia, Spodoptera). Evidence of scramble and contest competition among parasitoid larvae was also observed, manifested by a trade-off between parasitoid brood size and individual mass, and siblicidal behaviour, respectively. Finally, C. vanessae was also observed to cause indirect death of some parasitized Lepidoptera larvae, of up to 100% for some species, by suppressing their ability to pupate and to excrete food, even though the immune system of these individuals killed all parasitoid larvae at an early stage. The ability of C. vanessae to develop in different species of pestiferous Noctuidae suggests a potential role for this species as a biocontrol agent in pest control programs. 3.2. Introduction With about 25,000 described species, including 3,250 in America north of Mexico, the family Noctuidae is the most species-rich family of Lepidoptera, and it contains the greatest number of horticultural and agricultural pest species (Zahiri, 2012; Zhang, 1994). Many noctuid species – known by common names such as cutworms, armyworms, and semi- loopers – cause significant economic losses in North America and elsewhere. For example, between 1955 to 1975 in Idaho, Montana, South Dakota, Wyoming, Nebraska, Colorado, Kansas, and Oklahoma, the pale western cutworm, Agrotis orthogonia Morrison, caused an average annual loss of 17-million USD (the equivalent of ~90-million USD in 2016). During the same time period in Montana, South Dakota, Colorado, and Oklahoma, the army cutworm, Euxoa auxiliaris (Grote), caused a total annual average loss of nearly 8-million USD (~60- million USD in 2016) (USDA, 1977). Global losses due to cotton bollworm, Helicoverpa armigera (Hübner), are estimated to be 2-billion USD annually, in addition to over 1-billion USD spent on insecticide applications (Sharma et al., 2014). Over the last decade a noticeable increase in cutworm outbreaks in the agricultural region of the Canadian prairies has 30 Chapter 3 renewed interest in these pests, and biological control techniques are being investigated (WCCP, 2012, 2013). Cotesia (Hymenoptera: Braconidae) is a speciose genus of both gregarious and solitary parasitoids within the subfamily Microgastrinae. All microgastrines are koinobiont endoparasitoids of Lepidoptera larvae. Adult females lay small, hydropic eggs (= ostensibly yolk-free eggs that develop and expand within the host, enabled by an extraembryonic membrane that permits absorption of nutrients from the host’s haemolymph (Grbić et al. 1998; Harvey et al. 2013; King et al., 1969)). Larvae of many genera (e.g., Cotesia, Microplitis, and Protomicroplitis) feed essentially on the host’s haemolymph and fat tissues, and egress from the host’s sides prior to pupating. These characteristics, which are putatively apomorphic compared to solitary tissue feeding, allow these species to parasitize a wide range of host sizes, develop rapidly, and, for some species, utilize the moribund host as a body guard for their cocoons (Flanders, 1942; Harvey et al. 2008; Michel-Salzart et al., 2004; Shaw et al. 1991). Species in the genus Cotesia have been used in previous biological control programs; e.g., C. rubecula (Marshall) against Pieris rapae (Linnaeus) (Lepidoptera: Pieridae) (van Driesche et al., 2002), C. urabae Austin & Allen against Uraba lugens Walker (Lepidoptera: Nolidae) (Avila et al., 2011; Avila et al., 2013), C. flavipes (Cameron) against Chilo partellus (Swinhoe) (Lepidoptera: Crambidae) and its relatives (Cugala et al., 2001), and Cotesia marginiventris (Cresson) against Trichoplusia ni (Hübner) (Lepidoptera: Noctuidae) and other Noctuidae (Gillespie et al., 1999; Messelink, 2002). Only recently has the Palearctic species Cotesia vanessae (Reinhard) (Hymenoptera: Braconidae) been recognized as occurring in North America (Hervet et al., 2014). Because this species is known to parasitize species of Noctuidae and Nymphalidae in the western Palearctic region (Hervet et al., 2014; Nixon, 1974; Papp, 2011-2012; Stefanescu et al., 2012), it may help suppress pestiferous species in North America. 31 Chapter 3 Cotesia vanessae has many attributes that enhance its potential as a biocontrol agent. It is easy to mass rear because it usually parasitizes hosts as soon it is exposed to them, it can parasitize multiple hosts consecutively (e.g., 50 Euxoa ochrogaster (Guenée) (Lepidoptera: Noctuidae) larvae were parasitized by a single parasitoid in 20 minutes), it successfully parasitizes any host instar except the first, it is gregarious, it can be kept alive as adult usually up to a month, sometimes up to a month and half (using rearing conditions described below) and it has a facultative diapause that allows for continuous rearing in the laboratory. Further, maintaining thelytokous C. vanessae eliminates the problem of male- biased laboratory colonies and every individual can parasitize hosts. In the current study, C. vanessae host range and its fitness (parasitism rate, development time, brood size, brood mass, and individual parasitoid mass) was tested on possible host species to assess its potential use as a biological control agent against noctuid pests. Non-pest species and species in other families were also included to investigate the limit of its host range and potential non-target effects. Closely related species were tested to assess the robustness of a phylogenetic approach to help predict whether species closelyrelated to known hosts would also be hosts. Most species tested were pestiferous Noctuidae, partly because these were target species, but also because they represented the highest number of catches using UV light traps, and also because some of these species were commercially available. I also investigated the occurrence of scramble competition (i.e., “the finite resource is shared equally amongst the competitors so that the quantity of food per individual declines with increasing population density” (Den Berg et al., 2006)) and contest competition (i.e., “each successful competitor obtains all resources it requires for survival or reproduction; the remaining competitor being deprived of its resources, will not be able to function anymore” (Den Berg et al., 2006)) among parasitoid larvae within hosts. 32 Chapter 3 3.3. Materials and Methods 3.3.1. Insect collection The research was conducted at the Lethbridge Research and Development Centre, Agriculture and Agri-Food Canada (AAFC), Lethbridge, AB, Canada. The C. vanessae laboratory colony was initiated at the University of Windsor (Windsor, ON) and transferred to AAFC in 2011. Parasitoids originated from southern Ontario, where they were reared from larvae of tomato looper, Chrysodeixis chalcites (Esper), and cabbage looper, Trichoplusia ni (Lepidoptera: Noctuidae: Plusiinae), collected on vegetables in commercial greenhouses and fields in 2009 and 2010 (Hervet et al., 2014). In the laboratory, parasitoids were maintained on T. ni purchased from Insect Production Service (IPS), Natural Resources Canada, Sault Ste Marie, ON, Canada. Trichoplusia ni larvae were reared on an artificial diet, the McMorran diet with Grisdale’s modification (Grisdale, 1973; McMorran, 1965), also purchased from IPS. Species of Lepidoptera tested as potential hosts for C. vanessae were obtained using different methods. Adults were collected with two UV light traps in summer and fall of 2012 to 2014 in and near Lethbridge, Alberta (traps were usually at 49.6924 °N -112.8152 °E and 49.6953 °N -112.7674 °E) (Table 3.1). Eggs were obtained from gravid females, males were released. Other species were purchased as eggs from Benzon Research Inc., Carlisle, PA, USA (Table 3.1). To import species from the United States, Canadian Food Inspection Agency Import Permits (No. P-2011-04397, P-2013-02134, and P-2014-02394) were obtained. An additional ~50 species were collected by hand and sweep net as eggs or larvae, in and near Lethbridge, and near Magrath (Cardston County, AB) (49.4327 °N -112.8626 °E) (i.e., Noctuidae, Erebidae, Lasiocampidae, Plutellidae, Hesperidae, Nymphalidae, Pieridae, Papilionidae, and Tortricidae). Although these were tested as potential hosts of C. vanessae, many of them were either too diseased or too close to pupation to remain as larva long enough to support parasitoid development and were thus not included in the analyses. 33 Chapter 3 3.3.2. Insect identification The identity of the Lepidoptera species was determined based on the morphology of the adult and larval forms and confirmed using molecular identification. For morphological identification, I used taxonomic keys (Eichlin, 1975; Eichlin et al., 1978; Schmidt, 2015) and comparison with preserved adults and larvae in collections held at the Lethbridge Research and Development Centre. For molecular identification, mitochondrial cytochrome oxidase 1 (CO1) sequences were obtained from specimens using primers and general methods as described in Hebert et al. (2003). DNA extraction and amplification was done at AAFC Lethbridge; sequencing was done at the University of Calgary. Sequencher software was used to trim and align the sequences. The CO1 sequences thus obtained were compared with sequences deposited in GenBank (https://blast.ncbi.nlm.nih.gov/Blast.cgi?PAGE_TYPE=BlastSearch). Field-collected larvae may have been parasitized prior to collection. Therefore, parasitoids emerging in the laboratory were also identified with this molecular method if necessary to confirm the identity of C. vanessae. 3.3.3. Colony rearing Field-collected lepidopteran larvae were handled in one of two ways depending on their number. Species collected in low numbers were usually reared to the next generation to increase the number of larvae available, to produce parasitoid-free individuals, and reduce disease pressure. Species that were collected in large numbers (~50 individuals per species) were used directly in the experiment; i.e., Apamea sordens (Hufnagel), Dargida diffusa (Walker) (Lepidoptera: Noctuidae), Aglais milberti (Godart) (Lepidoptera: Nymphalidae), Malacosoma disstria Hübner (Lepidoptera: Lasiocampidae), and Pieris rapae. Between experiments, Lepidoptera of all stages were reared in Plexiglas cages (40 x 34 x 30 cm [length, width, height]). The cages had a 10 cm diameter screen on opposing sides for ventilation and 34 Chapter 3 on the front a 15 cm diameter hole to which a screen sleeve was attached for accessing the cage. Adult Lepidoptera were provided with water, crystal sucrose, and oviposition substrates: fine sand, cheese cloth, paraffin film, brown paper, and paper towel. When eggs were laid, the oviposition substrate was cut into small squares and placed in clean cages on top of blocks (22 ml) of McMorran diet. Sand was sieved to recover the eggs, and the eggs were placed in a shallow 20 cm diameter plastic lid in the middle of the cage. Blocks of McMorran diet were mainly placed along the sides and corners of the cages to ease caterpillar access and were replaced with fresh blocks every 5 to 7 days as needed. To slow evaporation, pieces of paraffin film (Parafilm®) (3 x 3 cm) were placed on the bottom and top of each diet block. Species that couldn’t be reared on McMorran diet were reared on their natural host plants (details in Chapter 4). Crumpled pieces of brown paper bag were placed in cages as shelter. Larvae were either used in experiments or used to generate a subsequent generation. In the latter case, pupae were placed in clean cages with paper towel for emerging moths to climb onto and provided with water, sucrose, and oviposition substrates. Both sexes were kept together to ensure that females were fertilized. Their eggs were handled as described above. Lepidoptera were reared in a growth cabinet at 12:12 h light:dark cycle, at constant 20 °C, and at 70 to 80% humidity to reduce evaporation of artificial diet and prevent desiccation of insect pupae. These conditions were monitored using a data logger (HOBO Pendant® UA-002-08) placed into an empty experimental cup and a thermo- and hygrometer (TempMinder® Remote Thermo-Hygro EMGR819LR(MRC800)) placed into an empty cage. The parasitoid colony of C. vanessae was reared in the same type of cages that was used for Lepidoptera. To increase their life span, adult parasitoids were maintained at 10 °C, 12:12 h light:dark cycle under low light intensity, with access to honey, water, and a piece of crumpled brown paper for shelter. The cage was moved to 20 °C for two hours to allow parasitoids to feed and drink each week. At the beginning of that time fresh drops of honey 35 Chapter 3 were placed on the upper corner of the cage where parasitoids spent the majority of their time and water was sprayed on the walls and ceiling of the cage. For offspring production, adult parasitoids (1 to 3 weeks post-emergence) were exposed to 4th instar Trichoplusia ni. To ensure that all larvae were parasitized, they were exposed individually to parasitoids inside of the parasitoid cage until one or two parasitism events were observed. Parasitized larvae were then transferred to clean cages (~20 larvae per cage) containing blocks of McMorran diet and reared in the same conditions as non-parasitized individuals. When individuals of a given Lepidoptera species were no longer required, they were typically exposed to several dozen parasitoids in cages (for 20 to 100 caterpillars per cage) to help regenerate the parasitoid colony. When placed into cages containing hosts, parasitoids didn’t seem to be able to locate them and the percent of parasitism was always very low (< 10%). However, when placed in direct contact with a Lepidoptera larva, and especially when dropped onto it, parasitism generally occurred instantly. Hence, to produce high in-cage percent parasitism, hosts were usually placed into a pile. Parasitoids were gathered in a vial with a mouth aspirator, then dropped onto the pile of caterpillars. To reduce incidence of diseases, all material (e.g., cages, experimental cups, and forceps) was thoroughly cleaned and disinfected with 70% bleach for five minutes between uses. 3.3.4. Host-range and host-fitness testing Cotesia vanessae host-range and fitness in different host species was tested using no-choice tests. Based on preliminary observations, 4th instars were selected to be parasitized as these appeared to yield the best parasitism results; i.e., compared to earlier instars, parasitoids egressed (= emerged as larvae from a host; term here used to prevent confusion with the “emergence” of adults from cocoons) from fully grown host larvae so they produced larger broods, parasitized hosts had higher survival rates, and they were more attractive to parasitoids – and compared to later instars, 4th instars displayed lower physical 36 Chapter 3 resistance and a higher percentage of successful parasitism. When too few 4th instars were available, other instars (usually 5th instar) of similar size were used. Because some species were larger than others, larvae parasitized were of different sizes between species. For Lepidoptera reared to the next generation in the laboratory, the goal was to use 30 to 35 larvae per species in the experiment. For some species, fewer individuals were used due to lack of availability. Field-collected larvae had a high mortality rate (from 51% with Aglais milberti, n = 49, to 100% with Malacosoma disstria, n = 60), due to diseases and field parasitism. Hence, for these species, 50 individuals were used whenever possible. The highest number of replicates used was 75 for Helicoverpa zea (Boddie). Initially 25 H. zea were exposed to C. vanessae, none of which produced parasitoids. However, exposing caged individuals to parasitoids revealed that a low percentage of H. zea were suitable hosts for C. vanessae. To capture the parasitism rate for this species, the experiment was repeated using another 50 H. zea. To obtain a parasitized Lepidoptera larva, an individual was picked up from a colony cage using thin soft metal forceps, then, still held with forceps, placed into the colony cage of parasitoids and put in contact with a parasitoid’s antennae. This usually resulted in immediate parasitism (i.e., insertion of the ovipositor for at least one second, and position of its wings and middle and hind legs upward (Fig. 3.1), a display that I previously observed with other braconid species and was assumed to be indicative of oviposition). When a parasitoid didn’t respond, the larva was presented to another parasitoid, until one eventually responded. Sometimes when few parasitoids were available, none of them were willing to parasitize hosts. In this case, a parasitoid was collected using a mouth aspirator, the host was placed in an experimental cup, and the parasitoid was rapidly thrown onto it. This usually triggered immediate parasitism. Parasitoids were quick to parasitize some species of lepidopterans, but required repeated exposure over extended periods before they parasitized other species. Eventually, 37 Chapter 3 however, they parasitized all exposed lepidopteran species with the exception of Spilosoma virginica Fabricius (Lepidoptera: Erebidae: Arctiinae) (~10 individuals exposed in 2013 and 30 in 2014). The heavy pilosity of these larvae probably prevented parasitism (although six individuals of another hairy Arctiinae, Estigmene acrea (Drury), were parasitized on their hairless ventral side). Larvae were slowly removed from the cage while being parasitized. When the parasitoid detached, both insects were quickly placed into an experimental container to allow for further parasitism. This was rarely observed. Parasitoids subsequently were removed after 48 h and returned to the colony, or discarded when dead. The experimental container was a translucent plastic cup (240 ml capacity, Polar Plastic Ltd.) containing a disc of filter paper (55 mm diameter) at the bottom, on top of which a block of diet was placed (7.3 ml). Each diet block had a square of paraffin film (3 x 3 cm) placed on top and below to reduce evaporation. A small drop of honey was placed on top of the upper paraffin film and some water was sprayed into the cup to sustain the parasitoid. Beforehand, numerous small holes (~0.5 mm diameter) were pierced in the bottom of the cups and on the lids to allow airflow and prevent the accumulation of moisture and the development of mold within the cups. Diet blocks were replaced approximately every 5 days as they were consumed or became too dry. At the same time, frass was discarded. For species that did not develop on McMorran diet; i.e., Aglais milberti, Pieris rapae, Habrosyne scripta (Gosse) (Lepidoptera: Drepanidae), and Hyles euphorbiae (Linnaeus) (Lepidoptera: Sphingidae), pieces of plant stems with a few leaves attached were used instead. Excised plant parts were placed onto a piece of paraffin film to avoid contact with the filter paper and sprayed with water daily to reduce wilting. Plant parts were replaced every 2 to 4 days as they were consumed or wilted. Tests were done in a growth cabinet (a different one than that used for colonies) at 12:12 h light:dark cycle, at constant 20 °C, and at 70 to 80% humidity. These conditions were monitored using a digital device measuring temperature and humidity and a data logger measuring temperature, humidity, and light intensity, which were each placed 38 Chapter 3 into two empty experimental containers (240 ml) to ensure that these conditions corresponded to those to which insects were exposed. Parasitized caterpillars remained in these conditions until they either died or completed development to adults. This last outcome took several months for diapausing species (up to 8 months for Mamestra configurata Walker (Lepidoptera: Noctuidae)). Of all the Lepidoptera species tested, only Hyles euphorbiae and Habrosyne scripta did not emerge from their pupae, which suggests that these two species require a cold treatment to break their winter diapause. All other species contained specimens that successfully matured to adults in the conditions described above, given sufficient time, although maturity was not synchronized among all individuals. The fitness of parasitoids on different host species was measured using a suite of parameters, including: - The percent parasitism (number of caterpillars from which parasitoids egressed / [total number of parasitized larvae - larvae that died prior to pupating without producing parasitoids]). - The development time of parasitoid broods within the host (from ovipotition to egression of larvae from the host). - The development time of parasitoid broods within their cocoon masses (from egression of larvae from the host to first adult emergence from the cocoon mass). - The number of adult parasitoids that emerged from each host, here referred to as “brood size” (parasitoids that didn’t fully emerge from cocoon masses were not counted in order to only consider individuals that could contribute to the next generation). - The dry mass of individual adult parasitoids (parasitoids were dried in their experimental container in a rearing room at 20 °C, 20% relative humidity). 39 Chapter 3 To measure parasitoid dry mass, the individual mass of 10 haphazardly selected parasitoids in pristine conditions from each brood was measured. However, to ensure completion of the data collection on time, for about half of the host species it was necessary to instead measure the mass of 30 parasitoids simultaneously from each brood; i.e., Abagrotis reedi Buckett, Actebia balanitis (Grote), Agrotis ipsilon (Hufnagel), Anagrapha falcifera (Kirby), Anaplectoides prasina (Denis & Schiffermüller), Apamea devastator (Brace), Apamea lignicolora (Guenée), Caradrina morpheus (Hufnagel), Chrysodeixis includens (Walker), Dargida diffusa, Feltia herilis (Grote), Helicoverpa zea, Heliothis virescens (Fabricius), Lacanobia grandis (Guenée), Mythimna unipuncta (Haworth), and Spaelotis clandestina (Harris) (Lepidoptera: Noctuidae). 3.3.5. Phylogeny of lepidopteran species Genetic relatedness of the Noctuoidea used in this experiment was compared using CO1 barcodes generated from specimens used in this experiment and others from GenBank (Benson et al., 2005) and Barcode of Life Data Systems (BOLD) (Ratnasingham et al., 2007) for species that were not tested in this experiment, but which are considered reliable hosts for C. vanessae (Hervet et al., 2014). A neighbour-joining tree (Kimura 2-parameter distance) was generated to assess the host quality of different clades of Noctuidae according to the maximum likelihood of evolutionary distance among species based on their CO1 sequences (Kimura et al., 1980), and to predict the suitability of non-tested species as hosts for C. vanessae. 