Coleoptera: Curculionidae) in the Landscape and Horticulture Industry

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DISTRIBUTION, BIOLOGY, AND MANAGEMENT OF Myllocerus undecimpustulatus undatus MARSHALL (COLEOPTERA: CURCULIONIDAE) IN THE LANDSCAPE AND HORTICULTURE INDUSTRY

ANITA SEEN NEAL

A DISSERTATION PRESENTED TO THE GRADUATE SCHOOL OF THE UNIVERSITY OF FLORIDA IN PARTIAL FULFILLMENT OF THE REQUIREMENTS FOR THE DEGREE OF DOCTOR OF PHILOSOPHY

UNIVERSITY OF FLORIDA

2018

To my family, friends, and fellow faculty, without their love and support, none of this would have happened

ACKNOWLEDGMENTS

I would like to express my sincere gratitude to my advisor Dr. Ronald Cave whose patience, support, and motivation provided guidance and advice in my research, in the writing of my dissertation, and my growth as a scientist.

In addition to my advisor, I would like to thank the rest of my committee, Dr.

Jennifer Gillett-Kaufman, Dr. Catharine Mannion, and Dr. Sandra Wilson, for their suggestions, comments, and support that made substantial improvements to my study and created a new-found enthusiasm for the study of insects. I sincerely thank Dr.

Pasco Avery, who provided me access to his laboratory, taught me how to use various instruments, and guided my research with new ideas. I thank Dr. Rodrigo Diaz for conveying the use of a species distribution model.

I am grateful to Michael Thomas, Florida Department of Agriculture and

Consumer Services, Division of Plant Industry and field inspectors for sharing their data collection locations and host plant information for the weevils. Special thanks are extended to Norman Platts, homeowner, for allowing the collection of weevils and Janet

Dawson at the Biological Control Research and Containment Laboratory for assistance with weevil colony maintenance.

I thank the faculty and personnel at the Biological Control Research and

Containment Laboratory and the Indian River Research and Education Center for all their help when doing my research. It has been a pleasure to get to know them all.

I would be amiss not to thank UF/IFAS administration for allowing me time to work on my Ph.D. Lastly, I thank my family, especially my mom, my husband and daughters, and my brothers for their loving support, understanding, and encouragement during this process. 4

TABLE OF CONTENTS

page

ACKNOWLEDGMENTS ...... 4

LIST OF TABLES ...... 7

LIST OF FIGURES ...... 8

ABSTRACT ...... 111

CHAPTER

1 INTRODUCTION ...... 13

2 LITERATURE REVIEW ...... 166

Distribution and Characteristics of Myllocerus Species ...... 166 Description and Life Cycle ...... 177 Host Plants and Insect Damage ...... 188 Management Strategies...... 1919

3 BIOLOGY AND REARING OF Myllocerus undecimpustulatus undatus MARSHALL (COLEOPTERA: CURCULIONIDAE) ...... 311

Materials and Methods ...... 322 Weevil Collection and Colony...... 322 Rearing System...... 322 Results ...... 344 Discussion ...... 355

4 ADULT COLD TOLERANCE AND POTENTIAL NORTH AMERICAN DISTRIBUTION OF Myllocerus undecimpustulatus undatus MARSHALL (COLEOPTERA: CURCULIONIDAE) ...... 51

Materials and Methods ...... 53 Cold Tolerance...... 53 Sustained Cold Exposure (SCE) Versus Repeated Cold Exposure (RCE)...... 55 Niche Modeling...... 56 Results ...... 57 Discussion ...... 59

5 MORTALITY AND FEEDING BEHAVIOR OF ADULT Myllocerus undecimpustulatus undatus MARSHALL (COLEOPTERA: CURCULIONIDAE) EXPOSED TO BIOPESTICIDES IN LABORATORY ASSAYS ...... 70

Materials and Methods ...... 71

Petri Dish Plant-based Bioassay...... 71 Plant and Insect Preparation...... 72 Fungal Formulations...... 72 Biochemical Formulations...... 73 Treatment Applications...... 74 Statistical Analysis...... 76 Results ...... 76 Discussion ...... 78

6 COMPARISON OF MORTALITY RATES AND FEEDING BEHAVIOR OF ADULT Myllocerus undecimpustulatus undatus MARSHALL (COLEOPTERA: CURCULIONIDAE) EXPOSED TO BIOPESTICIDES ON PEACH FOLIAGE IN A FIELD EXPERIMENT ...... 89