3.3.6. Investigating parasitized caterpillars that did not produce parasitoids Many parasitized caterpillars survived and metamorphosed to adults. To verify whether parasitoid eggs were actually laid in these larvae, H. zea (a host with a low rate of successful parasitism; n = 20) and Trichoplusia ni (a host with a high rate of successful 40 Chapter 3 parasitism; n = 20) larvae were exposed to C. vanessae until the parasitoid displayed the putative oviposition posture on each caterpillar for more than one second. Five larvae from each of these two species were dissected immediately following parasitism to allow for observation of parasitoid eggs. The remaining caterpillars were maintained in individual cups in conditions previously described, and were dissected throughout the following 11 days to observe different stages of the parasitoid and of the hosts’ immune reaction. To prevent gut contents from clouding the haemolymph, the head and the last rear abdominal segments of each caterpillar were excised and the digestive system removed. Haemolymph was then squeezed from the hemocoel onto a microscope slide, diluted with one drop of saline solution, and topped with a coverslip. The haemolymph was checked for eggs using a compound light microscope (400 x). Two H. zea dissected had too much fat, which clouded the haemolymph and rendered potential parasitoid eggs undetectable. To overcome this issue, remaining larvae were kept 2 days in a growth chamber without food to reduce their amount of fat tissues. Parasitized caterpillars occasionally died without producing parasitoids. These caterpillars are referred to as having died of “premature death”. Potential causes include: diseases, unsuitable rearing conditions, an indirect effect of parasitism by C. vanessae (discussed later), and death by other parasitoid species (for field-collected larvae only). Because it was unknown if these dead larvae would have produced C. vanessae under different conditions (some dissected dead caterpillars contained dead C. vanessae larvae), they were excluded from analyses. Premature death was observed in nearly all species tested, but was particularly evident in a few species; e.g., Spodoptera frugiperda (J. E. Smith) and Anarta trifolii (Hufnagel) (Lepidoptera: Noctuidae), all of which died following parasitism by C. vanessae, but without producing parasitoids, while conspecific non-parasitized individuals in the colony cages they came from were healthy. Some of these caterpillars were dissected upon death and their haemolymph observed with a dissecting microscope (15 x), 41 Chapter 3 but no parasitoid larvae were detected. New specimens of A. trifolii were obtained the following year using UV light traps to test experimentally whether C. vanessae induced premature death of these caterpillars. Larvae (n = 70) were reared individually in experimental containers (240 ml) using the setup previously described. Half of these larvae (n = 35) were parasitized by C. vanessae while the other half were not parasitized (control group). However, this time nearly all A. trifolii that were exposed to C. vanessae produced parasitoids (results of this test were included in the host-range and host-fitness experiment previously described). These different outcomes raised concerns regarding the conspecificity of individuals used in both tests. To verify the identity of specimens, CO1 sequences of specimens from both tests were compared (GenBank accession numbers: KX281193 and KX281194). They showed to be 100% identical and to match sequences of A. trifolii voucher specimens in GenBank. Since C. vanessae didn’t induce premature death of A. trifolii in the second test, the test was then repeated using the same protocol but this time with S. frugiperda. Time to death of parasitized S. frugiperda larvae and time to pupation and adult emergence of non-parasitized individuals were recorded. Haemolymph of dead S. frugiperda larvae was observed with a compound light microscope (400 x) immediately following death of individuals. 3.3.7. Data analyses A lepidopteran species was defined as a host if it produced at least one adult parasitoid. A χ2 test was used to compare percentages of parasitism among host species, followed by post hoc comparisons between pairs of species using Fisher’s exact tests with Bonferroni correction to detect which species differed. A χ2 test was also used to compare the percentage of premature death between parasitized and non-parasitized S. frugiperda to test whether C. vanessae induced premature death of this species. A Welsh’s approximate two-sample test (alternative to t-test since standard deviations of samples were unequal) 42 Chapter 3 was used to compare time to death of parasitized S. frugiperda and time to metamorphosis to adults of non-parasitized S. frugiperda. One-way analysis of variance (ANOVA) tests were performed to examine the effect of host species on parasitoid brood size and on parasitoid development time within cocoons. When results detected a significant effect of host species, Tukey’s range tests were performed to identify which host species had significant differences. Assumptions of normal distribution and homoscedasticity of variables were verified using residual-by-predicted plots. Kruskal-Wallis tests were performed to analyze C. vanessae development time in the host, total development time, and estimated total brood mass because ANOVA assumptions of homoscedasticity between host species and normality in a few species were violated. When Kruskal-Wallis tests detected a significant difference, post hoc tests were performed using Dunn tests (pairwise multiple comparisons using rank sums) to detect differences between groups. It was hypothesized that differences in parasitoid fitness parameters between host species might be confounded by other factors. For example, if there is scramble competition within hosts, larger parasitoid broods may result in smaller individuals. In addition, considering that smaller individuals have less body mass to produce, smaller parasitoids may develop faster than larger parasitoids, thus brood size may directly influence parasitoid development time. Thirdly, if smaller parasitoids develop faster within the host, they may also develop faster within their cocoons for the same reason. Conversely, the opposite may also occur: due to the fact that parasitoids from the same brood egress simultaneously from their host, broods that develop rapidly in the host may comprise less-developed individuals. Thus, parasitoids that have a short development time within their host may compensate by having a longer development time within their cocoon. For these reasons, three analyses of covariance (ANCOVA) tests were performed to test for an effect of: host species and parasitoid brood size on average individual parasitoid mass, host species and brood size on C. 43 Chapter 3 vanessae total development time, and host species and C. vanessae development time in the host on the development time of C. vanessae in cocoons. When a significant interaction between the main effects was detected, scatterplots and post hoc Pearson product-moment correlations with Bonferroni correction were used to identify the nature of the effects and which treatments differed. All analyses were conducted in R version 3.3.0 (Dinno, 2016; Fox et al., 2011; R Core Team, 2016), which was also used to generate the figures. For figures displaying results of statistical analyses, species were ordered along the x axis starting with the species providing best fitness for C. vanessae for the variable analyzed, and in such way that the lines showing lack of significant difference would be the least interrupted. Unless otherwise stated, all means are reported ± SE; in some instances, ranges are provided. 3.4. Results 3.4.1. Fitness of C. vanessae in relation to host species The experimental testing identified 31 previously unknown hosts of C. vanessae (see Table 3.1). The morphology of the species “Abagrotis sp.” closely matches that of Abagrotis orbis (Grote), but CO1 sequences from four specimens put it closer to Abagrotis baueri McDunnough (Lepidoptera: Noctuidae) than to A. orbis. It may be either one of those two species, or is an undescribed species. Larvae of this species feed on buds of grapevines, Vitis spp. (Vitaceae), and it appears to be an important pest of vineyards in British Columbia, Canada. Lepidoptera species that were already known hosts of C. vanessae confirmed to be hosts in this experiment were Anarta trifolii, Apamea sordens, and Trichoplusia ni (Hervet et al., 2014; Tobias, 1971, 1976, 1986). Species that did not produce C. vanessae in this study (n = 13) are listed in Tables 3.1 and 3.2. Parasitism rate was significantly different among host species (χ232 = 451.98, P < 0.001) (Fig. 3.2). Result from Amphipyra tragopoginis (Clerck) (Lepidoptera: Noctuidae) is not 44 Chapter 3 displayed in Fig. 3.2 because only a single individual was exposed to C. vanessae, which produced parasitoids. Results indicate that suitable hosts can be found in the families Noctuidae and Nymphalidae, and that hosts of similar parasitism rate are often closely- related; e.g., Plusiinae, Apameini, Euxoa spp., Abagrotis spp., Agrotis spp., Lacinipolia spp., and Spodoptera spp. However, there were exceptions; e.g., Noctuini, Hadenini and Feltia spp. (Fig. 3.3). The wide range of parasitism success detected among Noctuini species is likely an effect of the high number of species tested in this group. Non-Noctuoidea species for which CO1 sequences were obtained – thus are not displayed in Fig. 3.3, are Aglais milberti (GenBank accession number: KX281189) and Habrosyne scripta (GenBank accession number: KX281205). Development time of C. vanessae varied among host species in the host (Kruskal- Wallis, χ224 = 415.94, P < 0.001), in the cocoon (ANOVA, F24, 474 = 7.08, P < 0.001), and total development time (Kruskal-Wallis, χ224 = 383.41, P < 0.001) (Fig. 3.4). Total parasitoid development time for species from which fewer than five individuals produced parasitoids are not displayed in Fig. 3.4 because, due to a lack of power stemming from too few replicates, significant differences could not be detected; i.e., Agrotis ipsilon (26.8 ± 0.19 days, n = 4), Apamea sordens (30.7 ± 0.75 days, n = 2), Dargida diffusa (31.3 ± 1.32 days, n = 3), Euxoa messoria (Harris) (29 ± 2.52 days, n = 3), Helicoverpa zea (27.6 ± 0.61 days, n = 3), Lacanobia grandis (28.2 ± 24.5 days, n = 2), Mamestra configurata (27.82 ± 0.02 days, n = 2), and Spaelotis clandestina (28.5 ± 0.18 days, n = 2). Regarding the potential effect of brood size on parasitoid development time, there was a significant interaction between host species and C. vanessae brood size on C. vanessae total development time (ANCOVA, F24, 445 = 1.67, P = 0.03). There was a significant effect of species (ANCOVA, F24, 445 = 19.5, P < 0.001) (Fig. 3.5), but not of total development time (ANCOVA, F1, 445 = 0.31, P = 0.58) on brood size. Scatterplots and Bonferroni-corrected Pearson product-moment correlation test showed no effect of parasitoid brood size on total 45 Chapter 3 development time (all P ≥ 0.006). Parasitoid brood size for species from which fewer than five individuals produced parasitoids were not displayed in Fig. 3.5 because of low power; i.e., A. ipsilon (71.5 ± 17.97, n = 4), D. diffusa (28 ± 23.07, n = 3), H. zea (24 ± 16.33, n = 3), L. grandis (130 ± 24.50, n = 2), M. configurata (119 ± 16, n = 2), S. clandestina (120 ± 20.50, n = 2), A. tragopoginis (42, n = 1), and A. sordens (1, n = 1). Host species associated with longer parasitoid development time in the host were usually also associated with longer parasitoid cocoon development time; e.g., parasitoids from Feltia herilis showed both the longest development time in the host (32.7 ± 1.4 days) and in the cocoon (11.1 ± 0.1 days). There was a significant interaction between C. vanessae development time in the host and host species on the development time of C. vanessae in- cocoons (ANCOVA, F24, 449 = 4.09, P < 0.001). There was also a significant effect of host species (ANCOVA, F24, 449 = 8.20, P < 0.001), but not of development time in the host (ANCOVA, F1, 449 = 1.61, P = 0.21) on C. vanessae in-cocoon development time. Post hoc Pearson product- moment correlation tests and scatterplots detected a significant negative correlation between C. vanessae development time in the host versus development time in the cocoon only for a few host species; e.g., Agrotis vancouverensis, Anaplectoides prasina, and Noctua pronuba (Linnaeus) (all P < 0.001). There was no interaction between C. vanessae development time in the host and in the cocoon for the other host species. Average individual parasitoid mass per brood was different among host species (one- way ANOVA, F24, 484 = 13.13, P < 0.001) (Fig. 3.6). There was a significant interaction between host species and parasitoid brood size on individual parasitoid mass (ANCOVA, F24, 464 = 4.47, P < 0.001). There were also significant effects of host species (ANCOVA, F24, 464 = 18.17, P < 0.001) and brood size (ANCOVA, F1, 464 = 105.38, P < 0.001) on the mass of individual parasitoids. Scatterplots showed negative correlations between brood sizes and parasitoid individual mass for six species, as did Bonferroni-corrected Pearson product-moment correlation tests; i.e., Trichoplusia ni, Autographa californica (Speyer), Chrysodeixis includens, 46 Chapter 3 Actebia balanitis, Abagrotis reedi, and Euxoa satis (Harvey) (all r ≤ -0.70, df = 15 to 32, all P < 0.001). These species were amongst those that produced both the heaviest and the largest broods (Figs. 3.5 & 3.7). These six hosts also produced among the highest estimated brood masses, surpassed only by Anarta trifolii (22.94 ± 1.21 mg), Noctua pronuba (22.07 ± 1.50 mg), and Anaplectoides prasina (19.91 ± 1.25 mg), which were the three host species that also produced that largest parasitoid broods. Average individual parasitoid mass per brood for species from which fewer than five individuals produced parasitoids is not displayed in Fig. 3.6 due to low power; i.e., A. ipsilon (0.15 ± 0.003 mg, n = 4), D. diffusa (0.14 ± 0.023 mg, n = 3), H. zea (0.17 ± 0.023 mg, n = 3), L. grandis (0.13 ± 0.004, n = 2), M. configurata (0.15 ± 0.010 mg, n = 2), and S. clandestina (0.16 ± 0.024 mg, n = 2), and A. sordens (0.24 mg, n = 1). For similar reasons, estimated parasitoid brood mass for these species is not displayed in Fig. 3.7: A. ipsilon (10.60 ± 2.44 mg, n = 4), D. diffusa (3.90 ± 3.09 mg, n = 3), H. zea (4.35 ± 3.11 mg, n = 3), L. grandis (17.15 ± 3.74 mg, n = 2), M. configurata (18.42 ± 3.59 mg, n = 2), and S. clandestina (18.66 ± 0.48 mg, n = 2), and A. sordens (0.24 mg, n = 1). Even if host size was not measured, it was evident that the largest host species didn’t always produce the largest broods or the heaviest parasitoids; e.g., Eurois occulta (Linnaeus) larvae were noticeably larger than Anarta trifolii larvae, although A. trifolii produced significantly larger broods than E. occulta. In addition, E. occulta and N. pronuba were the largest species used in this experiment, but many other species of noticeably smaller size produced parasitoids of comparable mass (Fig. 3.6). In addition, brood size varied considerably both within and among host species (Fig. 3.5). For example, A. trifolii larvae produced parasitoid broods ranging from one to 250 individuals (250 being the largest brood observed in this experiment). Other hosts that produced broods of one were: A. sordens (one parasitoid larva egressed and formed a cocoon, which produced an adult), Feltia jaculifera (Guenée), and H. virescens (in both cases a few parasitoid larvae egressed and formed a cocoon mass from which only one adult emerged). Three H. virescens individuals produced 47 Chapter 3 one, one, and five parasitoid larvae, which failed to produce cocoons and eventually dried out and died. These H. virescens were considered to have died producing parasitoids, although they didn’t have an associated brood size. There was little variation in the mass of individual parasitoids from the same brood. For example, average individual parasitoid mass from T. ni hosts was 0.18 ± 0.0025 mg (n = 251 individuals from 26 broods). Their dry mass varied between 0.08 mg to 0.33 mg (individuals from two different broods). However, out of 26 broods, the average difference between the heaviest and the lightest of 10 individuals chosen randomly from the same brood was 0.06 ± 0.0048 mg. The smallest difference of dry mass between two individuals emerged from a brood was 0.02 mg (with 0.11 mg and 0.13 mg, n = 10), and the greatest difference was 0.12 mg (with 0.14 mg and 0.26 mg, n = 10). 3.4.2. Oviposition testing All larvae into which C. vanessae inserted its ovipositor (20 T. ni and 17 H. zea) contained either parasitoid eggs (observed up to 4 days following parasitism, at which time embryos were visible within) or larvae (observed starting 5 days after parasitism). Parasitoid eggs and larvae were observed within the haemolymph of each dissected T. ni (Fig. 3.8 A & B). None of them were encapsulated by the host’s hemocytes, which would have been a normal immune response for lepidopteran larvae to parasitism (Lavine et al., 2002). Cotesia vanessae larvae were observed to produce teratocytes (Fig. 3.8 B), which are macrocells known to have trophic, immunosuppressive, and secretory functions (Dahlman, 1990, 1991). Observations (> 5 days) showed that a few eggs failed to hatch (three in total), and even those were not encapsulated. While when oviposited into H. zea, parasitoid eggs were observed to become encapsulated four days after parasitism, and all H. zea dissected 11 days after parasitism contained numerous dead first-instar parasitoid larvae encapsulated by hemocytes (Fig. 3.8 C & D) and no live larvae, except for one individual. This individual 48 Chapter 3 contained both encapsulated and live larvae. All encapsulated larvae appeared rather small and shrivelled (Fig. 3.8 D), which suggests that they had died shortly after hatching, while live larvae were much bigger. These observations show that parasitoids laid eggs even within caterpillars that did not support their development. In all lepidopteran larvae that contained parasitoid larvae, always a great number of them (about half of all observed parasitoid larvae in a host) had been cut in half in a transverse plane, usually about half way down their body (Fig. 3.8 E). This was indicative that sororicide occured. Larval sororicide was later further evidenced by direct observation of the act (Fig. 3.8 F). 3.4.3. Testing if C. vanessae causes premature death of Lepidoptera larvae Results obtained for Spodoptera frugiperda showed that parasitism can cause the premature death of the host. All S. frugiperda parasitized died of premature death while all non-parasitized S. frugiperda (used as control) survived. These two outcomes were significantly different (χ21 = 70, n1 = 35, n2 = 35, P < 0.001). Larvae from the parasitized group matured to their last instar, but never pupated. Last-instar larvae slowly bloated until their integuments couldn’t inflate anymore. At this point they stopped feeding, dark areas eventually appeared on their cuticle and spread over time until larvae were entirely black. During this darkening process the larvae started to shrink, likely because of water loss. Eventually larvae were entirely dark and motionless, at which point they appeared dead, but were still responsive to touch. They were touched daily with forceps and assumed dead when unresponsive. The time taken for parasitized individuals to die was significantly longer than that required for non-parasitized individuals to metamorphose to adults (Welsh’s approximate two-sample t-test, t = -7.45, df = 40.35, P < 0.001). Dissection of dead larvae showed that their hemocoel was nearly depleted of haemolymph and that their gut was highly bloated and occupied nearly the entire hemocoel. Dissection of the gut showed that it was filled with a gelatinous orange substance similar to the artificial diet, which it was 49 Chapter 3 assumed to be (Fig. 3.9). Observation of their haemolymph revealed dead encapsulated first instar parasitoids, which, considering their small size, had died shortly after hatching and long before the death of the host. In these parasitized S. frugiperda larvae, as in H. zea, and a number of Mythimna unipuncta (> 10, dissected for unrelated reasons), encapsulated parasitoids were observed but none of them was also melanised. Although the relationship between premature death and parasitism was documented only in S. frugiperda, similar observations were made in a number of species. For example, time to premature death for A. ipsilon larvae (n = 20), once parasitized, averaged 98.5 ± 3.07 days (range: 74 to 120), about twice the duration observed for S. frugiperda; while individuals that metamorphosed to adults (n = 4) did so in 46.5 ± 4.5 days of parasitism. For A. trifolii (n = 40), larvae died only after 20.9 ± 2.2 days (range: 3 to 30) of parasitism. 3.5. Discussion The overall results show that C. vanessae has a broad fundamental host range within Noctuidae and may potentially become an important parasitoid of key pest species in North America. Hosts in the subfamily Plusiinae were among those that displayed the highest percent of successful parasitism, produced the largest broods, the heaviest individuals, and the fastest-developing parasitoids. In Canada, C. vanessae has only been recovered in the wild from Plusiinae; i.e., Chrysodeixis chalcites, Trichoplusia ni, and Autographa californica (Hervet et al., 2014), confirming that Plusiinae are natural hosts. However, it has been demonstrated with other parasitoid species, including with Cotesia sesamiae (Cameron) (Branca et al., 2011), that conspecific parasitoids from different geographic regions do not display a similar specificity on the same host species (Dubuffet et al., 2006). Because C. vanessae from a single source population was used in the current experiment, it may be that C. vanessae from other populations have different host specificities. 50 Chapter 3 Nymphalini appear to be the primary hosts of Cotesia vanessae in the southern- western part of its Palearctic range where temperatures are warm enough for parasitoids to refrain from diapause (Stefanescu et al., 2012), but in the northern part of its range C. vanessae must necessarily overwinter within an overwintering host larva. Nymphalini commonly either overwinter as adults or migrate to southern latitudes for the winter (Dvořák et al., 2002; Scott, 1979); thus, parasitoids of northern latitudes must find other hosts for the winter period. Noctuidae parasitized by C. vanessae have been collected in October (Italy) and May (England), which suggests that C. vanessae overwinters within overwintering noctuid larvae in the northern part of its range (Nixon, 1974). This would explain the affinity of C. vanessae to Noctuidae, which may be stronger in overwintering northern populations. In the current study, parasitoids were observed to develop slower on overwintering hosts, thus supporting the previous claim that they overwinter within overwintering hosts. Individuals of univoltine lepidopteran species that overwinter as larvae necessarily are at the larval stage during the fall, and do not occur in the larval stage any other time of the year. Such species were exposed to C. vanessae in the fall while they were diapausing or soon to do so, although they remained at 20 °C in experimental conditions throughout their entire development. In Fig. 3.4, the species between (and including) Apamea devastator and Cryptocala acadiensis (Bethune) (Lepidoptera: Noctuidae) both overwinter as larvae and are univoltine throughout their range, except for Euxoa tristicula (Morrison), a species that can be multivoltine. However, personal observations and previous studies (Jacobson, 1969) show that E. tristicula must necessarily go through a diapause at the larval stage, which lasts at least two months at 20 °C. Parasitoids took the longest time to develop within these diapausing hosts. This indicates that development within overwintering hosts likely triggered parasitoids to initiate overwintering, but after a short diapause (from a few days, up to two months in Cryptocala acadiensis (Fig. 3.4)) parasitoids eventually resumed their development, likely because of conditions non-optimal for overwintering (including too high 51 Chapter 3 temperature, too long light cycle, and too strong light intensity). Regarding the other species in Fig. 3.4, only Abagrotis spp., Agrotis vancouverensis, and Anaplectoides prasina also overwinter as larvae and are likely to be obligately univoltine throughout their range. The reason why parasitoids did not attempt to diapause within these hosts is unknown. Putting aside overwintering hosts, parasitoids usually developed faster in host species that also produced larger broods and heavier individual parasitoids. This observation supports previous findings that fast development can be associated with higher fitness (Doyon et al., 2005; Gao et al., 2016), and that parasitoids may display different fitness when using different host species (Greenblatt et al., 1981). Host species in which parasitoids required longer development period were also usually those that were associated with longer parasitoid cocoon development time. There were two exceptions to the trend that univoltine species that overwinter as larvae had longer development time in both the host and the cocoon: Apamea lignicolora and Cryptocala acadiensis. Both species produced parasitoids that had amongst the longest development time in the host and amongst the shortest development time in the cocoon. On the other hand, brood size and parasitoid mass did not appear to always be associated with host size. This effect has also been observed in other species, and may be partly explained by the suitability of host species to the parasitoid species, and by how adequate the diet is for these hosts (Greenblatt et al., 1981). Comparison of genetic relatedness allows for tentative predictions regarding what additional pest species in North American might be hosts for C. vanessae. Results indicate that many species in the subfamily Plusiinae, in the tribes Noctuini, and Hadenini, and in the genera Apamea and Mythimna are likely to be hosts. Hosts were found in the four subfamilies of Noctuidae tested. This indicates that other subfamilies of Noctuidae likely also contain hosts. Most of the main pest species of Noctuidae in North America were tested in this experiment, the majority of which supported the development of C. vanessae. Other important pests that were not tested that are likely hosts include: the cotton bollworm, 52 Chapter 3 Helicoverpa armigera, the pale western cutworm, Agrotis orthogonia, and the granulate cutworm, Feltia subterranea (Fabricius). Any species in the tribe Nymphalini are also likely to be hosts. On the other hand, species in the family Erebidae and in the genera Lacinipolia and Spodoptera are not likely to be hosts. This experiment did not test 30 species previously reported to be hosts of C. vanessae: Aglais io (L.), Aglais urticae L., Aglais urticae turcica (Staudinger), Nymphalis polychloros (L.), Vanessa atalanta (L.), Vanessa cardui (L.) (Nymphalidae: Nymphalinae), Limenitis camilla (L.) (Nymphalidae: Limenitidinae), Argynnis aglaja (L.) (Nymphalidae: Argynninae), Autographa gamma (L.), Chrysodeixis chalcites (Esper), Cornutiplusia circumflexa (L.), Macdunnoughia confusa (Stephens) (Noctuidae: Plusiinae), Actebia praecox (L.), Actebia fugax (Noctuidae: Noctuinae), Acontia lucida (Hufnagel) (Noctuidae: Acontiinae), Lacanobia oleracea (L.), Mamestra brassicae (L.), Mythimna litoralis (Curtis) (Noctuidae: Hadeninae), Helicoverpa armigera (Hübner), Heliothis viriplaca (Hufnagel) (Noctuidae: Heliothinae), Nycteola revayana (Scopoli) (Noctuidae: Sarrothripinae), Calophasia opalina (Esper) (Noctuidae: Cuculliinae), Simyra dentinosa Freyer (Noctuidae: Acronictinae), Cerura vinula (L.), Notodonta ziczac (Notodontidae), Cnaemidophorus rhododactylus (Denis & Schiffermüller) (Pterophoridae: Pterophorinae), Lasiocampa trifolii Denis & Schiffermüller, Malacosoma neustria (L.) (Lasiocampidae: Lasiocampinae), Loxostege sticticalis (L.) (Pyralidae: Pyraustinae), Thaumetopoea processionea (L.) (Thaumetopoeidae) (Aubert, 1966; Aue, 1938; Doğanlar, 1982; Fahringer, 1936; Hervet et al., 2014; Karimpour et al., 2001; Meyer, 1927; Morley, 1906; Obregón et al., 2015; Özbek et al., 2012; Papp, 1988, 2011-2012; Schwarz et al., 2000; Shaw et al., 2009; Telenga, 1955; Tobias, 1954, 1971, 1976, 1986; Yu et al., 2016; Zhumanov, 1980). Spodoptera exigua (Hübner) has been reported as a host of C. vanessae (Tobias, 1971), although it didn’t support the development of C. vanessae in the current study. 53 Chapter 3 Results indicate that testing a lepidopteran species multiple times may yield different outcomes (as illustrated by the tests using Anarta trifolii and Helicoverpa zea). This shows that results from only one set of parasitized caterpillars should be viewed with caution, especially for species listed as non-hosts. Thus, failing to confirm the host status of S. exigua does not invalidate this previous report. However, previous reports of parasitoid-host association do not often provide strong evidence of accurate identification of species, either for field-collected host larvae or emerged parasitoids (Nixon, 1974). Thus, some of these reports may be considered doubtful (Hervet et al., 2014). In light of the results and other reliable sources (Shaw et al., 2009; Stefanescu et al., 2012), the records considered to be most reliable, or at least to be the most common hosts, identify species of Noctuidae and Nymphalini. But, given its broad host range in these taxa, host species in other Lepidoptera families seem probable. Many Cotesia species are known to be generalists. During the current study C. vanessae was observed to insert its ovipositor and assumed the putative oviposition posture on all exposed lepidopteran species (except Spilosoma virginica (Fabricius) (Lepidoptera: Erebidae)), which belonged to 11 families; i.e., Drepanidae, Erebidae, Hesperidae, Lasiocampidae, Noctuidae, Nymphalidae, Papilionidae, Pieridae, Plutellidae, Sphingidae, and Tortricidae. Many of these species were not previously mentioned because they didn’t produce parasitoids and the number of exposed larvae was low (< 6 per species). Hence, no conclusion could be drawn regarding their possible non-host status, but this illustrates how opportunistic C. vanessae is. A review of the Cotesia species in Taxapad (Yu et al., 2012) (n = 266) showed that for those that have at least one known host species (n = 214), over one third of them (n = 88) have been reported from Noctuidae. This illustrates that there is an affinity of Cotesia species for members of this family. For the Cotesia species that had at least two known host species (n = 135), over half of them (n = 75) were associated with hosts in different families, which often included micro- and macrolepidopterans, as well as moths and 54 Chapter 3 butterflies. Some Cotesia species of particular economic importance have been extensively studied, and for some of them this has yielded the discovery of a high number of host species; e.g., Cotesia ruficrus (Haliday): 83 host species in 15 families, Cotesia marginiventris (Cresson): 42 host species in 6 families, Cotesia flavipes Cameron: 37 host species in 6 families, and Cotesia congregata (Say): 28 host species in 5 families (Yu et al., 2016). This illustrates the number and diversity of hosts associated with these parasitoids, although these numbers should be taken with caution as many reports likely stem from misidentifications (Nixon, 1974). Although C. vanessae accepted nearly all the Lepidoptera species in the current experiment, it was sometimes necessary to wait for parasitoids to get closer to the end of their life for them to accept certain host species. Plusiinae were always readily accepted regardless of parasitoid age, and to a lesser extent other Noctuidae too, but Lepidoptera in other families were generally not accepted by young adult parasitoids. This effect has been described as an aspect of “dynamic state variable theory”, known as “partial preference”, which states that the acceptance of less desirable species depends on to the parasitoid egg load, fat reserves, and age (Clark et al., 2000; Mangel, 1989). This means that the host range of a parasitoid species is likely to change over time, with older parasitoids accepting an increasing number of species as hosts. The plant species that the hosts feed on are a crucial aspect of parasitoid-host specificity. Some Cotesia species, although they have been reared from taxonomically distant hosts, appear to be associated with few plant species. For example, Cotesia ferruginea (Marshall) has been reared only from three host species, each of them belonging to a different family: Archanara geminipuncta (Haworth) (Lepidoptera: Noctuidae), Phragmataecia castanea (Hübner) (Lepidoptera: Cossidae), and Chilo phragmitella (Hübner) (Lepidoptera: Pyralidae) (Wilkinson, 1945). The two first species feed solely on Phragmites spp. and the third one feeds both on Phragmites and Glyceria maxima (Hartm.) Holmb. 55 Chapter 3 (Poaceae). This suggests that C. ferruginea is rather associated with specific plant species than specific host species. The hypothesis that some parasitoid species are associated with plant species rather than host species, and to this effect parasitize taxonomically distantly related species simply because they feed on the same plant species, has previously been formulated (Cushman, 1926). To test this hypothesis, Tawfik (1957) exposed Cotesia glomerata, a known parasitoid of Pieris spp. (Lepidoptera: Pieridae) associated with Brassica spp. (Brassicaceae), to a non-host species, Mamestra brassicae, which also feeds on Brassica spp. In the laboratory, C. glomerata oviposited within M. brassicae. Although the eggs didn’t hatch within this species because they became encapsulated, this showed a key biology aspect of the parasitoid for selecting hosts. Similarly, the solitary parasitoid Cotesia marginiventris and the gregarious parasitoid Cotesia congregata, which were thought to be specific on Noctuidae and Sphingidae respectively, were exposed to the other species’ natural host in the laboratory; i.e., C. marginiventris to Manduca sexta (L.) (Lepidoptera: Sphingidae) and C. congregata to T. ni (Noctuidae). Wasps of both species were found to readily parasitize these larvae, and C. marginiventris even did so in the presence of its natural host S. exigua. Surprisingly, individuals of both lepidopteran species produced parasitoids, some of which successfully matured to adults (Beckage et al., 2003; Beckage et al., 2002). These results show that Cotesia parasitoids are not as specific as they seem to be to particular host species, and that distantly related lepidopteran species may be suitable hosts. However, species found to be hosts in the laboratory may never be hosts in natural settings. A number of steps are necessary to lead to successful parasitism (i.e., host habitat location, host location, host acceptance, and host suitability), which are each essential to complete parasitism (Vinson, 1980). The current study focuses on host suitability. Therefore, resuts reflect the fundamental, but not ecological, host range of C. vanessae. To understand the ecological host range of C. vanessae it is essential to at least know the plant species it is associated with. Plant species from which lepidopteran larvae 56 Chapter 3 parasitized by C. vanessae have been collected include: thistles (tribe Cynareae, family Asteraceae), common mallow, Malva sylvestris (L.) (Malvaceae) (Stefanescu et al., 2012), tomatoes, Solanum lycopersicum Linnaeus (Solanaceae), cucumbers, Cucumis sativus Linnaeus (Cucurbitaceae), cabbages, Brassica spp. (Hervet et al., 2014), and Chenopodiastrum murale (L.) S. Fuentes, Uotila & Borsch (Amaranthaceae) (Papp, 2011- 2012). Known host plants of previously reported host species may also indicate an affinity of C. vanessae for: nettles, Urtica spp., beet, Beta vulgaris Linnaeus (Amaranthaceae), poplars, Populus spp., and willows, Salix spp. (Salicaceae) (Yu et al., 2016). The affinity of C. vanessae for both Nymphalinae and Noctuidae may have originated in the common food source of its hosts. Species of Noctuidae have been reported from plant species also used by larvae of V. atalanta, V. cardui, and A. urticae (DBIF, 2016; Shaw et al., 1990; Stefanescu et al., 2012). Serendipitous encounter of parasitoids with prospect hosts on plants may have led to the extension of the host range of C. vanessae from Nymphalidae to Noctuidae, or from Noctuidae to Nymphalidae. Results also identified the occurrence of both scramble and contest competition among parasitoid larvae of the same brood. In the current study, certain host species produced particularly large broods considering the sizes of both parasitoid and host (Le Masurier, 1987), some of which displayed a trade-off between parasitoid brood size and individual mass, which is typical of scramble competition (Den Berg et al., 2006; Gu et al., 2003; Harvey et al., 2013). The little mass variation between individuals of the same brood suggests that available resources are divided somewhat equally among individuals of that brood. Contest competition among sibling parasitoids of the same brood is known to occur in other gregarious species (Giron et al., 2004). However, some of these observations were made when immature parasitoids were artificially injected into hosts containing sibling broods, so it is unclear whether this behaviour also occurs in ‘natural broods’ (i.e., without 57 Chapter 3 artificial injection of immature parasitoids into hosts) (Grbić et al., 1992; Giron et al. 2007). To our knowledge, larval siblicide has not previously been reported in gregarious Braconidae. Contest competition was evidenced by direct observation of larval siblicide. Solitary parasitoids, including within Braconidae, are well known to exhibit siblicidal behaviour, and it is considered a necessary behaviour because the relatively small size of their hosts usually only allows for the development of a single parasitoid (Balduf, 1926; Mayhew et al., 1999; Pemberton et al., 1918; Pexton et al., 2002; Salt, 1961; Willard, 1920; Yu et al., 2008). These species are known to either puncture the integument of their rivals and leave them draining of their haemolymph if they have piercing mandibles, or to messily chew any part of their rivals’ bodies if they have chewing mandibles (Salt, 1941; Yu et al., 2008). However, larvae of C. vanessae, which have chewing mandibles, were observed to always be cut in a straight- across section, suggesting an acquired efficiency. Sibling rivalry seems to take place only to reduce the number of competitors within the host, as cannibalism didn’t appear to occur. Gregarious Cotesia are known to display interspecific competition through physical and physiological competition, and intraspecific competition via scramble competition only (Fidgen et al., 2000; Reitz et al., 1995; Takagi, 1986; Waage et al., 1984). Theoretical work suggests that gregariousness has evolved multiple times from solitary ancestors within the genus Cotesia and their relatives, and that for this evolution to be possible, individuals must lose either their antagonistic behaviour toward their siblings or their motility (Mayhew, 1998; Mayhew et al., 1999; Michel-Salzat et al., 2004; Pexton et al., 2009; Pexton et al., 2002, 2004, 2005; Pexton et al., 2003). Observations here showed that C. vanessae is siblicidal and likely also has some motility in order to reach and kill siblings in the host’s hemocoel, yet it is gregarious. It has been hypothesized that spines may aid the locomotion of larvae between host tissues (Pinheiro et al., 2010). This hypothesis is supported by observations that show that first and last instar Cotesia flavipes and C. marginiventris have both functional mandibles and 58 Chapter 3 cuticular spines, but their second instar have neither (Boling et al., 1970; Pinheiro et al., 2010; Yu et al., 2008). The same observations were made with C. vanessae (for the last instar, a single row of small spines was observed on the middle of each thoracic and abdominal segment dorsally, and what looked like pairs of pseudopods ventrally). Both first and last instars need locomotion, to reach rivals and to egress from the host, respectively. In addition, observations showed that first instar C. vanessae did not appear to have a caudal appendage to aid their mobility. Thus, their mobility within the host is likely limited. The transition from first to second instar being accompanied with a loss of both physical antagonism and cuticular spines could suggest that spines also provide first instars with some protection against physical attacks from siblings. As previously noted, another defense against other parasitoids is to quickly kill surrounding contestants before being killed by them (Salt, 1941). Larvae of C. vanessae displayed protective mechanisms to survive attacks from the host’s immune system and possibly also from their siblings. To acquire protection against encapsulation they produce teratocytes. These are macrocells released into the host’s hemocoel that serve various functions, including production of Bracovirus that target and destroy the host’s hemocytes (Dahlman, 1990, 1991). Observation from dissected H. zea showed that in some hosts, parasitoids from the same brood survived encapsulation while others did not. It is unknown what caused this in the current study. It has previously been observed that given certain host species and rearing temperatures, a few parasitoids from a brood could successfully hatch from encapsulated eggs (Blumberg, 1977; Blumberg et al., 1991-1992; Blumberg et al., 2001). Other studies found that the rate of encapsulated individuals decreased as clutch size (i.e., number of eggs a parasitoid laid within a host) increased (Kapranas et al., 2012; Kitano et al., 1978). This suggests that clutch size and the timing of egg hatching for some individuals may have prevented them from succumbing to encapsulation. 59 Chapter 3 Cotesia vanessae are very fertile. As previously stated, adult female C. vanessae can parasitize many hosts in a short amount of time, and appear to remain very fertile throughout their adult life. Considering the high number of larvae killed through sororicide and yet the large size of broods produced by hosts raised the question of whether this species was polyembryonic or if it carried a large egg load. The absence of males in the laboratory colony suggested that polyembryony was a reasonable hypothesis, but it was invalidated by observation of single, non-dividing, eggs within hosts, from the time they were laid until they hatched. The hypothesis of large egg load was validated by the observation of large number of eggs within the ovaries of a female. Although, the large number of eggs combined with their small size and their tendency to stick to each other and to the pin used for dissection didn’t allow for a count. Unlike some other parasitoid species (Charnov et al., 1984; Hasan et al., 2010; Zaviezo et al., 2000), female C. vanessae did not appear to adjust clutch size according to host size, and that a supernumerary number (i.e., an excess number) of eggs is almost always laid within hosts. Supernumerary larvae are then destroyed by their siblings through contest competition, thus self-adjusting the size of their brood. This adjustment appears imperfect as dissected hosts that contained clutches small enough that all individuals would likely have had sufficient resources to mature also contained a large number of larvae cut in half. Nonetheless, it is hypothesized that this self-regulation of brood size by the larvae has the advantage of maximizing the use of all available resources within a host, and to better compete with other parasitoid species. This gives the ability of C. vanessae to develop within hosts of a wide range of sizes. The egression of few, sometimes even single, parasitoids from small hosts was observed (these parasitoids developed slowly, and hosts were parasitized at an early stage). This suggests the ability of C. vanessae to undergo facultative gregarious development. In solitary species, the laying of supernumerary eggs within hosts has been attributed to be an adaptation for efficiently competing in multiparasitized hosts (i.e., hosts 60 Chapter 3 parasitized by more than one parasitoid species). In this case, all parasitoid larvae compete until only one survives. For these, a larger number of offspring within a host means a greater chance that one of them survives (Pexton et al., 2005). In the western Mediterranean area, Cotesia vanessae must compete with a number of other parasitoid species, particularly throughout the winter as hosts become scarce (Stefanescu et al., 2012). In such an environment, the presence of supernumerary siblings within a host would likely provide a competitive advantage. A negative consequence of this evolutionary strategy is that females must dedicate resources to producing supernumerary eggs, whose fate is to die. Other Cotesia species that have been reported to display both gregarious and solitary development and to develop in both micro- and macrolepidoptera, such as C. jucunda (Marshall) (Yu et al., 2016), may display similar behavioural adaptations. Another biological aspect of some populations of C. vanessae is thelytoky (Hervet et al., 2014; Stefanescu et al., 2012). Symbiotic bacteria in the genus Wolbachia are known to induce thelytoky in some insects (Czarnetzki et al., 2004; Dong et al., 2006; Kumm et al., 2008; Louis et al., 1993). These bacteria have previously been detected within Cotesia species (Rattan et al., 2011). Using primers for the Wolbachia-specific wsp gene and methods reported in previous studies (e.g., Floate et al., 2006, Li et al. 2015), the presence of Wolbachia within our laboratory C. vanessae was tested, but not detected. Thelytoky can be induced by other effects, including other symbiotic bacteria (Zchori-Fein et al., 2001). Elucidating the origin of thelytoky in this species was not the goal of this study, so was not investigated further. Thelytokous populations of C. vanessae are known to occur in the Mediterranean region, but male-female populations are known from other regions of Europe (Stefanescu et al., 