Materials and Methods ...... 90 Sleeve Cages...... 90 Plants and Insects...... 91 Fungal and Biochemical Formulations...... 92 Experimental Design...... 92 Leaf Sample Processing...... 94 Statistical Analysis...... 95 Results ...... 95 Discussion ...... 97

7 CONCLUSIONS ...... 111

APPENDIX: PERSONNAL COMMUNICATION ...... 114

LIST OF REFERENCES ...... 115

BIOGRAPHICAL SKETCH ...... 127

LIST OF TABLES

Table page

3-1 Host plant species from which adult Myllocerus undecimpustulatus undatus were collected...... 377

3-2 Mean and range of morphometric details and duration of biological stages of Myllocerus undecimpustulatus undatus development...... 50

4-1 Details of sustained cold exposure and repeated cold exposure experiments on Myllocerus undecimpustulatus undatus adults collected from the field in August 2014 and January 2015...... 64

4-2 Mean percentage survival of field-collected Myllocerus undecimpustulatus undatus adults following exposure to 0° C or -5° C ...... 65

6-1 Mean number of Beauveria bassiana CFUs/mm2 from sprayed and unsprayed peach leaves...... 109

6-2 Mean number of Metarhizium anisopliae and Isaria fumosorosea CFUs/mm2 from sprayed and unsprayed peach leaves...... 110

LIST OF FIGURES

Figure page

2-1 Distribution of Myllocerus undecimpustulatus undatus in Florida based on collection information from the Florida Department of Agriculture and Consumer Services, Division of Plant Industry from 2000 through 2015...... 233

2-2 Adult Myllocerus undecimpustulatus undatus head, rostrum, and antenna...... 244

2-3 Adult Myllocerus undecimpustulatus undatus with dark mottled elytra...... 255

2-4 Adult Myllocerus undecimpustulatus undatus, tridentate metafemur...... 266

2-5 Adult Myllocerus undecimpustulatus undatus compared with Artipus floridanus...... 277

2-6 Adult male and female Myllocerus undecimpustulatus undatus...... 288

2-7 Life stages of Myllocerus undecimpustulatus undatus...... 2929

2-8 Myllocerus undecimpustulatus undatus adult feeding damage...... 300

3-1 Mesh-screened Bug Dorm used to maintain field collected Myllocerus undecimpustulatus undatus for experiments and observation...... 43

3-2 Clear plastic tubes with brown paper covering to simulate darkness...... 44

3-3 Myllocerus undecimpustulatus undatus eggs...... 45

3-4 Myllocerus undecimpustulatus undatus immature stages...... 46

3-5 Ventral, lateral, and dorsal habitus of the pupa of Myllocerus undecimpustulatus undatus with distinct mandibular cusps...... 47

3-6 Teneral adult of Myllocerus undecimpustulatus undatus with distinct mandibular cusps...... 48

3-7 Myllocerus undecimpustulatus undatus male and female mating...... 49

4-1 Myllocerus undecimpustulatus undatus adults in plastic containers with Chrysobalanus icaco leaves and a Petri dish with moistened paper towel...... 63

4-2 Adult Myllocerus undecimpustulatus undatus survival response models over time at 0° C and -5° C. The green line represents observed data...... 66

4-3 Number of dead summer- and winter-collected adult Myllocerus undecimpustulatus undatus after repeated cold exposure and sustained cold exposures...... 67

4-4 Leaf area consumed by summer- and winter-collected adult Myllocerus undecimpustulatus undatus after repeated cold exposure and sustained cold exposures...... 68

4-5 Isothermal line predicting the northern limits of Myllocerus undecimpustulatus undatus distribution based on daily air temperatures of 0° C for at least 4 d. Colored areas in North America where M. undecimpustulatus undatus is predicted to occur by DIVA-GIS BIOCLIM utilizing two climate variables, isothermal temperature, and annual mean precipitation...... 69

5-1 Petri dish cages modified to study Myllocerus undecimpustulatus undatus treatment interactions on treated contained Chrysobalanus icaco leaves for 15-day period...... 82

5-2 Plant Damage Rating Index examples of adult Myllocerus undecimpustulatus undatus feeding on treated Chrysobalanus icaco leaves...... 83

5-3 Comparison of Myllocerus undecimpustulatus undatus survival curves for each treatment trial and combined treatment trials using Kaplan-Meier survival analysis...... 84

5-4 Comparison of combined mean survival time in days of adult Myllocerus undecimpustulatus undatus on treated Chrysobalanus icaco leaves...... 85