2012). This indicates that the North American population may have originated from the southwestern Palearctic or Afrotropic regions, perhaps accidentally introduced with Chrysodeixis chalcites, a species also endemic to these regions whose 61 Chapter 3 appearance in southern Ontario coincides with that of C. vanessae (Hervet et al., 2014; Murillo et al., 2013). An interesting observation was the premature death of Lepidoptera larvae caused by C. vanessae, to such extent that all parasitized S. frugiperda died without producing parasitoids. Such extreme cases of mutually antagonistic interactions are rarely reported. There were indications that within S. frugiperda, parasitoid larvae were killed by the caterpillar’s immune system shortly after hatching. These caterpillars continued to develop apparently normally until they reached their final instar, but failed to pupate. Their digestive system eventually stopped functioning, which was evidenced in some species by the anus of individuals not fully closing and partly expelled droppings remained attached and dried there (particularly observed in T. ni), and food accumulation within their gut (particularly observed in S. frugiperda). These larvae remained in their final instar much longer than non-parasitized individuals (Fig. 3.9). If parasitism prevented the pupation of some or all individuals in some species, it didn’t in other species. Some species even seemed to be only partly affected. This was particularly evident with Agrotis ipsilon, for which most individuals had a prolonged larval development time and eventually died, or turned into larval-pupal intermediates that died following pupation. Other A. ipsilon died as larvae producing parasitoids, or produced normal pupae that matured to adults. These individuals didn’t seem to display a delay of development time. This shows that parasitism can yield a wide range of possible outcomes, even amongst individuals of the same species. It also shows that this parasitoid species appears more adapted for development using certain host species compared to others. Previous authors found that wasp calyx fluid, alone or combined with venom, polydnaviruses, and teratocytes alone, can induce some or all of these effects. For example, it has been showed that Chrysodeixis includens larvae injected with 0.01 to 0.10 wasp equivalent calyx fluid from Microplitis demilotor Wilkinson (Hymenoptera: Braconidae) developed into non- viable larval-pupal intermediates; while Heliothis virescens was relatively unaffected by 62 Chapter 3 injections of any amount of M. demilotor calyx fluid (Strand et al., 1991). It has also been shown that a combination of venom and calyx fluid injected by female Cardiochiles nigriceps (Viereck) (Hymenoptera: Braconidae) during oviposition suppress the ability of the host’s prothoracic glands to produce ecdysteroid, thus inhibiting the ability of the host to pupate (Tanaka et al., 1991). It has also been shown that a polydnavirus (Bracovirus) released by parasitoids can inhibit the feeding and gut contractions of their hosts, possibly to prevent the host from predating on emerging parasitoid larvae (Beckage, 2012; Cooper et al., 2010; Pruijssers et al., 2009), and arrest development of the host at the prepupal stage (Lanzrein et al., 2012; Soller et al., 1996). Finally, it has also been shown that the presence of braconid teratocytes in H. virescens’ hemocoel could prolong the larval stage of this noctuid, followed by either its pupation or its death, and the pupation resulted in either a normal pupa or a non-viable larval-pupal intermediate (Vinson, 1970; Zhang et al., 1989). The type of response was associated with the number of teratocytes injected, the age of these teratocytes, and the maturity of the host (Zhang et al., 1989). Results show that the North American population of Cotesia vanessae displays adaptations that should enhance survival in different environments, including those with low host availability and high interspecific competition; i.e., 1) Adult females are opportunistic in their selection of hosts and are able to develop within numerous host species. They carry large egg loads, they lay supernumerary eggs within hosts, and their larvae display an antagonistic behaviour, which allows them to: 2) maximize the available resources to produce large broods whenever possible, 3) develop within hosts of a wide range of sizes, 4) display high intra- and inter-specific competitiveness. In addition, their ability to facultative diapause allows them to survive in environments with hot and cold winters. These ecological adaptations, and the known geographic range of this species (Hervet et al., 2014), are indications that C. vanessae should be able to colonize and thrive in many parts of North America, and that it may become an important parasitoid species for the natural biological 63 Chapter 3 control of noctuid pests of certain crops. However, field studies would be needed to elucidate this. The results show that Cotesia vanessae has a broad fundamental host range on Noctuidae species, but to uncover its ecological host range it would be necessary to investigate the plant species that are most attractive to this parasitoid, and more generally the host finding and selection processes, and how the diet of the host influences parasitoid development and fitness (Haye et al., 2005). 64 Table 3.1. Identity of the species studied (“-“ = “none”). Scientific name Common name Status Suitability for Winter diapause Collection Collection Collection location C. vanessae (North America) stage method Hymenoptera Cotesia vanessae (Reinhard) - Beneficial In host In host Hands Essex and Chatham- Kent counties (ON) Lepidoptera Abagrotis sp. - Pest Suitable Larva Adult Hands Agassiz (BC) Abagrotis reedi Buckett - Pest Suitable Larva Adult Hands Agassiz (BC) Actebia balanitis (Grote) - Not a pest Suitable Larva Adult UV trap Lethbridge (AB) Aglais milberti (Godart) Milbert’s tortoiseshell Not a pest Suitable Adult Larva Hands Lethbridge (AB) Agrotis ipsilon (Hufnagel) black cutworm Pest Suitable Pupa Egg Purchased Benzon Research (PA) Agrotis vancouverensis Grote Vancouver dart Not a pest Suitable Larva Adult UV trap Lethbridge (AB) Amphipyra tragopoginis (Clerck) mouse moth Not a pest Suitable unknown Larva Hands Lethbridge (AB) Anagrapha falcifera (Kirby) celery looper Pest Suitable Larva Adult Sweep Lethbridge (AB) net Anaplectoides prasina green arches moth Not a pest Suitable Larva Adult UV trap Lethbridge (AB) (Denis & Schiffermüller) Anarta trifolii (Hufnagel) clover cutworm Pest Suitable Pupa Adult UV trap Lethbridge (AB) Anticarsia gemmatalis Hübner velvetbean caterpillar Pest Not suitable - Egg Purchased Benzon Research (PA) Apamea devastator (Brace) glassy cutworm Pest Suitable Larva Adult UV trap Lethbridge (AB) Apamea lignicolora (Guenée) wood-coloured quaker Not a pest Suitable Larva Adult UV trap Lethbridge (AB) Apamea sordens (Hufnagel) rustic shoulder-knot Pest Suitable Larva Larva Hands Lethbridge (AB) Autographa californica (Speyer) alfalfa looper Pest Suitable Pupa Larva Sweep Lethbridge (AB) net Caradrina Morpheus (Hufnagel) mottled rustic Not a pest Suitable Larva Adult UV trap Lethbridge (AB) Chrysodeixis includens (Walker) soybean looper Pest Suitable Pupa Egg Purchased Benzon Research (PA) Cryptocala acadiensis (Bethune) catocaline dart Not a pest Suitable Larva Adult UV trap Lethbridge (AB) Dargida diffusa (Walker) wheat head armyworm Pest Suitable Pupa Larva Hands Brooks (AB) Chapter 3 65 Eurois occulta (Linnaeus) great brocade Pest Suitable Larva Adult UV trap Lethbridge (AB) Euxoa auxiliaris (Grote) army cutworm Pest Suitable Larva Adult UV trap Lethbridge (AB) Euxoa messoria (Harris) darksided cutworm Pest Suitable Egg Adult UV trap Lethbridge (AB) Euxoa ochrogaster (Guenée) redbacked cutworm Pest Suitable Egg Adult UV trap Lethbridge (AB) Euxoa satis (Harvey) - Not a pest Suitable Egg Adult UV trap Lethbridge (AB) Euxoa tristicula (Morrison) early cutworm Pest Suitable Egg Adult UV trap Lethbridge (AB) Feltia herilis (Grote) master’s dart Pest Suitable Larva Adult UV trap Lethbridge (AB) Feltia jaculifera (Guenée) dingy cutworm Pest Suitable Larva Adult UV trap Lethbridge (AB) Habrosyne scripta (Gosse) scribe moth Not a pest Not suitable Pupa Adult UV trap Lethbridge (AB) Helicoverpa zea (Boddie) corn earworm Pest Suitable - Egg Purchased Benzon Research (PA) Heliothis virescens (Fabricius) tobacco budworm Pest Suitable Pupa Egg Purchased Benzon Research (PA) Hyles euphorbiae (Linnaeus) spurge hawk-moth Not a pest Not suitable Pupa Larva Hands Lethbridge (AB) Lacanobia grandis (Guenée) grand arches moth Not a pest Suitable Pupa Adult UV trap Lethbridge (AB) Lacinipolia renigera (Stephens) bristly cutworm Pest Not suitable Egg Adult UV trap Lethbridge (AB) Lacinipolia sareta (Smith) - Not a pest Not suitable Larva Adult UV trap Lethbridge (AB) Mamestra configurata Walker Bertha armyworm Pest Suitable Pupa Adult UV trap Lethbridge (AB) Mythimna unipuncta (Haworth) true armyworm Pest Suitable Larva Adult UV trap Edmonton (AB) Noctua pronuba (Linnaeus) winter cutworm Pest Suitable Larva Adult UV trap Lethbridge (AB) Pieris rapae (Linnaeus) small cabbage white Pest Not suitable Pupa Larva Hands Lethbridge (AB) Sideridis rosea (Harvey) rosewing moth Not a pest Not suitable Pupa Adult UV trap Lethbridge (AB) Spaelotis clandestina (Harris) w-marked cutworm Pest Suitable Larva Adult UV trap Lethbridge (AB) Spilosoma virginica (Fabricius) yellow woolly bear Not a pest Not suitable Pupa Adult UV trap Lethbridge (AB) Spodoptera eridania (Stoll) southern armyworm Pest Not suitable - Egg Purchased Benzon Research (PA) Spodoptera exigua (Hübner) beet armyworm Pest Not suitable - Egg Purchased Benzon Research (PA) Spodoptera frugiperda (J. E. Smith) fall armyworm Pest Not suitable Pupa Egg Purchased Benzon Research (PA) Trichoplusia ni (Hübner) cabbage looper Pest Suitable - Egg Purchased Benzon Research (ON) Trichordestra lilacina (Harvey) aster cutworm Not a pest Not suitable Pupa Egg Hands Lethbridge (AB) Xestia c-nigrum (Linnaeus) spotted cutworm Pest Not suitable Larva Adult UV trap Lethbridge (AB) Chapter 3 66 Chapter 3 Figure 3.1. Cotesia vanessae parasitizing an early 4th instar Trichoplusia ni. The ovipositor is inserted and the wings, middle and hind legs are placed upward in a putative oviposition position. 67 Chapter 3 Table 3.2. Lepidoptera pecies that didn’t produce C. vanessae. Lepioptera species Number of larvae that: Survived Died Anticarsia gemmatalis 39 6 Habrosyne scripta 27 3 Hyles euphorbiae 12 5 Lacinipolia renigera 22 3 Lacinipolia sareta 31 3 Pieris rapae 12 27 Sideridis rosea 26 9 Spilosoma virginica 30 0 Spodoptera eridania 19 3 Spodoptera exigua 27 2 Spodoptera frugiperda 5 13 Trichordestra lilacina 19 39 Xestia c-nigrum 17 16 68 Chapter 3 Died producing parasitoids Died of premature death (i.e., before pupating and w witihthooutu tp rpordoudcuicnign gp apraarsaistiotoidisd)s ) Pupated (i.e., survived parasitism) Figure 3.2. Fate of Lepidoptera larvae of species that produced C. vanessae. All individual larvae were parasitized, with the exception of S. virginica that could not be stung, likely because of its pubescence. Parasitism rate (comparing the number of larvae that “died producing parasitoids” with the number of larvae that “pupated”) of each host species was analyzed with Fisher’s exact tests comparing pairs of species (α = 0.05, corrected according to Bonferroni). Significance is indicated by horizontal lines above the boxes. Each “x” refers to a particular species, and the line it stands on indicates the species that didn’t have a significantly different parasitism rates from the species represented by the “x”. 69 Chapter 3 TRIBE SUBFAMILY FAMILY Abagrotis reedi (KX281188) Abagrotis sp. (KX281230) Xestia c-nigrum (KX281223) Spaelotis clandestina (KX281231) Eurois occulta (KX281198) Noctuini 1 Anaplectoides prasina (KX281192) Cryptocala acadiensis (KX281197) Noctua pronuba (KX281212) Lacinipolia renigera (KX281208) Eriopygini Lacinipolia sareta (KX281209) Sideridis rosea (KX281215) Trichordestra lilacina (KX281213) Mamestra configurata (KX281210) Hadenini 1 Anarta trifolii (KX281194) Lacanobia grandis (KX281207) Feltia herilis (KX281203) Noctuinae 1 Feltia jaculifera (KX281204) Actebia balanitis (KX281187) Agrotis vancouverensis (KX281190) Noctuidae 1 Agrotis ipsilon (KX281226) Actebia praecox (CGUKA867-09) Noctuini 2 Euxoa messoria (KX281224) Euxoa tristicula (KX281202) Euxoa ochrogaster (KX281201) Euxoa auxiliaris (KX281199) Euxoa satis (KX281200) Apamea devastator (KX281186) Apamea lignicolora (KX281195) Apameini Apamea sordens (KX281217) Mythimna unipuncta (KX281211) Leucaniini Mythimna litoralis (GBLAA229-14) Helicoverpa zea (KX281206) Heliothinae Heliothis virescens (KX281229) Dargida diffusa (KX281228) Hadenini 2 Spodoptera exigua (KX281220) Noctuinae 2 Spodoptera eridania (KX281219) Prodeniini Spodoptera frugiperda (KX281221) Anticarsia gemmatalis (KX281227) Panopodini Eulepidotinae Spilosoma virginica (KX281216) Arctiini Erebidae Arctiinae Amphipyra tragopoginis (KX281225) Amphipyrini Amphipyrinae Caradrina morpheus (KX281218) Caradrini Noctuinae 3 Anagrapha falcifera (KX281191) Plusiini Autographa californica (KX281196) Noctuidae 2 Trichoplusia ni (KX281222) Plusiinae Chrysodeixis chalcites (KC846709.1) Argyrogrammatini Chrysodeixis includens (KX550951) Figure 3.3. Neighbour-joining tree (Kimura 2-parameter distance) for CO1 sequences of known hosts of C. vanessae and other tested species within Noctuoidea, generated using Mega 5.2 (allowing for visual representation of relative interspecific divergence, but not a robust phylogenetic analysis). The scale bar represents the percent divergence of the Kimura 2-parameter. Bootstrap values indicate the percentage of times, out of 5000, that clades were paired with one another. Bootstrap values < 70 are not shown are these are considered too low to be meaningful. Accession numbers underlined and in bold font refer to sequences obtained from GenBank and BOLD, respectively. Other accession numbers refer to sequences generated from specimens from this study and deposited in GenBank. Colour of species represents C. vanessae parasitism success; i.e., green: excellent host (81-100% parasitism); blue: intermediate host (21-80% parasitism); orange: poor hosts (1-20% parasitism); red: not detected as a host; black: species that were considered reliably reported as hosts but were not tested. 70 Chapter 3 Figure 3.4. Cotesia vanessae total development time on each host species. Because of low statistical power, species represented by fewer than 5 individuals were excluded from analyses. The boxes are delimited by the 25th and 75th percentiles. The upper and lower whiskers indicate Q3 + 1.5 IQR and Q1 – 1.5 IQR, respectively. Medians and means are represented by solid lines and dotted lines, respectively. Significance (Kruskal-Wallis test followed by post hoc Dunn test, α = 0.05 corrected according to Bonferroni) is indicated by horizontal lines above the boxes. Each species is represented by an “x” standing on a line that indicates ot her species with similar parasitism rates. 71 Chapter 3 Figure 3.5. Cotesia vanessae brood sizes on each host species. Because of low statistical power, species represented by fewer than 5 individuals were excluded from analyses. The boxes are delimited by the 25 th and 75th percentiles. The upper and lower whiskers indicate Q3 + 1.5 IQR and Q1 – 1.5 IQR, respectively. Medians and means are represented by solid lines and dotted lines, respectively. Significance (one-way ANOVA followed by post hoc Tukey’s HSD, α = 0.05) is indicated by horizontal lines above the boxes. Each species is represented by an “x” standing on a line that indicates other species with similar brood sizes. 72 Chapter 3 Figure 3.6. Cotesia vanessae average adult individual dry mass per brood (average of 30 or fewer wasps per brood) on each host species. Because of low statistical power, species represented by fewer than 5 individuals were excluded from analyses. The boxes are delimited by the 25 th and 75th percentiles. The upper and lower whiskers indicate Q3 + 1.5 IQR and Q1 – 1.5 IQR, respectively. Medians and means are represented by solid lines and d otted lines, respectively. Significance (one-way ANOVA followed by post hoc Tukey’s HSD, α = 0.05) is indicated by horizontal lines above the boxes. Each species is represented by a “x” standing on a line that indicates other species with similar individual parasitoid mass. 73 Chapter 3 Figure 3.7. Cotesia vanessae estimated brood mass per host (average mass of 30 or fewer wasps per brood x brood size) on each host species. Because of low statistical power, species represented by fewer than 5 individuals were excluded from analyses. The boxes are delimited by the 25th and 75 th percentiles. The upper and lower whiskers indicate Q3 + 1.5 IQR and Q1 – 1.5 IQR, respectively. Medians and means are represented by solid lines and dotted lines, respectively. Significance (Kruskal-Wallis test followed by post hoc Dunn tests, α = 0.05 corrected according to Bonferroni) is indicated by horizontal lines above the boxes. Each species is represented by a “x” standing on a line that indicates other species with similar individual parasitoid mass. 74 Chapter 3 t av A 50 µm B 100 µm C 100 µm D 100 µm E 200 µm F 200 µm Figure 3.8. Immature Cotesia vanessae A. Egg, two days after parasitism within Trichoplusia ni; B. Neonate larva producing teratocytes, five days after parasitism within Trichoplusia ni. Head on the left, anal vesicle on the right, thoracic and first 7 abdominal segments each partly surrounded on their dorsal and lateral sides by a row of cuticular spines projecting backward, t: teratocyte, av: anal vesicle (photo courtesy of Stephanie Harris, Agriculture and Agri-Food Canada, Saskatoon, SK); C. Egg becoming encapsulated by hemocytes, four days after parasitism within Helicoverpa zea; D. First instar larva encapsulated, eleven days after parasitism within Helicoverpa zea larva; E. First instar larvae, seven days after parasitism within Mythimna unipuncta. One larva with open mouth (centre), and the fore half of a killed larva on its right; F. Larva biting a sibling, and a fore half of a younger larva on the bottom right, seven days after parasitism within Mythimna unipuncta. 75 Chapter 3 bg ld gc A B C Figure 3.9. Spodoptera frugiperda A. Live larva 50 days after parasitism; B. Same larva dissected (ld), with bloated gut dissected (bg), and gut content extracted (gc); C. Boxplot showing time to death of parasitized S. frugiperda larvae compared to time to pupation and to adult emergence of non-parasitized individuals (six pupae died in the non-parasitized group). The boxes are delimited by the 25th and 75th percentiles. The upper and lower whiskers indicate Q3 + 1.5 IQR and Q1 – 1.5 IQR, respectively. Medians and means are represented by solid lines and dotted lines, respectively (non-visible mean lines are superposed with median lines). 76 Chapter 4 CHAPTER 4: A REVIEW OF THE MCMORRAN DIET FOR REARING LEPIDOPTERA SPECIES WITH ADDITION OF A FURTHER 39 SPECIES (Published as: Hervet, V. A. D., Laird, R. A., & Floate, K. D. (2016). A review of the McMorran diet for rearing Lepidoptera species with addition of a further 39 species. Journal of Insect Science, 16(1), 1-7) 4.1. Abstract Research on cutworms led to the exploration the use of the McMorran diet to rear lepidopteran species, mainly Noctuidae, under laboratory conditions. This chapter reports the development of 104 lepidopteran species, including 39 species not previously reported in the literature, on this diet. Given its low cost, ease of preparation, and wide species’ acceptance, this diet provides a powerful tool for facilitating Lepidoptera and other insects rearing and research in the laboratory. 4.2. Introduction Laboratory colonies of herbivorous insects are commonly reared on artificial diets to reduce the labor, time, space, and associated costs of growing their host plants. These diets also simplify the synchronization of insect development with the availability of food and can be optimized to increase insect fitness above that of insects reared on natural foods (McMorran, 1965). Furthermore, the nutritional quality of these diets can be manipulated to facilitate research on topics including insect development, entomopathogens, insecticides, and plant resistance factors (George et al., 1960). The McMorran diet was used to rear primarily pestiferous species of Noctuidae. This diet has been used as a suitable media to rear at least 154 species of insects, mainly Lepidoptera (Table 4.1). Adkisson et al. (1960) were the first to use wheat germ as an ingredient in an artificial diet they developed to rear the pink bollworm, Pectinophora 77 Chapter 4 gossypiella (Sauders) (Gelechiidae). This recipe was modified by Vanderzant et al. (1962) to rear the corn earworm, Helicoverpa zea (Boddie), and further modified by Berger (1963) to rear the corn earworm and the tobacco budworm, Heliothis virescens (Fabricius) (Lepidoptera: Noctuidae). McMorran (1965) modified this later recipe to rear the spruce budworm, Choristoneura fumiferana (Clemens) (Lepidoptera: Tortricidae). Grisdale (1973) subsequently added linseed oil as an ingredient to this latter recipe to reduce the incidence of wing deformities observed in other Lepidoptera species. There are publications that provide artificial diet recipes with rearing methods of insects and their relatives and it is not the objective here to review these articles. However, it is important to note the existence of key publications that summarize diet recipes and rearing procedures, such as Smith (1966), Singh (1977), Singh and Moore (1985), and Wong (1972). This chapter provides a list of Lepidoptera that have been reared (i.e., larvae were able to develop) on the McMorran diet in previous studies and in the current study. Comments indicating the suitability of the diet for rearing success are indicated for each species (Table 4.1). During four years of research on Lepidoptera, an increasing number of species that successfully develop on the McMorran diet was serendipitously discovered. However, assessing rearing success of insect species on this diet was not an objective so no data on specific fitness of each species reared are displayed. Nonetheless, the information presented here should greatly ease rearing of Lepidoptera larvae for many studies. The research conducted included rearing of field collected Lepidoptera larvae until either parasitoid egression or until they metamorphosed to adults for moths and parasitoids identification, and investigation of the host range of a parasitoid. The use of McMorran diet was ideal for these investigations. It allowed for a rapid handling of high numbers of field-collected caterpillars of various species. Use of the McMorran diet saved tremendous time, space, and 78 Chapter 4 money because it eliminated the need to grow plants and to develop species-based rearing knowledge as in most cases Noctuidae would develop on this diet. It also facilitated insect maintenance as the food remained good for at least 2 weeks at 4 ᵒC and 5-7 days at room temperature (as per rearing conditions described under Materials and Methods section below), and the rearing cages and cups remained relatively clean until insects completed their development. 4.3. Materials and Methods To compile the list of Lepidoptera previously reported to have been reared on the McMorran diet, research articles reported in the Web of Science, Scopus, Google, Google Scholar citing “McMorran 1965” and “Grisdale 1973” were examined (Table 4.1). For laboratory use laboratory, McMorran diet ingredients were purchased from the Insect Production Services (IPS) (Great Lakes Forestry Centre, Canadian Forest Service, Sault Ste. Marie, ON, Canada). Ingredients were preserved for up to 6 months in air-sealed packages (excluding linseed oil and potassium hydroxide (KOH), which were kept in bottles) at 2.5 ᵒC. Fresh diet was prepared on a weekly basis in our laboratory. This diet was held for up to 2 weeks at 4 ᵒC before use. The diet was prepared as per the recipe provided to us by Insect Production Services, which corresponds to that of Grisdale’s (1973) modification of the recipe developed by McMorran (1965). Ingredients for 2-liter diet: 1,720 ml distilled water, 34.72 g agar, 70 g casein, 70 g sucrose, 61.39 g toasted wheat germ, 20 g Wesson’s salt, 10 ml 4 M KOH, 10 ml linseed oil, 10 ml alphacel, 8 g ascorbic acid, 4.2 g aureomycin, 3 g methyl paraben, 2 g choline chloride, 1ml 37% formalin, 20 ml vitamin solution. Ingredients for 2 l vitamin solution: 2 l water, 2 g nicotinic acid, 2 g calcium pantothenate, 2 g riboflavin, 0.5 g thiamine hydrochloride, 0.5 g pyridoxine hydrochloride, 0.5 g folic acid, 0.04 g biotin, and 0.004 g vitamin B12. To better preserve the vitamins, 200 ml (for 2 l of diet) of water were replaced 79 Chapter 4 by 200 g of crushed ice, which was blended with the hot mixture in a blender for one minute just before adding the vitamin solution. This was implemented in order to reduce temperature of the mixture. When the diet was ready, the head foam was scraped away, and the diet was poured into 22 ml plastic cups. When the diet was set (~30 min after pouring), these cups were refrigerated until use. Considering that the ingredients were purchased already pre-divided for 2-liter batches, it took only one hour to prepare one batch including washing of equipment. Most lepidopteran species were purchased as eggs or recovered from adults collected from UV light traps. Commercial sources were Insect Production Services (Sault Ste. Marie, ON, Canada) and Benzon Research Inc. (Carlisle, PA). Importation of species from the United States was done with import permits issued by the Canadian Food Inspection Agency (Permit No. P-2011-04397, P-2013-02134, and P-2014-02394). Light traps were operated in the city of Lethbridge, Alberta, Canada, during the summer and fall of 2012-2014. Additional species were collected as larvae or eggs on various plants in and near Lethbridge, and near Magrath (Cardston county), Alberta. Eggs were recovered from field-collected adults by placing female moths in Plexiglass cages (40 cm by 34 cm by 30 cm [length x width x height]). A screened window (10 cm in diameter) on each side of the cage provided ventilation. The front of the cage had an opening (15 cm in diameter) with an attached cloth sleeve to provide access into the cage. Females were provided with sucrose crystals, water, and a choice of oviposition materials; i.e., cheese cloth, paraffin film (“Parafilm”), brown paper, paper towel, and fine sand. Eggs on the oviposition substrate were placed into new cages of the same design in which the hatched caterpillars were reared until pupation or until used in experiments. Usually about 50 caterpillars were reared per cage (or 30 for Plusiinae). Eggs obtained commercially were similarly handled. For species maintained for more than one generation, pupae were placed in clean cages of this same design for emergence, mating, and oviposition. 80 Chapter 4 In the laboratory, insects were reared in growth cabinets at 20 ᵒC, 70-80% humidity, and 12:12 (L:D) hours cycle under low incandescent and fluorescent lighting. Caterpillars were provided with blocks of diet (20 cm3) positioned mainly near the sides and corners of the cage. Small pieces of paraffin film (~9cm2) placed beneath and on top of each diet block facilitated its removal, reduced desiccation, and allowed caterpillars to feed while standing on the underside of the upper piece of the paraffin film (Fig. 4.1) as they would under a plant leaf in natural conditions. Fresh diet was added about every 5-7 days when the previous one became too old or dry. The diet had to be replaced about four times to sustain complete larval development of most species reared. For cages with first- to third-instar larvae, and any instar of Plusiinae (i.e., “loopers”), fresh diet blocks were placed as close as possible to the old blocks to facilitate the movement of caterpillars onto the fresh diet as these caterpillars have difficulty moving on the floor of the cage. Blocks of old diet were removed from cages after about 10 days. Variously sized pieces of brown paper (laid flat or slightly crumpled) were placed in the cages of Noctuidae species (except Plusiinae) to provide sheltered areas. This noticeably reduced diurnal activity of the caterpillars and biting of each other. Cannibalism of aggressive species (e.g., bertha armyworm and corn earworm) was further reduced by placing an increasing amount of crumpled brown paper, using bigger cages, having fewer caterpillars in a cage, and adding of a few centimeters of vermiculite on the bottom of the cage (Fig. 4.2). Lepidoptera larvae were also reared individually in translucent plastic (Polar Plastic Ltd.) 8 oz (240 ml) containers (Fig. 4.3). These included caterpillars hatched from eggs in the laboratory or collected outside (same species as these reared in cages). A non-threaded sewing machine was used to ease piercing of holes (0.5 mm in diameter) in the bottom and lid of containers to prevent the accumulation of condensation within. This reduced the incidence of mold on the diet and prevented early-instar larvae from becoming trapped and dying in water droplets. Each cup contained a piece of filter paper (55 mm in diameter) onto 81 Chapter 4 which a block of diet was placed (6.7 or 10 cm3, depending upon the size of the larva), with a piece of paraffin film beneath and on top as described previously. Rearing conditions were as described for caterpillars reared in cages. The caterpillars pupated either in their cages or individual cups. They remained this way under the conditions described above until moth emergence. 4.4. Results and Discussion The results of the literature search and laboratory studies are reported in Tables 4.1 and 4.2. Many studies do not report if the diet used contained linseed oil as per Grisdale’s modification (1973) or any other modification. Table 4.1 reports Lepidoptera species successfully reared on the McMorran diet, whereas Table 4.2 reports species that failed to develop on this diet. Grisdale (1973) reports successfully rearing grasshoppers, Melanoplus spp. (Orthoptera: Acrididae) on the McMorran diet for at least one generation. Rearing successive generations of Lepidoptera on nutritionally unbalanced diets can cause wing deformities (Morris 1967; personal observation), reductions in insect weight, fecundity, longevity, and increased mortality of all life stages (Morris, 1967). The suitability of the McMorran diet for rearing successive generations of the species listed in Table 4.1 typically was not assessed. Some species were unwilling or unable to feed on McMorran diet as early instars but fed on the diet in later instars to complete their development; i.e., Hyles euphorbiae (L.) (Sphingidae) (n = ~17; most adults had deformed wings) and Polychrysia esmeralda (Oberthür) (Noctuidae) (n = ~60; no deformed wings observed). These are indications that further modification of the McMorran diet is needed to maintain laboratory colonies of these species. Minor modifications to the McMorran diet can enhance rearing success. For example, Syme and Green (1972) reared larvae of European pine shoot moth, Rhyacionia buoliana (Denis & Schiffermüller) (Tortricidae) with the addition of 1 ml soya bean oil per 100 82 Chapter 4 ml of McMorran diet. Gardiner (1970) reared to adults larvae of 49 identified species (and additional unidentified species) of Cerambycidae (Coleoptera), 17 of them from the egg, by adding 5 ml of dried pulverized host plant material to 100 ml McMorran diet. Similarly, Trudel successfully reared immature stages of the white pine weevil, Pissodes strobi Peck (Coleoptera: Curculionidae), and maintained adults, by adding 5% pulverized white pine bark to the McMorran diet. Other diets similar in composition to the McMorran diet include that of Adkisson et al. (1960), Vanderzant et al. (1962), and the “Vanderzant-Adkisson special wheat germ diet” (marketed by MP Biomedicals, headquartered in Santa Ana, CA), which have also been used to rear a wide range of Lepidoptera and some Coleoptera species. This ability has been attributed to their wheat germ base (Cohen, 2004). Research that requires maintenance of laboratory colonies over multiple generations or aiming to study species performance should try to determine suitability of the McMorran diet for specific species, which may require adjustments of the recipe and further investigation into insect performance, to ensure maintenance of healthy colonies. Nonetheless, it is shown here that the McMorran diet can be used to facilitate the laboratory rearing of numerous species, both in cages and in 240 ml containers, and can likely be used to rear many more species than those reported here. 83 Table 4.1. Species of Lepidoptera reared (i.e., were able to grow) on McMorran diet as reported in the literature and from the current study. Species Reference Comments Erebidae (Arctiidae) Estigmene acrea (Drury) (Barber et al., 1993); This study Reared from egg to at least 3rd instar (Barber 1993); [Saltmarsh caterpillar] Partly developed larvae reared to adults (n = 3) (this study) Hyphantria cunea (Drury) (Morris, 1967) Reared over one generation and a partial second generation, severe [Fall webworm] wing deformities, degeneration Lymantria monacha (Linnaeus) (Grijpma et al., 1987) Reared for at least one full generation [Nun moth] Orgyia antiqua (Linnaeus) (Grant, 1977) Likely at least one full generation [Rusty tussock moth] Orgyia cana Edwards (Grant, 1977) Likely at least one full generation Orgyia definita Packard (Grant, 1977) Likely at least one full generation [Definite tussock moth] Orgyia leucostigma (J.E. Smith) (Grisdale, 1973; Percy et al., Reared over multiple generations [Whitemarked tussock moth] 1971) Orgyia pseudotsugata (McDunnough) Non-modified (Grisdale, 1973; Reared over multiple generations [Douglas-fir tussock moth] Morris, 1970) and modified recipe (Lyon et al., 1966) Spilosoma congrua Walker (Barber et al., 1993) Reared from egg to at least 3rd instar [Agreeable tiger moth] Spilosoma virginica (Fabricius) (Barber et al., 1993); This study Reared from egg to adults (n = ca. 200) [Yellow woollybear] Gelechiidae Filatima obscuroocelella (Chambers) This study Partly developed larvae reared to adults (n = 7) Geometridae Lambdina fiscellaria fiscellaria (Otvos et al., 1973) Reared for at least one full generation (Guenée) [hemlock looper] Chapter 4 84 Operophtera bruceata (Hulst) (Ives et al., 1980) Reared from egg to pupa [Bruce spanworm] Operophtera brumata (Linnaeus) (Feeny, 1968) Reared at least from eggs to pupae; modified recipe [Winter moth] Lasiocampidae Malacosoma disstria Hübner (Grisdale, 1973; Stairs, 1965) Reared at least for one full generation [Forest tent caterpillar] Noctuidae Abagrotis sp.1 This study 2nd instars reared to adults (n = ca. 80). Abagrotis reedi Buckett This study Reared from egg to adult (n = ca. 50) Actebias balanitis (Grote) This study Reared from egg to adult (n = ca. 20) Actebia fennica (Tauscher) ? Mentioned on the IPS website (when visited in 2016) [Black army cutworm] Agrotis ipsilon (Hufnagel) (Barber et al., 1993); This study Reared for two generations (n = ca. 100) [Black cutworm] Agrotis vancouverensis (Grote) This study Reared from egg to adult (n = ca. 100) [Vancouver dart] Anagrapha falcifera (Kirby) (Barber et al., 1993); This study Reared from egg to adult (n = ca. 80) [Celery looper] Anaplectoides prasina (Denis & This study Reared from egg to adult (n = ca. 20) Shciffermüller) [Green arches moth] Anaplectoides pressus (Grote) (Barber et al., 1993) Reared from egg to at least 3rd instar [Dappled dart] Anarta (Hadula) trifolii (Hufnagel) This study Reared from egg to adult (n = ca. 100); few specimens with deformed [Clover cutworm] wings Anticarsia gemmatalis (Hübner) This study Reared from egg to adult (n = ca. 70) [Velvetbean caterpillar] Apamea devastator (Brace) This study Reared from egg to adult (n = ca. 2) [Glassy cutworm] Chapter 4 85 Apamea lignicolora (Guenée) This study Reared from egg to adult (n = ca. 20) [Wood-colored Apamea] Apamea sordens (Hufnagel) This study Partly developed larvae reared to adults (n = 5) [Rustic shoulder-knot] Autographa bimaculata (Stephens) This study Last instar larvae reared to adults (n = 2) [Two-spotted looper] Autographa californica (Speyer) This study Reared from egg to adults (n = ca. 200) [Alfalfa looper] Autographa flagellum (Walker) (Barber et al., 1993) Reared from egg to at least 3rd instar [Silver whip] Autographa precationis (Guenée) (Barber et al., 1993) Reared from egg to at least 3rd instar [Common looper] Caradrina morpheus (Hufnagel) This study Reared from egg to adults (n = ca. 100) [Mottled rustic moth] Caradrina multifera (Walker) (Barber et al., 1993) Reared from egg to at least 3rd instar [Speckled rustic moth] Cerastis salicarum (Walker)2 (Barber et al., 1993) Reared from egg to at least 3rd instar [Willow dart] Chrysodeixis includens (Walker) (Grisdale, 1973); This study Reared from egg to adults (n = ca. 60) [Soybean looper] Crocigrapha normani (Grote) (Barber et al., 1993) Reared from egg to at least 3rd instar [Climbing cherry cutworm] Cryptocala acadiensis (Bethune) (Barber et al., 1993); This study Reared from egg to adults (n = ca. 30) [Catocaline dart] Cucullia intermedia Speyer This study Three last instars reared to adults (n = 1); the younger instars didn’t [Goldenrod cutworm] feed very willingly on the diet Dargida diffusa (Walker) This study Partly developed larvae reared to adults (n = ca. 20) [Wheat head armyworm] Diachrysia aereoides (Grote) This study Partly developed larvae reared to adults (n = 5) [Dark-spotted looper] Chapter 4 86 Diarsia rubifera (Grote) (Barber et al., 1993) Reared from egg to at least 3rd instar [Red dart] Diarsia jucunda (Walker) (Barber et al., 1993) Reared from egg to at least 3rd instar [Smaller pinkish dart] Eueretagrotis attenta (Grote) (Barber et al., 1993) Reared from egg to at least 3rd instar [Attentive dart] Eupsilia tristigmata (Grote) (Barber et al., 1993) Reared from egg to at least 3rd instar [Brown fruitworm] Eurois occulta (Linnaeus) This study Reared from egg to adults (n = ca. 60) [Great brocade] Euxoa auxiliaris (Grote) This study Reared from egg to adults (n = ca. 50) [Army cutworm] Euxoa messoria (Harris) This study Reared for two generations (n = ca. 150) [Darksided cutworm] Euxoa ochrogaster (Guenée) This study Reared for three generations (n = ca. 300); ca. 5% adults had deformed [Redbacked cutworm] wings Euxoa satis (Harvey) This study Reared from egg to adult (n = at least 1) Euxoa tristicula (Morrison) This study Reared for five generations (n = ca. 600) [Early cutworm] Feltia herilis (Grote) This study Reared from egg to adult (n = ca. 30) [Herald dart] Feltia jaculifera (Guenée) This study Reared for two generations (n = ca. 60) [Dingy cutworm] Fishia illocata (Walker) (Barber et al., 1993) Reared from egg to at least 3rd instar [Wandering brocade] Helicoverpa zea (Boddie) (Grisdale, 1973); This study Reared for two generations (n = ca. 80) [Corn earworm] Heliothis virescens (Fabricius) This study Reared for two generations (n = ca. 80) [Tobacco budworm] Chapter 4 87 Homorthodes furfurata (Grtote) (Barber et al., 1993) Reared from egg to at least 3rd instar [Northern scurfy quacker moth] Hyppa xylinoides (Guenée) (Barber et al., 1993) Reared from egg to at least 3rd instar [Cranberry cutworm] Lacanobia grandis (Guenée) (Barber et al., 1993); This study Reared from egg to adult (n = ca. 40) [Grand arches moth] Lacanobia radix (Walker) (Barber et al., 1993) Reared from egg to at least 3rd instar [Garden arches moth] Lacinipolia renigera (Stephens) This study Reared from egg to adult (n = ca. 40) [Bristly cutworm] Lacinipolia vicina (Grote) This study Reared from egg to adult (n = ca. 50) Leucania multilinea Walker (Barber et al., 1993) Reared from egg to at least 3rd instar [Many-lined Wainscot moth] Mamestra configurata Walker (Grisdale, 1973);This study Reared from egg to adult (n = ca. 50); last instar was highly cannibalistic [Bertha armyworm] Melanchra picta (Harris) This study Partly developed larvae reared to last instar (n = 3); two of them died [Zebra caterpillar] from parasitoids, the last one from disease. Melanchra pulverulenta (Smith) (Barber et al., 1993) Reared from egg to at least 3rd instar Morrisonia latex (Guenée) (Barber et al., 1993) Reared from egg to at least 3rd instar [Fluid arches moth] Mythimna unipuncta (Haworth) (Barber et al., 1993); This study Reared for over five generations (n = ca. 1000) [True armyworm] Noctua pronuba (Linnaeus) This study Reared from egg to adult (n = ca. 20); few specimens with deformed [Large yellow underwing] wings Orthosia revicta (Morrison) (Barber et al., 1993) Reared from egg to at least 3rd instar [Rusty whitesided caterpillar] Peridroma saucia (Hübner) (Birch et al., 1976) Reared for four generations [Variegated cutworm] Phlogophora iris Guenée (Barber et al., 1993) Reared from egg to at least 3rd instar [Olive angle shades moth] Chapter 4 88 Plusia putnami Grote This study Last instars reared to adults (n = 2) [Lempke’s gold spot] Pyrrhia exprimens (Walker) (Barber et al., 1993) Reared from egg to at least 3rd instar [Purple-lined sallow moth] Sideridis rosea (Harvey) This study Reared from egg to adult (n = ca. 20); most adults with deformed wings [Rosewing moth] Spaelotis clandestina (Harris) This study Reared from egg to adult (n = ca. 20) [W-marked cutworm] Spodoptera eridania (Stoll) This study Reared for three generations (n = ca. 200) [Southern armyworm] Spodoptera exigua (Hübner) This study Reared for two generations (n = ca. 200) [Beet armyworm] Spodoptera frugiperda (J.E. Smith) This study Reared for two generations (n = ca. 100) [Fall armyworm] Trichoplusia ni (Hübner) (Grisdale, 1973); This study Reared for at least three generations, multiple times (n >10,000) [Cabbage looper] Trichordestra lilacina (Harvey) This study Reared from egg to adult (n = 12) [Aster cutworm] Xestia c-nigrum (Linnaeus) (Barber et al., 1993); This study Reared from egg to adult (n = ca. 40) [Setaceous Hebrew character] Xestia smithii (Snellen) (Barber et al., 1993) Reared from egg to at least 3rd instar [Smith’s dart] Xylena nupera (Lintner) This study Partly developed larva reared to adults (n = 1); wings of adult deformed [American swordgrass moth] Zanclognatha pedipilalis (Guenée) (Barber et al., 1993) Reared from egg to at least 3rd instar [Grayish zanclognatha] Pieridae Pieris napi (Linnaeus) (Barber et al., 1993) Reared from egg to at least 3rd instar [Green-veined white] Chapter 4 89 Pyralidae Dioryctria reniculelloides (Hamel, 1977) Field collected larvae to? Mutuura & Munroe [Spruce coneworm] Dioryctria abietivorella (Grote) (Trudel et al., 1995) Fourth instars to adults [Fir coneworm] Ostrinia nubilalis (Hübner) (Grisdale, 1973) Reared for at least one full generation [European corn borer] Saturnidae Hyalophora cecropia (Linnaeus) (Grisdale, 1973) Reared for at least one full generation [Cecropia moth] Sphingidae Daphnis nerii (Linnaeus) (Retnakaran et al., 1985) Reared over 30 generations [Oleander hawk-moth] Hyles galii (Rottemburg) This study Penultimate instar to pupae (n = 5) [Bedstraw hawk-moth] Manduca sexta (Linnaeus) ? Mentioned on the IPS website (when visited in 2016) [Tobacco hornworm] Tortricidae Acleris gloveranus (Walsingham) (Gray et al., 1993) Field collected caterpillars reared until parasitoid emergence [Western blackheaded budworm] Acleris variana (Fernald)3 ? Mentioned on the IPS website (when visited in 2016) [Eastern blackheaded budworm] Archips cerasivorana (Fitch) (Grisdale, 1973) Reared for at least one full generation [Uglynest caterpillar] Choristoneura conflictana (Walker) (Grisdale, 1973) Reared for at least one full generation [Large aspen tortrix] Choristoneura fumiferana (Clemens) (Grisdale, 1973; McMorran, 1965) Reared over multiple generations [Spruce budworm] Chapter 4 90 Choristoneura occidentalis Freeman (Clancy, 1991) Reared over multiple generations; modified ingredients [Western spruce budworm] Choristoneura pinus pinus Freeman (Allen et al., 1958) Reared for multiple generations, degeneration observed [Jack pine budworm] Choristoneura rosaceana (Harris) (Barber et al., 1993) Reared to at least 3rd instar [Obliquebanded leafroller] 1 Species called “Abagrotis orbis (Grote)” in Hervet et al. (2016b). Further investigations revealed that it could be another species. A CO1 sequence of this species was deposited into GenBank (GenBank accession number: KX281230) (http://www.ncbi.nlm.nih.gov/genbank/) for later reference. 2 Species misspelled as “Cerastsis salicarum” in Hervet et al. (2016b) 3 Species not listed in Hervet et al. (2016b) Chapter 4 91 Chapter 4 Table 4.2. Lepidoptera species that did not successfully develop on McMorran diet in the present study. Species Comments Drepanidae Habrosyne scripta (Gosse) First instars (n = ca. 60) [Lettered Habrosyne] Gelechiidae Syncopacma sp. Partly developed larvae collected on Lupinus argenteus Pursh (Fabaceae) (n = ca. 10) Hesperiidae Epargyreus clarus (Cramer) Partly developed larvae (n = 2) [silverspotted skipper] Nymphalidae Aglais milberti (Godart) Partly developed larvae (n = ca. 60) [Milbert’s tortoiseshell] Coenonympha tullia (Müller) First instars (n = ca. 15) [Common ringlet] Limenitis arthemis (Drury) Partly developed larvae (n = 3) [White admiral] Papilionidae Papilio Canadensis (Rothschild & Jordan) First instars (n = 1) [Canadian tiger swallowtail] Papilio multicaudata Kirby First instars (n = 1) [Two-tailed swallowtail] Pieridae Colias philodice Godart Partly developed larvae (n = ca. 20) [Clouded sulphur] Pieris rapae (Linnaeus) Partly developed larvae (n = ca. 50) [Small white] Saturniidae Hemileuca eglanterina (Boisduval) Partly developed larvae (n = 2) [Sheep moth] 92 Chapter 4 Figure 4.1. Set up used to rear caterpillars in cages on the McMorran diet (on this picture: Trichoplusia ni). 93 Chapter 4 Figure 4.2. Set up for rearing cannibalistic species in cages. This picture shows a corner of a 60 x 47 x 44 cm Plexiglas cage used to rear 50 Mamestra configurata larvae (pupa visible in vermiculite). Three caterpillars died of disease, the others pupated, but none were cannibalized. 94 Chapter 4 Figure 4.3. Polar Plastic Ltd. cup (240 ml) used to rear individual caterpillars (here a Euxoa auxiliaris). 95 Chapter 5 CHAPTER 5: MULTI-TROPHIC EFFECTS OF PROTEIN-RESTRICTED DIET ON THE DEVELOPMENT AND FITNESS OF THE CABBAGE LOOPER, TRICHOPLUSIA NI (LEPIDOPTERA: NOCTUIDAE), AND ITS PARASITOID COTESIA VANESSAE (HYMENOPTERA: BRACONIDAE) “It’s a kind of scientific integrity, a principle of scientific thought that corresponds to a kind of utter honesty–a kind of leaning over backwards. For example, if you’re doing an experiment, you should report everything that you think might make it invalid–not only what you think is right about it: other causes that could possibly explain your results […]. Details that could throw doubt on your interpretation should be given, if you know them. […] If you make a theory, for example, and advertise it, or put it out, then you must also put down all the facts that disagree with it, as well as those that agree with it. […] The idea is to try to give all of the information to help others to judge the value of your contribution; not just the information that leads to judgment in one particular direction or another.” Richard P. Feynman (1999). Cargo Cult Science. 5.1. Abstract Understanding the trophic effects of unbalanced diets on the parasitoids of crop-pest insects can improve our understanding of parasitism in the field. In this study, casein levels in an artificial diet were manipulated to test the effects of a diet primary metabolite: protein, on a Lepidoptera larva, the cabbage looper Trichoplusia ni, and on its parasitoid, Cotesia vanessae. Host larvae reared on diets with less than 25.5 g/L of protein exhibited delayed growth and lighter pupae. Similarly, parasitoids emerging from hosts reared on diets containing less than 25.5 g/L of protein exhibited delayed growth in the host and smaller broods. This shows that host development is linked to the protein content of its diet. Parasitoid fitness is likely linked to its host’s diet protein content through variation of host quality. However, the percentage of hosts successfully parasitized and the mass of individual parasitoids were not affected by diet quality. An increase in cabbage looper death was observed only for individuals being reared on the most protein-poor diet (8.1 g/L of protein), which suggests that cabbage loopers can tolerate high variation in protein availability. As a compensatory mechanism, cabbage loopers increasing their food consumption on diets of lower protein content. Food consumption of cabbage loopers was also higher in parasitized individuals (only on high- 96 Chapter 5 protein diets), likely to compensate for nutrients lost to parasitoids. No parental effect of hosts’ diet quality on parasitoids was found: offspring of parasitoids that developed on caterpillars reared on high-protein diet and offspring of parasitoids that developed on caterpillars reared on the most protein-poor diet showed no difference of development time, brood size, individual parasitoid mass, and brood mass (offspring were reared on caterpillars fed the same high-protein diet). The results also show that C. vanessae can produce a large variety of brood sizes and individual parasitoid masses, and that average individual parasitoid mass is not affected by the quality of the food of its host. These aspects could contribute to the parasitoid success in different environments. 5.2. Introduction The quality of diet for herbivores affects not only their fitness, but also the fitness of members of higher trophic levels. Most previous studies have investigated the effect of secondary metabolites; e.g., glucosinolates, terpenes, and toxins, in the food of insect herbivores on higher trophic levels (Hopkins et al., 2009; Katsanis et al., 2016; Ponsard et al., 2002; Price et al., 1980), but few studies have investigated aspects of nutritional balance by primary metabolites in tri-trophic contexts involving parasitoids (Bloem et al., 1990; Pimentel, 1966). A key component of insect diet is nitrogen, predominantly from proteins. In theory, any nutrient can limit growth and fitness, but certain nutrients may never be limiting because of their excess in natural foods (Harrison et al., 2012). Proteins are often limiting in the natural diets of herbivores (Mattson Jr., 1980; Scriber, 1984; White, 1993). For example, small increases of protein in artificial diets, relative to that of natural foods, often increase the fitness of herbivorous insects (Huberty et al., 2006; McMorran, 1965; White, 1993). It has been hypothesized that if nitrogen is truly a limiting factor in the food of herbivores, natural selection should result in the adoption of strategies to maximize their intake of nitrogen-rich 97 Chapter 5 food sources (Mattson Jr., 1980; Milton, 1978). Indeed, “early-season feeders” are species that evolved to synchronize their main feeding stages with the occurrence of nitrogen-rich sprouts and seedlings (Baker, 1972; McNeil et al., 1978; Schwertzer, 1979). The preference of early-season feeding Noctuidae (e.g., cutworms) for seedlings and buds makes them particularly damaging as pests of crops. The effect of fertilizer to elevate levels of nitrogen in plants and subsequently the fitness of herbivorous insects is well known (Waring et al., 1992). However, an enhanced understanding of the ecology of crop-pest insects also requires knowledge of how variation in nitrogen content in the food of herbivorous insects affects the fitness of their natural enemies. A number of studies have measured the fitness of insects and of their parasitoids by looking at development time, survival rates, length of body or of body part, body mass, female egg load, adult longevity, mating success, number of hosts parasitized, time spent searching for hosts, efficiency in exploiting host patches, etc. Body size (as measured by its length or mass) is the most common variable used by researchers to assess the fitness of parasitoids (Roitberg et al., 2001). It has been shown that body size strongly relates with other metrics of fitness, such as longevity and fecundity of parasitoids and other organisms (Bennett et al., 1998; Bloem et al., 1990; Denno et al., 1985; Gao et al., 2016; Godfray, 1994; Karowe et al., 1989; Kazmer et al., 1995; López et al., 2009; Nicol et al., 1999; Pimentel, 1966; Roff, 1986; Salt, 1941; Vet et al., 1994). In the current study, it was assumed that insects of larger body size had higher fitness, but a number of other fitness parameters were also considered. Here, the fitness consequence of nitrogen-poor diets of an herbivorous insect on its parasitoid was examined in a laboratory experiment. Specifically, I manipulated the protein content of an artificial diet, the McMorran diet (McMorran, 1965), on which I reared cabbage loopers, Trichoplusia ni (Hübner) (Lepidoptera: Noctuidae), to serve as hosts for the parasitoid Cotesia vanessae (Reinhard) (Hyenoptera: Braconidae). Diets containing different 98 Chapter 5 amounts of protein were used to test the following hypotheses: 1) Because protein-restricted diets lead to reduced fitness for herbivores, decreasing proteins in McMorran diet should lead to a decrease in fitness and an increase of larval development time for T. ni; 2) Because parasitoid larvae necessarily derive all of their nutrients from their hosts, parasitoids should have highest fitness and quickest developmental time when developing in hosts reared on high-quality diets; 3) Because the resources provided by a host are finite and because parasitoid larvae egress in synchrony from the host, parasitoid larvae developing within the same host are expected to share among themselves all available resources thus should exhibit scramble competition; 4) The metabolism of parasitods is highly efficient (i.e., a high proportion of the food ingested by the larva is transformed into parasitoid tissues, as illustrated by the small amount of meconium expelled by the parasitoid prior to pupating (Fig. 2.4), which is the only waste accumulated during larval development (Marian et al. 1982; Yu et al. 2008)), therefore, if there is indeed scramble competition, then the total mass of parasitoids produced by a host (= mass of the brood) should remain relatively constant and not be correlated with the number of parasitoids in the host (= size of the brood), when hosts develop on diets of similar quality. 5) Considering the tolerance to low-quality diets observed in other polyphagous Noctuidae (Bloem et al., 1990; Karowe et al., 1989), the decrease of fitness of T. ni and that of C. vanessae are expected to be low or negligible on diets with small protein-restrictions, but significant on diets with high protein-restriction; 6) Because insects in cocoon/pupae do not feed, the development time of T. ni and C. vanessae at these stages should not be affected by diet quality; 7) Because T. ni likely needs to ingest a minimum amount of protein in order to complete its development, T. ni should increases its food consumption on protein-restricted diets; 8) Because parasitoids consume nutrients from their host, to compensate for this loss it is expected that parasitized T. ni larvae would consume more food than non-parasitized larvae; 9) As nutrients used to produce eggs are ingested by the mother, there could be an inter-generational effect in which the fitness of 99 Chapter 5 the progeny of parasitoids from hosts that developed on low-quality diets is lower than these from hosts that developed on higher quality diets, as has been found in previous studies (Bonduriansky et al., 2007; Einum et al., 1999; Gould, 1988; Hafer et al., 2011; Marshall et al., 2006; Wellington, 1976, 1977). 5.3. Materials and methods 5.3.1. Studied insects A thelytokous strain of the gregarious larval koinobiont endoparasitoid Cotesia vanessae was used in the experiment. These individuals descended from parasitoids originally collected in 2009 in southern Ontario, Canada (Hervet et al., 2014). Females lay several eggs in the body of their hosts in a single oviposition event. The eggs hatch into larvae that feed on their host’s haemolymph, and likely on fat tissues too, as do other Cotesia species (Harvey et al., 2008). When larvae reach maturity, they egress in synchrony from the sides of their host. If the host reaches larval maturity before the parasitoids do, the host ceases any activity until parasitoid egression, and never pupates. Immediately upon egression, larvae spin cocoons that collectively form a cocoon mass surrounded by frothy silk. In most cases, the host regains some movement immediately upon parasitoid egression, often crawls a few centimeters and dies within a day. Adult parasitoids usually emerge in synchrony from the cocoon mass. The hosts of C. vanessae are Lepidoptera larvae in the family Noctuidae, in tribe Nymphalini (Nymphalidae), and potentially in other taxa as well (Hervet et al., 2014). The host used in this experiment is the cabbage looper Trichoplusia ni, a polyphagous pest of vegetables in North America (Capinera, 2001). Cabbage looper eggs were obtained from Insect Production Services (IPS), Natural Resources Canada, Sault Ste. Marie, Ontario, Canada. To facilitate rearing and to allow for protein manipulation in their diet, cabbage loopers were reared on artificial diet. 100 Chapter 5 5.3.2. Diet preparation The artificial diet was prepared as indicated in Hervet et al. (2016b), as per McMorran’s recipe (1965) and Grisdale’s modification (1973). Diet ingredients were purchased from IPS. In this artificial diet, the two main sources of nitrogen were casein and wheat germ. As casein is a pure protein while wheat germ contains other nutrients (USDA, 2015), the amount of casein but not that of wheat germ was modified. Diets with six different amounts of protein (Table 5.1) were used, with diet #1 being considered the control as it contains the original amount of casein (i.e., 35 g/L) as per McMorran’s recipe (McMorran, 1965). Because dry wheat germ contains about 26.07 g protein /100 g (USDA, 2015), and because there is 30.695 g/L wheat germ in McMorran diet, each of the six diets contain about 8 g/L protein from wheat germ (Table 5.1). Diet #1 (containing 43 g/L of protein) and diet #6 (containing 8.1 g/L of protein) respectively contain greater and lower amounts of protein than that of the natural foods of cabbage looper. All other diets (diets #2 to #5) contain similar amounts of protein to known host-plants; e.g., Brussel sprouts, broccoli, cauliflower, as well as leaves of cabbages, spinach, lettuce, watercress, and dandelion (Capinera, 2001; USDA, 2015). Because potassium hydroxide (KOH) is used in the McMorran diet to dissolve the casein, correspondingly reduced amounts of KOH were used in the preparation of diets with reduced amounts of casein (Table 5.1). To avoid artificially increasing the concentration of all other diet components, the same mass of extracted casein + KOH was replaced by pure cellulose (purchased as Alphacel (or Alpha-CelTM) from MP Biomedicals, Santa Ana, California, USA). Cellulose is already an ingredient in the McMorran diet and is the main structural component of the cell walls of plants, thus a natural component of T. ni ‘s natural diet; e.g., leaves of the cilion cabbage, Brassica oleracea L. variety capitata L. (Brassicaceae), which is a host plant of T. ni, are about 8% cellulose (Komolka et al., 2012). Because caterpillars cannot 101 Chapter 5 digest cellulose (Martin, 1983; Martin, 1991), increasing its amount in the diet was an appropriate way of maintaining the concentration of the non-casein components. Fresh diet blocks were kept in sealed plastic bags in a Styrofoam cooler in a 4 °C refrigerator until needed. 5.3.3. Insect handling Upon receipt, Trichoplusia ni eggs were placed at room temperature and hatching was monitored. On the first day of egg hatching, neonate caterpillars were placed individually in translucent plastic containers (240 ml, Polar Plastic Ltd.) (Fig. 4.3 (Chapter 4)). Filter paper (55 mm in diameter) was placed at the bottom of each container, on top of which was placed a 6.7 cm3 block of diet. On top and beneath each diet block, squares (9 cm2) of paraffin film (Parafilm®) were placed to slow evaporation and allow the caterpillars to stand beneath the upper film while feeding, as they would under a leaf in natural conditions. The blocks of diet were replaced every ~5 days as they became too old (turned from yellow to orange) or dry. The containers were placed in shuffled locations in the growth cabinet every time the diet was replaced. Small holes were pierced on the bottom and on the lids of the containers beforehand to reduce the humidity, to avoid the development of mould on the diet and caterpillar frass as well as the formation of droplets within the containers’ walls that are death traps for neonate caterpillars. Throughout the experiment the insects were reared in a growth cabinet at 70 to 80% humidity and a 12:12 h light-dark cycle under low incandescent and fluorescent lighting. The temperature of the cabinet was adjusted so that the insides of the rearing containers were at a constant 20 °C, monitored by temperature loggers placed inside of two containers. Caterpillars were either allowed to complete development to adults or were parasitized by C. vanessae. For the latter group, caterpillars that recently molted to their 4th instar were exposed to C. vanessae until the parasitoid’s ovipositor was inserted once. It was 102 Chapter 5 previously determined that this stage showed the best success for C. vanessae (higher host survival rate, larger parasitoid brood size, and higher attractiveness to parasitoids compared to earlier stages; and, higher rate of successful parasitism and lower physical resistance to parasitoid attacks of hosts compared to later stages). Each oviposition event lasted approximately 10 seconds. For consistency, caterpillars were not kept if parasitoids inserted their ovipositors more than once, or if there was suspicion that they might have just probed and rejected the host; i.e., if the thrust lasted less than one second, and if the parasitoid didn’t put its wings, mid- and hind lend upward, which was considered a sign of oviposition (Fig. 3.1). These two groups (i.e., parasitized and non-parasitized) were themselves subdivided into 6 subgroups, which represented the diets containing 6 different amounts of protein (Table 5.1). On each diet, a target number of 45 caterpillars were allowed to develop to adults and a target number of 45 were parasitized (final n values were between 39 and 45, with 10 of the 12 treatment combinations having n = 45; Table 5.2). For logistical reasons, 10 and 35 replicates of each treatment were initiated on February 28th and June 17th, 2014, respectively. Supplementary caterpillars were reared on the diet containing 8.1 g/L of protein due to the high rate of “premature death” (i.e., died as larvae and without producing parasitoids) on this particular diet. 5.3.4. Measurements 5.3.4.1. Data inclusion / exclusion A small number of data points were excluded due to minor errors during data collection. Sample sizes for the same treatments indicated in boxplots may vary between figures because within individuals a missed data point for one variable did not always impede collection of another variable (e.g., failing to record development time of an individual did not hinder the measurement of its mass). No outliers were excluded from the analyses. 103 Chapter 5 5.3.4.2. Measurements for non-parasitized T. ni The effect of diet on T. ni was assessed by measuring the development time and survival of larvae, and the mass of pupae (male, female). In order to allow for the pupae exoskeletons to harden before manipulation and to make consistent measurements, pupae were removed from their cocoons the day following pupation. They were then weighed using a 1 mg precision balance and sexed using morphological features (Fig. 5.1). Individuals that did not reach pupation (i.e., died of premature death or produced parasitoids) were neither sexed nor weighed. 5.3.4.3. Variables measured for C. vanessae The effect of host diet on C. vanessae was assessed by measuring the development time inside the host (eggs then larvae) and inside the cocoon mass (prepupae then pupae), the number of parasitized hosts from which parasitoid larvae egressed, the number of adult parasitoids produced per host, and the dry mass of both individual parasitoids and parasitoid broods. Brood mass was assessed using an average individual mass calculated from the mass of 30 (or fewer depending on availability) intact adults haphazardly chosen from each brood and weighed simultaneously. To test for an inter-generational effect of host quality, the fitness of F2 parasitoids whose mothers developed in hosts fed high- and low-quality diets were compared. Specifically, 14 parasitoids (from different mothers) that developed on hosts fed the control diet (containing 43 g/L of protein) and 14 parasitoids that developed on hosts fed the diet containing the lowest amount of casein (containing 8.1 g/L of protein) each inserted their ovipositor once into one early 4th instar T. ni. These caterpillars were reared on the control diet in the same conditions as described above. The development and fitness of these F2 parasitoids were measured using the same criteria as for the F1 parasitoids described above. 104 Chapter 5 5.3.4.4. Measurement of food eaten by T. ni The amount of food consumed by caterpillars was also measured to help estimate the expected relative damage of noctuid larvae on high- versus low-quality food sources, and of parasitized versus non-parasitized caterpillars. Knowing the extent of feeding in these different situations would help plan integrated pest management strategies, particularly in regard to the use of C. vanessae to control noctuid pests, given that the parasitized pests will not be killed right away and may continue to feed. To estimate the amount of food consumed by caterpillars, the mass of moist diet blocks was measured just prior to being given to caterpillars using a 1 mg precision scale. After 5 to 7 days the partly eaten diet blocks were removed from the experimental containers and placed in individual 57 x 89 cm paper envelopes, which were placed in a dryer for 12 to 14 months (the samples were likely dry after a few days but the unit was also used for storage) before being weighed again using the same scale. The dry diet was weighed immediately after being removed from the dryer to prevent re-hydration. The mass lost between the two weighing events was of two natures: diet eaten by caterpillars and drying. Because I didn’t know how much mass of each of ingredient would be lost by drying, I measured it experimentally using 90 diet blocks (15 of each of the 6 diet types) of about 22 ml from the same diet batches used to feed the caterpillars in the experiment. These diet blocks were not fed to caterpillars but were manipulated in the same time and in similar ways as the experiment to mimic experimental conditions. The percent mass lost by drying of these blocks was found to be significantly different between diet types (F5, 84 = 82.86, P < 0.001), which corresponded to the mass of KOH used in these diets. The results of this manipulation were used to estimate the dry mass of food consumed by each caterpillar in the experiment. 105 Chapter 5 5.3.4.5. Data analyses Data for non-parasitized T. ni were analyzed using two-way ANOVA tests with diet and sex as explanatory variables and larval development time, pupal mass, and food eaten as response variables. Data for parasitized T. ni were analyzed using a one-way ANOVA with diet as the explanatory variable and food eaten as response variable (sex of host was not a variable because T. ni were sexed based on pupal morphology, and since C. vanessae kills its hosts before they pupate, parasitized individuals were not sexed). Data for C. vanessae were analyzed using one-way ANOVA tests with diet as the explanatory variable and development time in host, development time in cocoon, total development time (i.e., in host + in cocoon), brood size (i.e., number of parasitoids produced per host), average individual parasitoid dry mass, and brood mass (i.e., estimated dry mass of all the parasitoids from a brood) as response variables. ANCOVA tests were used to investigate the effects of diet type and development time within the host on cocoon development time, the effects of diet type and brood size on individual parasitoid mass, the effects of diet type and brood mass on individual parasitoid mass, the effects of diet type and brood size on the mass of broods, the effect of diet type and brood size on development time in the host, and to investigate the effect of diet type and brood size on food consumption. Each of these tests provide evidence to support or not support hypotheses mentioned in the Introduction. χ2 tests were used to compare the rates of premature death (i.e., percent T. ni larvae that died before pupating and without producing parasitoids) of both non-parasitized and parasitized T. ni, and to compare the rate of successful parasitism (i.e., number of T. ni that produced parasitoids compared to these that survived to pupation) between diet types. When an ANOVA test detected a treatment effect, post hoc Tukey’s range tests were performed to identify which treatments differed. When an interaction between the main effects was detected in two-way ANOVA, interaction plots were used to identify the nature 106 Chapter 5 of the interaction. When an interaction between the main effects was detected in ANCOVA, scatterplots and post hoc Pearson product-moment correlations with Bonferroni correction for α = 0.05/6 = 0.0083 were used to identify the nature of the effects and which treatments differed. When a χ2 test was significant, comparisons between pairs of treatments using χ2 with Bonferroni correction for α = 0.05/(((6x6)-6)/2) = 0.0033 were performed to identify which treatments differed. Assumptions of normal distribution and homoscedasticity of samples were verified using residual-by-predicted plots. None of the samples showed strong deviations from these assumptions. All means reported ± SEM. All analyses were conducted in R version 2.14.1 (R Development Core Team, 2011). 5.4. Results The eventual outcomes of the caterpillars used in the different treatments in this experiment are detailed in Table 5.3. In this section, no to moderate protein reduction refers to 43 to 25.5 g/L of protein (diets #1, #2, and #3), whereas pronounced protein reduction refers to 16.75 to 8.1 g/L of protein (diets #4, #5, and #6). 5.4.1. Trichoplusia ni Lower casein content of diet led to significantly longer development time of T. ni larvae (two-way ANOVA, F5, 233 = 462.42, P < 0.001) when the protein reduction was pronounced (Fig. 5.2). There was no significant effect of sex on larval development time (two-way ANOVA, F1, 233 = 3.30, P = 0.07), nor was there an interaction between sex and diet type (two-way ANOVA, F5, 233 = 0.63, P = 0.72). The mass of male pupae was significantly greater than that of female pupae (two- way ANOVA, F1, 232 = 53.53, P < 0.001). Pronounced protein reduction led to lower pupal mass 107 Chapter 5 (two-way ANOVA, F5, 232 = 56.47, P < 0.001) (Fig. 5.3). There was no significant interaction between sex and diet type (two-way ANOVA, F5, 232 = 1.42, P = 0.22). 5.4.2. Trichoplusia ni rates of survival and successful parasitism Rate of premature death of T. ni larvae was significantly different between diet types for both non-parasitized (χ2 = 50.27, df = 5, P < 0.001) and parasitized (χ2 = 15.05, df = 5, P = 0.01) groups (Table 5.3). Pairwise χ2 comparisons found that premature death was only higher on the 8.1 g/L protein diet compared to each of the other diet types in the non- parasitized group, but found no significant difference in the parasitized group after Bonferroni correction, although it was visibly higher on the 8.1 g/L protein diet (Table 5.3). In the parasitized group this rate was artificially reduced because the number of larvae that died prior to the parasitism event was not recorded. There was no difference of successful parasitism rate between the different diet types (χ2 = 5.66, df = 5, P = 0.34) (Table 5.3). 5.4.3. Trichoplusia ni food consumption For non-parasitized caterpillars, decreasing diet quality (two-way ANOVA, F5, 158 = 334.88, P < 0.001), but not sex (two-way ANOVA, F1, 158 = 0.01, P = 0.93), was associated with increased food consumption. There was no significant interaction between diet quality and sex (two-way ANOVA, F5, 158 = 0.45, P = 0.82). Non-parasitized caterpillars developing on the 8.1 g/L protein diet consumed on average 3.63 times more food than these developing on the control diet (food consumption of 1.1710 ± 0.0397 g and 0.3227 ± 0.0090 g respectively). Parasitized caterpillars also consumed more food with decreasing diet quality (One-way ANOVA, F5, 139 = 100.10, P < 0.001); e.g., parasitized caterpillars that developed on the 8.1 g/L protein diet consumed on average 2.21 times more food than those developing on the 43 g/L 108 Chapter 5 protein diet (food consumption of 1.0743 ±0.0360 g and 0.4857 ±0.0134 g respectively) (Fig. 5.4). There were significant main effects on food consumption of both parasitism status (two-way ANOVA, F5, 330 = 28.15, P < 0.001) and diet quality (two-way ANOVA, F5, 330 = 388.18, P < 0.001), as well as an interaction between these two main effects (two-way ANOVA, F10, 330 = 4.31, P < 0.001) (both sexes combined). On diets containing 43 g/L protein (control) to 11.5 g/L protein, parasitized caterpillars ate more food than their non-parasitized counterparts, but only on the diet containing 8.1 g/L protein (the lowest quality diet) did non-parasitized caterpillars eat more than parasitized ones (Fig. 5.4). There was a significant interaction between diet type and C. vanessae brood size on T. ni food consumption (ANCOVA, F5, 127 = 2.46, P = 0.04), as well as a main effect of diet type (ANCOVA, F5, 127 = 44.12, P < 0.001), but no main effect of brood size (ANCOVA, F1, 127 = 1.05, P = 0.31). Scatterplots and Post hoc Pearson-product moment correlation showed a possible correlation between brood size and food consumption only on 8.1 g/L protein diet (r = 0.55, df = 21, P = 0.007). 5.4.4. Cotesia vanessae development time Moderate protein reduction of the host’s diet had no effect on C. vanessae development time within the host, but pronounced reduction significantly delayed development time within the host (one-way ANOVA, F5, 183 = 21.78, P < 0.001) (Fig. 5.5). There was a significant effect of the host’s diet type on the development time of C. vanessae within their cocoons (F5, 186 = 3.76, P = 0.003). However, a post hoc test indicated that the only significant difference was between groups on 43 g/L protein diet and 11.5 g/L protein diet (P = 0.013) (Fig. 5.6). Pronounced protein reduction led to a significant increase of C. vanessae total development time (one-way ANOVA, F5, 183 = 24.91, P < 0.001) (Fig. 5.7). 109 Chapter 5 There was a significant interaction between development time within the host and diet type on development time in cocoons (ANCOVA, F5, 177 = 3.75, P = 0.003). There was also a significant main effect of diet type (ANCOVA, F5, 177 = 1.64, P = 0.002), but not of development within the host (ANCOVA, F1, 177 = 1.64, P = 0.20). Post hoc Pearson-product moment correlation found a significant negative correlation between development time within the host and within the cocoon for the two highest quality diets (43 g/L protein diet : r = -0.71, n = 28, P < 0.001; 34.25 g/L protein diet : r = -0.53, n = 32, P = 0.002), but no correlations in the other diet types (|r| ≤ 0.14, n = 26~38, P > 0.05) (Fig. 5.8). Thus, on the 43 g/L protein and 34.25 g/L protein diets, the parasitoid broods that developed faster within their host subsequently developed slower within their cocoons, and vice versa. 5.4.5. Cotesia vanessae numbers and mass Hosts developing on lower-quality diets produced significantly fewer parasitoids per brood (one-way ANOVA, F = 15.31, P < 0.001) (Fig. 5.9), and broods of lower masses (one- way ANOVA, F5, 183 = 21.26, P < 0.001). However, there was no effect of diet type on the mass of individual parasitoids (ANOVA, F5, 189 = 1.30, P = 0.27) (Fig. 5.10). The size of broods reflects both the number of eggs laid in hosts and the survival rate of parasitoids. ANCOVAs showed significant main effects of brood size (F5, 183 = 152.82, P < 0.001) and brood mass (F5, 183 = 152.82, P < 0.001) on parasitoid individual mass. There was no interaction between diet type and brood size (F5, 183 = 1.17, P = 0.33) and between diet type and brood mass (F5, 183 = 0.27, P = 0.93). This indicates that larger and heavier broods correspond with lower mass of individual parasitoids (Fig. 5.11). There were significant main effects of diet type (ANCOVA, F5, 183 = 165.89, P < 0.001) and number of parasitoids per brood (ANCOVA, F1, 183 = 1357.19, P < 0.001) on the mass of broods. There was also a significant interaction between diet type and number of parasitoids per brood (ANCOVA, F5, 183 = 5.50, P < 0.001). Post hoc Pearson product-moment correlation 110 Chapter 5 found significant positive correlations between brood size and brood mass on each diet type (r ≥ 9.1, n = 27 to 38, P < 0.001). A scatterplot indicated that, even if diets of reduced protein contents were associated with reduced brood size and mass, the correlation between brood size and mass was similar across diet types (Fig. 5.12). There were significant main effects of diet type (ANCOVA, F5, 175 = 28.25, P < 0.001) and brood size (ANCOVA, F1, 175 = 3.92, P = 0.049) on C. vanessae development time. There was also a significant interaction between diet type and total development time (ANCOVA, F5, 175 = 5.69, P < 0.001). Scatterplots and post hoc Pearson product-moment correlation showed a correlation between brood size and development time only on 25.5 g/L protein diet (r = - 0.71, df = 26, P < 0.001), mainly caused by the inexplicably long development time of five small broods (1 to 33 parasitoids). 5.4.6. Cotesia vanessae F2 development, numbers, and mass There was no effect of diet (one-way ANOVAs, P > 0.05) on the measured fitness parameters for the F2 progeny of parasitoids emerging from hosts reared on control versus - 99.7% casein diet; i.e., developmental time within the host, developmental time within cocoons, number of emergent wasps per host, mean dry individual adult mass per host, and the estimated total dry adult parasitoid mass per host (Fig. 5.13). 5.5. Discussion Analyses did not detect a decrease of fitness for either T. ni and C. vanessae for diets containing at least 25.5 g/L of protein, but did detect a decrease of fitness for both T. ni and C. vanessae associated with a decrease of protein on diets containing less than 25.5 g/L of protein. This indicates that the decrease of fitness of C. vanessae is linked with that of its host. It also indicates that T. ni can develop on a wide range of protein concentrations in its diet, a tolerance that has also been observed in other polyphagous Noctuidae (Bloem et al., 111 Chapter 5 1990; Karowe et al., 1989). Across all treatments, the number of premature deaths was consistently higher in parasitized groups (Table 5.3) (although not significant). Some of the parasitized T. ni that died prematurely appeared constipated, as did caterpillars observed in Chapter 3, an effect attributed to parasitism disabling the host’s digestive system. T. ni larvae appeared to exhibit a compensatory feeding response when reared on diets of reduced quality, which likely contributes to their ability to develop on diets of different quality. On diets containing 43, 34.25 and 25.5 g/L of protein, non-parasitized larvae had similar development time and pupal mass, but their food consumption increased for each reduction of protein. However, on diets containing 16.75, 11.5, and 8.1 g/L protein, although food consumption increased, larval development time increased and pupal mass decreased. Similarly, parasitized T. ni ate more food than non-parasitized ones, likely to compensate for nutrients consumed by parasitoids. This effect has been observed with other koinobiont endoparasitoid species (Shi et al., 2002), although the opposite effect has also been documented (Doetzer et al., 1998; Elzinga et al., 2003). Another physiological effect detected with C. vanessae was the trade-off of time spent in the host and in the cocoon when hosts were reared on the two diets having the highest protein concentration; i.e., 43 and 34.25 g/L. This effect might be because larvae develop until they egress in synchrony from the host upon perception of probable egression cues. Thus, the longer larvae spend in the host, the more mature they are, and the less time they need to spend in the cocoon. This effect might not represent actual “egg-larva” development time (from laying of egg to pupation of larva) and “pupa” development time and might simply reflect the time spent in the cocoon as prepupa before pupating. Indeed, a few C. vanessae that didn’t form a cocoon following their emergence from hosts allowed for observation that they displayed various length of time between their egression and their pupation. For these, pupation occurred between one to at least three days post egression. Thus, there might not be a trade-off between actual egg-larval and pupal development times. 112 Chapter 5 This effect was not detected on lower-quality diets. This was perhaps because of the general increase of C. vanessae development time in the host (Fig. 5.5) when hosts were fed low- protein diets. Previous authors noticed that the egg-larval development time of both solitary and gregarious parasitoids was associated with the stage of the host being parasitized, with injection of eggs into earlier stages inducing longer parasitoid egg-larvae development time (Allen et al., 1991; Avilla et al., 1987; Bell et al., 2003; Elzinga et al., 2003; Nakamura et al., 2001; Obrycki et al., 1985; Shi et al., 2002). For the solitary endoparasitoid Encarsia formosa Gahan (Hymenoptera: Aphelinidae), this effect is attributed to the delayed development of the 1st instar parasitoid, which molts to 2nd instar only when the host reaches its 4th instar (Nechols et al., 1977). The closely related Encarsia tricolor Förster (Hymenoptera: Aphelinidae) displayed a similar increase in egg-larval development time when earlier stages of the host were parasitized, but in this case the effect is mainly due to a delay in egg hatching (Avilla et al., 1987). In the latter study, only when the pharate adult host is parasitized does the pupation time of the parasitoid increase. Otherwise, none of these studies found a correlation between parasitoid egg-larval and pupal or cocoon stages development time, as did the present study. In the current study, it seems unlikely that host maturity had any effect on parasitoid development time in the host as all the hosts were parasitized at the early 4th instar. Parasitoid individual mass significantly decreased with increasing brood size and mass. This effect has also been found in previous studies (Fidgen et al., 2000; Reitz et al., 1995; Takagi, 1986; Waage, 1986; Waage et al., 1984). This effect can be explained by the finite resources provided by a host, which translates into scramble competition; i.e., finite resources are split between all individuals present, thus a higher number of parasitoids results in smaller individuals. However, on all diets, larger broods were also heavier. This contradicts the hypothesis that in an environment of scramble competition all the usable resources are 113 Chapter 5 consumed and transformed into parasitoid mass, thus brood mass should not correspond to brood size. A possible explanation could be that larger broods have a longer development time or induced their host to feed more to allow the host to produce more resources (Elzinga et al., 2003). This does not appear to apply here because brood size was neither correlated with parasitoid development time nor host food consumption. Another possible explanation could be that with increasing brood size, parasitoids increasingly feed on less desirable host tissues. But if this was the case then why not always feed on these tissues if it creates a gain of mass with no cost to development time? Perhaps the cost of such behaviour, if it exists, could be due to a fitness variable that was not measure; e.g., longevity and fertility. Perhaps part of the answer can be found in Harvey et al. (2000), who argue that while tissue-feeding parasitoids favor maximization of body mass and usually display a trade-off between development time and body mass, haemolymph-feeding Microgastrinae show more complicated interactions with their host and favour fast development time over large body size. Haemolymph feeders do not consume all the available resources and their development time may not correlate with the amount of available resources (i.e., host size). Because C. vanessae larvae are gregarious and exhibit synchronized egression from the host, it could be that the intense pressure to develop quickly reduces the ability of parasitoid larvae to exploit all the resources from their host, perhaps by lack of time to explore all areas of the haemocoel, thus leaving behind haemolymph and fat in the unoccupied areas. Indeed, the concentration of parasitoids was observed to always be higher in the posterior part of the host, and with small broods there are often few individuals emerging from the thorax area of the host. Thus, I predict, although did not test, that body mass of individuals that developed in the posterior part should be lower than these that developed in the anterior part. This situation would explain why even small broods that do not consume all available resources display scramble competition and why brood mass increases with brood size. 114 Chapter 5 If resources are truly limiting in a host, there should be a point at which brood size becomes so large that parasitoids exploit all the resources available and increase of brood size do not correspond to an increase of brood mass anymore. Fig. 5.12 shows that the largest parasitoid brood (from a host that developed on the 43 g/L protein diet) is not the heaviest one. This indicates a possible threshold (approximately 180 parasitoids per brood) at which an increase in brood size does not result in an increase of brood mass, and perhaps even a decrease of brood mass. Another parasitoid brood that contained over 180 parasitoids seemed to confirm this pattern (brood mass of 23.1 g for 201 parasitoids), but it was excluded from the analysis because the host was parasitized twice. Although at the individual level, parasitoids from small broods have the ability to grow bigger and gain a fitness advantage, the greater number of smaller-size parasitoids produced by larger broods may not necessarily result in a disadvantage from the parents’ perspective. The variation of parents’ realized fitness due to the number or the size of offspring is dependent on ecological conditions; e.g., number of hosts available, competition with other parasitoids, availability in space and time of hosts (Vos et al., 2003). Thus, the ability of C. vanessae to produce a large diversity of brood sizes (Fig. 5.9) with individuals of various masses (Fig. 5.10) and its ability to maintain individual masses on hosts feeding on diets of different qualities (Fig. 5.10) would likely contribute to its success in different environments. There was no inter-generational effect of poor-quality diet on the fitness of F2 parasitoids. Studies on other organisms from various taxa found that parents that developed in more favorable environments generally produced higher quality offspring (Bonduriansky et al., 2007; Einum et al., 1999; Gould, 1988; Hafer et al., 2011; Marshall et al., 2006; Wellington, 1976, 1977), although not always (Myers et al., 2011). However, differences in offspring quality might only be noticeable if offspring develop in low-quality environments, as compensatory mechanisms can help low-quality offspring to quickly improve from their 115 Chapter 5 initial poor condition in high-quality environments (Donelson et al., 2009; Rombough, 1994). In the present study, F2 C. vanessae developed in hosts being reared on high-quality diet (43 g/L protein), which is a favorable environment. This could be why offspring quality from high- and low-condition parents was similar. The similar overall effects of host food quality on the development of T. ni and C. vanessae indicate that the fitness of the parasitoid is linked with the quality of the diet of its host’s food. The fitness of T. ni and of C. vanessae follow similar patterns. This could indicate that parasitoid fitness is closely associated with host fitness. In this case, the association between host food quality and parasitoid fitness might only exist because of the associations between host food quality and host quality, and between host quality and parasitoid fitness. Because host mass decreased with decreasing food quality, and because parasitoids fully rely upon the availability of host tissues to develop, it may be that host mass is the only true factor influencing parasitoid fitness. In this study, the mass of parasitized hosts was not measured, but because host mass is closely associated with diet quality, we cannot link parasitoid fitness to diet quality independently of host mass. To investigate if parasitoid fitness is an indirect effect of host mass (or size), a future study comparing the fitness of parasitoids produced by hosts of similar mass and size reared on different quality diets would be necessary. 116 Chapter 5 Table 5.1. McMorran diet manipulations. Total protein amount was estimated as to the amount of casein plus 8 g/L of protein from wheat germ. Diet # Total protein Casein Casein % removed 4 M KOH Cellulose amount (g/L) amount (g/L) compared to amount amount (g/L) original recipe (ml/L) 1 43 35 0% (Control) 5 0.5 2 34.25 26.25 25% 3.75 9.25 3 25.5 17.5 50% 2.5 18 4 16.75 8.75 75% 1.25 26.75 5 11.5 3.5 90% 0.5 32 6 8.1 0.1 99.7% 0.15 35.4 117 Chapter 5 Table 5.2. Summary of the different treatments at the start of the experiment, not accounting for replication later discarded due to errors and supplementary replications added to the 8.1 g/L protein diet to compensate for the high rate of premature death of caterpillars developing on this diet. Diet # Protein amount n non-parasitized n parasitized (g/L) caterpillars1 caterpillars 1 43 45 (16♀, 29♂) 45 2 34.25 45 (22♀, 21♂) 45 3 25.5 39 (23♀,13♂) 42 4 16.75 45 (18♀, 18♂) 45 5 11.5 45 (20♀, 23♂) 45 6 8.1 45 (16♀, 19♂) 45 1Individuals that died before pupating were not sexed 118 Chapter 5 Table 5.3. Outcome of Trichoplusia ni individuals at the end of the experiment. Protein Premature Produced Pupated, Turned amount (g/L) death parasitoids pupa died to moth Group A: 43 (n = 45) 0 NA 0 45 non- 34.25 (n = 43) 0 NA 0 43 parasitized 25.5 (n = 39) 2 NA 1 36 16.75 (n = 45) 2 NA 1 42 11.5 (n = 45) 0 NA 1 (+1)1 43 8.1 (n = 50) 15 NA 1 34 Group B: 43 (n = 37) 1 31 2 3 parasitized 34.25 (n = 45) 3 35 0 7 25.5 (n = 42) 3 29 4 6 16.75 (n = 45) 2 38 1 4 11.5 (n = 43) 2 37 1 3 8.1 (n = 45) 10 27 0 8 1Pupa accidentally killed during manipulation 119 Chapter 5 ♀ rough depressions genital pore anus cremaster segment 9 & 10 V shaped toward the 0.5 mm genital pore ♂ pair of divided tubercles on either side of the genital pore pair of rounded pads cremaster anus 0.5 mm Figure 5.1. Trichoplusia ni pupae, female (top) and male (bottom), posterior end, ventral side. 120 Chapter 5 Figure 5.2. Trichoplusia ni larvae (males and females combined) development time on diets containing different amounts of protein. The boxes are delimited by the 25th and 75th percentiles. The upper and lower whiskers indicate Q3 + 1.5 IQR and Q1 – 1.5 IQR, respectively. Medians and means are represented by solid lines and dotted lines, respectively. Significance (Tukey’s HSD, α = 0.05, following one-way ANOVA) is indicated by a letter above each box (i.e., boxes with different letters represent significantly different means). 121 Chapter 5 Females Males Figure 5.3. Trichoplusia ni mass of pupae from larvae that developed on diets containing different amounts of protein. The boxes are delimited by the 25th and 75th percentiles. The upper and lower whiskers indicate Q3 + 1.5 IQR and Q1 – 1.5 IQR, respectively. The median and mean are represented by a solid line and a dotted line respectively. Significance (Tukey’s HSD, α = 0.05, following one-way ANOVAs analyzing males and females separately) is indicated by a letter above each box (i.e., boxes with different letters represent significantly different means). Boxplots were analyzed separately. 122 Chapter 5 Non-parasitized Parasitized Figure 5.4. Estimated dry mass of food consumed by non-parasitized (top) and parasitized (bottom) T. ni larvae developing on diets containing 43 to 8.1 g/L of protein. The boxes are delimited by the 25th and 75th percentiles. The upper and lower whiskers indicate Q3 + 1.5 IQR and Q1 – 1.5 IQR, respectively. The median and mean are represented by a solid line and a dotted line respectively. Significance (Tukey’s HSD, α = 0.05, following one-way ANOVAs analyzing separately non-parasitized and parasitized T. ni) is indicated by a letter above each box (i.e., boxes with different letters represent significantly different means). Boxplots were analyzed separately. 123 Chapter 5 Figure 5.5. Cotesia vanessae development time within T. ni larvae that developed on diets containing different amounts of protein. The boxes are delimited by the 25th and 75th percentiles. The upper and lower whiskers indicate Q3 + 1.5 IQR and Q1 – 1.5 IQR, respectively. The median and mean are represented by a solid line and a dotted line respectively. Significance (Tukey’s HSD, α = 0.05, following one-way ANOVA) is indicated by a letter above each box (i.e., boxes with different letters represent significantly different means). 124 Chapter 5 Figure 5.6. Cotesia vanessae development time within their cocoons from T. ni hosts that developed on diets containing different amounts of protein. The boxes are delimited by the 25th and 75th percentiles. The upper and lower whiskers indicate Q3 + 1.5 IQR and Q1 – 1.5 IQR, respectively. The median and mean are represented by a solid line and a dotted line respectively. Significance (Tukey’s HSD, α = 0.05, following one-way ANOVA) is indicated by a letter above each box (i.e., boxes with different letters represent significantly different means). 125 Chapter 5 Figure 5.7. Cotesia vanessae total development time in T. ni hosts that developed on diets containing different amounts of protein. The boxes are delimited by the 25th and 75th percentiles. The upper and lower whiskers indicate Q3 + 1.5 IQR and Q1 – 1.5 IQR, respectively. The median and mean are represented by a solid line and a dotted line respectively. Significance (Tukey’s HSD, α = 0.05, following one-way ANOVA) is indicated by a letter above each box (i.e., boxes with different letters represent significantly different means). 126 Chapter 5 Figure 5.8. Scatterplot with trend lines showing Cotesia vanessae development time inside their host in relation to their development time inside cocoons, on each diet type; i.e., 43 g/L (black, slope = -0.67; r = -0.71), 34.25 g/L (blue, slope = -0.43; r = -0.53), 25.5 g/L (green, slope = 0.05, r = 0.07), 16.75 g/L (yellow, slope = 0.04, r = 0.05), 11.5 g/L (orange, slope = -0.06, r = - 0.14), 8.1 g/L (red, slope = 0.04, r = 0.14). 127 Chapter 5 Figure 5.9. Cotesia vanessae brood size per type of diet from T. ni hosts that developed on diets containing different amounts of protein. The boxes are delimited by the 25th and 75th percentiles. The upper and lower whiskers indicate Q3 + 1.5 IQR and Q1 – 1.5 IQR, respectively. The median and mean are represented by a solid line and a dotted line respectively. Significance (Tukey’s HSD, α = 0.05, following one-way ANOVA) is indicated by a letter above each box (i.e., boxes with different letters represent significantly different means). 128 Chapter 5 Figure 5.10. Average dry mass of individual adult Cotesia vanessae per brood from T. ni hosts that developed on diets containing different amounts of protein. The boxes are delimited by the 25th and 75th percentiles. The upper and lower whiskers indicate Q3 + 1.5 IQR and Q1 – 1.5 IQR, respectively. The median and mean are represented by a solid line and a dotted line respectively. Significance (Tukey’s HSD, α = 0.05, following one-way ANOVA) is indicated by a letter above each box. No significant difference was found between any treatment. 129 Chapter 5 Figure 5.11. Scatterplot with trend lines showing C. vanessae brood size in relation to average dry mass of individual parasitoids emerged from T. ni fed diets containing different amounts of protein; i.e., 43 g/L (black, slope = -0.005, r = -0.87), 34.25 g/L (blue, slope = - 0.0013, r = -0.62), 25.5 g/L (green, slope = -0.0005, r = -0.64), 16.75 g/L (yellow, slope = - 0.0007, r = -0.67), 11.5 g/L (orange slope = -0.0007, r = -0.59), 8.1 g/L (red, slope = -0.0011, r = -0.66). 130 Chapter 5 Figure 5.12. Scatterplot with trend lines showing number of C. vanessae adult per brood in relation to estimated total wasp mass of these broods, from T. ni fed diets containing different amounts of protein; i.e., 43 g/L (black), 34.25 g/L (blue), 25.5 g/L (green), 16.75 g/L (yellow), 11.5 g/L (orange), 8.1 g/L (red). All slopes between 0.10 and 0.16, and r ≥ 9.1. 131 Chapter 5 A C B D Figure 5.13. F2 C. vanessae: A. Development time within host, B. Development time within cocoon, C. Brood size, D. Adults average dry mass per brood; from T. ni hosts that developed on diets containing 43 and 8.1 g/L of protein. The boxes are delimited by the 25th and 75th percentiles. The upper and lower whiskers indicate Q3 + 1.5 IQR and Q1 – 1.5 IQR, respectively. The median and mean are represented by a solid line and a dotted line respectively. Significance (Tukey’s HSD, α = 0.05, following one-way ANOVA) is indicated by a letter above each box (i.e., boxes with different letters represent significantly different means). Boxplots were analyzed separately. 132 Within cocoons Within hosts Adults average dry mass (mg) Chapter 6 CHAPTER 6: GENERAL DISCUSSION “The scientist has a lot of experience with ignorance and doubt and uncertainty, and this experience is of very great importance, I think. When a scientist doesn’t know the answer to a problem, he is ignorant. When he has a hunch as to what the result is, he is uncertain. And when he is pretty darn sure of what the result is going to be, he is still in some doubt. We have found it of paramount importance that in order to progress we must recognize our ignorance and leave room for doubt. Scientific knowledge is a body of statements of varying degrees of certainty–some most unsure, some nearly sure, but none absolutely certain.” Richard P. Feynman (1988). The Value of Science. Results of this study show that thelytokous Cotesia vanessae occur in Canada and have a broad fundamental host range within the family Noctuidae. Within this family, species in the subfamily Plusiinae (the “loopers”) appear to be the best hosts. Cotesia vanessae can also develop on species of Nymphalini, as previously reported (Stefanescu et al., 2012). In no-choice tests, C. vanessae attempted to parasitize larvae of nearly all Lepidoptera species it was exposed to. Thus it may be able to develop on species in other lepidoperan families, as suggested by data in Yu (2016). The current study shows that parasitoid fitness is affected by host species, and that the fitness of the parasitoid and that of its host are affected by the quality of the host’s diet in term of protein content. Trichoplusia ni hosts were able to complete development on diet of various nutritional qualities, although development time and individual mass was affected. Diets that affected the host also affected the parasitoid, in terms of development time, brood size and brood mass, but not individual parasitoid mass. The next generation of parasitoids appeared unaffected by host’s diet quality. During this study, lepidopteran individuals of some species emerged with deformed wings (not caused by problems during emergence from the pupa), including individuals never exposed to parasitoids. Based on previous research (Morris, 1967), maladapted diet was considered the likely cause. Results in the Chapters 3 and 5 indicate that the McMorran diet is likely not optimal for all the species reared. This initially provided grounds for investigating parasitoid fitness on hosts fed diets of different qualities (Chapter 5 study), but misshapen 133 Chapter 6 wings were observed in both good and poor host species (see Chapter 4). This suggests no strong link with the cause of these deformities and parasitoid fitness. Cotesia vanessae was found to display scramble competition in certain species (brood size inversely correlated with individual mass), and contest competition in all species dissected (siblicidal first instar). Contest competition through larval siblicide has not previously been observed in gregarious Braconidae species. The parasitoid population studied was gregarious and parthenogenetic, hence, offspring in a brood must be genetically very close to one another. We can only speculate as whether or not larval siblicide also occurs in male-female populations of C. vanessae. This siblicidal behaviour seems to contradict previous hypotheses that gregariousness and close relatedness among offspring should be associated with altruistic behaviour (Hamilton, 1963; Segoli et al., 2009; Giron et al. 2004; Godfray, 1994; Pexton et al., 2002; Rosenheim, 1993). This could either indicate that gregariousness in C. vanessae is still evolving and altruism among brood siblings might eventually arise, or that siblicidal behaviour is a desirable trait (Tuda et al., 1998). This behaviour appears to be an inefficient strategy because of the costs for the mother of producing extra eggs, the food wasted on larvae that are killed (which do not appear to be consumed by their siblings), and the time spent by larvae searching for siblings and killing them rather than feeding. However, adult females devote little resources for individual egg production because eggs are hydropic, i.e., small and apparently yolkless. Devoting fewer resources for individual eggs allows for the production of a larger number of eggs in the ovaries (Strand, 2000). Laying of extra eggs into hosts may increase the overall fitness of this species by increasing its intra- and interspecific competitiveness in super- and multiparasitized hosts, and in maximizing the number of offspring produced by hosts of wide range of sizes. In addition, the loss of the caudal appendage of C. vanessae larvae should be associated with a reduced motility of individuals. This means that contest competition likely 134 Chapter 6 only occurs among larvae in close proximity within the host’s hemocoel, and is likely a key anatomical feature that permits gregariousness in this species. High host specificity is a desirable trait when selecting species to use in biological control programs due to reduced risk of parasitism on non-target species (Brodeur, 2012; Hervet et al., 2016a). For example, the use of the non-host specific Cotesia glomerata (L.) in a classical biological control effort for reducing populations of the cabbage white, Pieris rapae (L.), in North America resulted in the extirpation from part of its range of the native non-pest mustard white butterfly, Pieris oleracea Harris (Benson et al., 2003). On the other hand, the wide host-range of Cotesia marginiventris (Cresson) within the family Noctuidae did not impede investigation as a candidate for the biological control of these pests in greenhouse, which provided efficient control of Trichoplusia ni (Gillespie et al., 1999; Messelink, 2002). Unfortunately, the high costs associated with the production of C. marginiventris led to the discontinuity of this biological control program. Like C. marginiventris, C. vanessae could also be considered for the biological control of T. ni and other noctuid pests. Advantageous traits of C. vanessae life history may result in it being more economical to produce than C. marginiventris: 1) Adult Cotesia vanessae in our laboratory commonly lived up to one month, and on some occasions up to a month and half (as per the rearing conditions described in Chapters 3 and 5). This reduces the need to frequently regenerate the laboratory colony and reduces the risk of accidentally losing the colony. In contrast, the longevity of adult female C. marginiventris is about 10 to 14 days (Boling et al., 1970; Gillespie et al., 1997a); 2) Cotesia vanessae is very fertile throughout its adult life. It is able to parasitize a large number of hosts in a short amount of time; e.g., one individual was observed to parasitize 50 Euxoa ochrogaster (Guenée) larvae in 20 minutes; although these larvae were not further observed to determine parasitism success and size of broods. It is a gregarious species that produces relatively large broods per host; i.e., in Chapter 5, parasitized T. ni reared on control diet produced an average 92 ± 6.16 (SE) parasitoids (range: 16 to 199) (n = 31). Cotesia 135 Chapter 6 marginiventris being solitary, each parasitized larva produces a single parasitoid, thus requiring the rearing of many more hosts to obtain a similar number of parasitoids; 3) Thelytokous C. vanessae do not need to find mates, and each individual can lay eggs; whereas C. marginiventris populations are bisexual with a male-bias (Gillespie et al., 1997b; Jalali et al., 1987); 4) Cotesia vanessae usually parasitize the host upon first encounter and is fertile one day after emergence from the cocoon. Cotesia marginiventris starts to lay eggs two days after emergence (Gillespie et al., 1997a). Some unfavorable aspects are that 1) C. vanessae likely requires more time to develop (about 17 days to develop in the host and an additional 10 days in the cocoon, at 20 °C) than does C. marginiventris (about 7 to 9 days in the host and 6 to 8 days in the cocoon, at 25 °C); 2) Cotesia vanessae does not prevent maturation of its hosts prior to killing it, like C. marginiventris, thus parasitism does not directly prevent plant damage; 3) parasitized hosts consume more food than non-parasitized individuals. This means that its use in biological control programs can only be used to control pest populations over more than one generation. To elucidate the ecological host range of C. vanessae, it would be essential to investigate its association with plant species and conduct tests in more natural conditions. 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However, once after having handled Trichoplusia ni larvae, I introduced my hand into a cage containing a few hundred Cotesia vanessae, one of which landed on the dorsal side of my right index finger and immediately inserted its ovipositor into the skin. While doing so, it positioned its wings, mid and hind legs upward as displayed in Figure 3.3 (Chapter 3), as if parasitizing a host. The event lasted about 10 seconds and was followed by the appearance of a small (~1 mm diameter) slightly itchy red swell, which disappeared an hour later. It has been shown that parasitoids can be fooled by artificial hosts, given that they have the right shape, odor, and texture (Dindo et al., 2002; Grenier, 2012; House, 1978). As displayed in Chapter 3, C. vanessae is opportunistic, thus could have been misled by a host’s smell on a finger. It might also have been close to the end of its life, as such individuals become more likely to engage unlikely hosts (personal observation). Because I have handled tens of thousands of C. vanessae the chance to observe unusual behavior was increased and this is certainly not typical of the species. A.2. Ovipositor attachment Sometimes parasitoids were observed to struggle to, or even to be unable to, remove their ovipositor from a host. Hosts were sometimes observed to carry dead parasitoids attached by their ovipositor. Attempting to remove stuck individuals with tweezers sometimes resulted in the ripping of their abdomen, leaving behind the ovipositor and abdominal organs attached. Cotesia vanessae was observed to possess a barbed spear alongside its ovipositor. A feature shared with other Cotesia species (Thomas, 2012), which likely helps them remain firmly attached to their host during oviposition. A.3. Climbing prior to parasitoid egression Parasitized hosts usually climb shortly prior to parasitoid egression. This behavior was also observed with Autographa californica in canola fields parasitized with another Cotesia species (GenBank accession number: KF640233), and with Aglais milberti parasitized with yet another Cotesia species (unidentified), which egressed from hosts at the top of plants. What causes this behaviour is unknown, but one possible benefit to parasitoids is to ensure that they do not find themselves buried when egressing from a hiding host. A.4. Parasitoid larvae egression Cotesia vanessae larvae egress simultaneously from both of its host’s sides (as displayed by exit holes in Fig. A.1), although the larvae located toward the posterior of the host always egress slightly prior to anterior ones, and rarely from the dorsal side. In the experiments described in Chapter 3 and Chapter 5, about 10 host larvae were dissected after egression. Dead parasitoid larvae that failed to egress were always seen inside of them. Hosts (Trichoplusia ni) from the experiment described in Chapter 5 only contained a few parasitoid larvae (<10), all of which appeared to have reached their full size (Fig. A.2), but 158 hosts (Euxoa messoria) from the Chapter 3 experiment contained various numbers of dead parasitoid larvae. One E. messoria that died of premature death contained over 300 C. vanessae larvae of different stages (from nearly visible to the naked eye to virtually fully mature). In the Chapter 3 experiment the mother wasp remained 48 hours with its host, thus it was able to parasitize it multiple times, whereas in the Chapter 5 experiment each host was parasitized only once, thus reducing parasitoid crowdedness within the host. This should explain the different numbers of dead parasitoid larvae remaining within hosts. However, the total numbers of emerged parasitoid from T. ni host were similar in both experiments (Chapter 3 and 5). Immediately after leaving their host, each parasitoid larvae form individual cocoons. Each cocoon is attached to the surrounding siblings’ cocoons by silk, thus forming a cocoon mass. In small cocoon masses, individual cocoons are usually attached to the substrate from their sides, whereas in big cocoon masses (as in Fig. A.3) the cocoons are usually attached to the substrate by their bottom (where the rear end of the parasitoid is located). Hosts usually crawl away after the last parasitoid has egressed from their body, thus allowing parasitoid larvae that egressed from both sides of the host to reunite and form a single cocoon mass. Sometimes the host remained where the parasitoids emerged from its body and died there, and in other rare occasions the hosts traveled during parasitoid larvae egression. Both events usually resulted in parasitoids forming two or more cocoon masses. To avoiding misunderstanding, in the experiments described in Chapters 3 and 5 a “cocoon mass” refers to all the cocoons from a brood of parasitoids that egressed from one caterpillar. An extra ~2 mm of fluffy silk is produced to surrounds the cocoon mass, likely a protection against hyperparasitoids and unfavorable weather. It was observed that parasitoids larvae that happened to be far from other larvae (≥ 2 mm) often did not manage to fully form a cocoon and eventually dried out and died, either as larvae or pupae. Within cocoon masses the cocoons were always relatively well-aligned and staggered (Figs. A.3 & A.4), in a way that is reminiscent of the structure of nests of colonial wasps (Vespinae and Polistinae) and bees (Apis spp.). Parasitoid larvae can probably sense the position of their surrounding siblings and place themselves precisely between them. Even individuals that reunite themselves after having egressed from either side of a host end up well staggered. This organization may provide each adult parasitoid with a fair chance to successfully emerge from the cocoon mass considering that the anterior end of the cocoon is not obstructed by other cocoons, which may not be the case for the Cotesia species that do not appear to have any organization in their cocoon mass (personal observations with a Cotesia sp. (GenBank accession number: KF640232) egressed from Autographa californica larvae collected from alfalfa fields in southern Alberta in 2013 and 2014). A.5. Larvae color according to host taxa Surprisingly, the C. vanessae larvae that egressed from Noctuidae hosts were always grey, while those that egressed from Nymphalidae were always bright yellow (observed from Aglais milberti and Vanessa cardui hosts. The latter observation stems from a communication with Mark R. Shaw, National Museums of Scotland). This bright yellow color was also observed in larvae of Cotesia atalantae (Packard) reared from A. milberti collected in southern Alberta in 2016. These Nymphalidae species were reared on stinging nettle, Urtica dioica. In contrast, Noctuidae hosts were reared on McMorran diet, or on Bock choy, Brassica rapa, broccoli, Brassica oleracea, and canola, Brassica napus, between experiments, but never on nettle. Either the host identity or its food source could have induced this color difference. 159 3 mm Figure A.1. Last instar Trichoplusia ni larva that died following egression of Cotesia vanessae from its body. The dark circles on the side of the caterpillar are the exit holes made by parasitoid larvae while egressing. Near the centre, a parasitoid larva that died without completing egression is visible. 160 3 mm Figure A.2. Same T. ni larva as in Fig. A.1 dissected dorsally. Four fully-developed dead parasitoid larvae that failed to egress from the host are visible. The two red bodies surrounded by yellow tissue are the gonads of the host. The thick longitudinal white mass is fat surrounding the digestive system of the caterpillar. 161 3 mm Figure A.3. Top view of a Cotesia vanessae cocoon mass, with surrounding silk and open opercula of cocoons removed (some opercula are displayed below the cocoon mass). The few unopened cocoons contain dead parasitoids. 162 2 mm Figure A.4. Cotesia vanessae cocoon mass lateral side, with surrounding silk and open opercula of cocoons removed. A dead parasitoid adult that opened its cocoon but failed to come out of it is visible in the middle of the first row. 163