5-5 Adult Myllocerus undecimpustulatus undatus mortality after treatments at the end of 15 days...... 86

5-6 Myllocerus undecimpustulatus undatus adults after infection with Beauveria bassiana, BotaniGard...... 87

5-7 Plant damage ratings of treated Chrysobalanus icaco leaves fed on by adult Myllocerus undecimpustulatus undatus at 15 days after treatment application...... 88

6-1 Prunus persica, Peach leaves damaged by adult Myllocerus undecimpustulatus undatus...... 101

6-2 Sleeve cage...... 102

6-3 Peach trees at two locations: Row 16E located on the south side and Row E located on the north side of 33rd Street at the Florida Research Center for Agriculture Sustainability in Vero Beach, FL...... 103

6-4 Plant Damage Rating Index examples of adult Myllocerus undecimpustulatus undatus feeding on treated peach leaves...... 104

6-5 Peach leaf discs taken from leaf samples...... 105

6-6 Adult Myllocerus undecimpustulatus undatus mortality rates 15 days after treatment applications...... 106

6-7 Plant damage ratings of treated peach leaves exposed to adult Myllocerus undecimpustulatus undatus for 15 days after treatment application...... 107

6-8 Myllocerus undecimpustulatus undatus adult mycosing after infection with Beauveria bassiana...... 108

Abstract of Dissertation Presented to the Graduate School of the University of Florida in Partial Fulfillment of the Requirements for the Degree of Doctor of Philosophy

DISTRIBUTION, BIOLOGY, AND MANAGEMENT OF Myllocerus undecimpustulatus undatus MARSHALL (COLEOPTERA: CURCULIONIDAE) IN THE LANDSCAPE AND HORTICULTURE INDUSTRY

Anita Seen Neal

August 2018

Chair: Ronald D. Cave Major: Entomology and Nematology

The Sri Lankan weevil, Myllocerus undecimpustulatus undatus Marshall, is a non-native species of concern to ornamental and tropical fruit growers in Florida.

Establishing feral populations in southern Florida in 2000, there is relatively little information on this weevil’s biology or management. This dissertation examined the life cycle, biotic and abiotic factors that influence survival, and effective control measures.

A plant-based rearing system was developed to capture morphometric details and establish the duration of biological stages in the weevil’s life cycle at 25° C. Eggs hatched in about 6-8 days, larval development ranged from 34 to 43 days, and pupation was approximately 22 days. Total development time from egg to adult ranged from 61 to

74 days.

Two experiments tested the cold tolerance of adults. The first experiment exposed weevils to 0° C and -5° C for various time periods. After 2 days of exposure, survival was 60% and 18% at 0° C and -5° C, respectively. After 4 days of exposure,

11% and 4% of the weevils survived 0° C and -5° C, respectively. In the second

11 experiment, weevils collected in summer and winter were exposed to -5° C, through either a sustained cold period of 10 hours or a repeated cold exposure of 2 hours.

Summer-collected weevils exposed to repeated cold exposure survived more than 3 times longer than those subjected to sustained cold exposure. Average leaf area consumed by winter-collected weevils was 4 times greater than that consumed by summer-collected weevils. A niche distribution model based on collection localities in

Florida projects the potential of the weevil to become established in the southeastern and western USA.

Entrust evaluated in the laboratory and BotaniGard in the laboratory and the field, reduced survival time and consistently performed well in killing M. undecimpustulatus undatus adults. In the laboratory, adult weevil mortality was 169% higher with Entrust and BotaniGard than with PyGanic, AzaMax, PFR-97, Met52, and Sevin. On peach leaves in the field, weevil mortality rates with BotaniGard were approximately fourfold higher than with PFR-97, MET52, and AzaMax. The results of this study should be included in an integrated pest management strategy.

CHAPTER 1 INTRODUCTION

In 2015, the Florida nursery and landscape industry exceeded $21 billion in total output sales, with $4.6 billion in nursery and greenhouse crop production (Hodges et al.

2017), ranking second in the nation behind California. Nursery plants are one of the largest agricultural commodity groups in Florida, together with fruits, vegetables, and forest products (U.S. Department of Agriculture, National Agriculture Statistics Service

2017). Adult leaf-feeding beetles damage the foliage of ornamental plants, fruit trees, and vegetables, and their larvae often injure root systems. One of these beetles,

Myllocerus undecimpustulatus undatus Marshall (Coleoptera: Curculionidae), is a serious pest of concern to the subtropical and tropical fruit, ornamental, and vegetable industries in Florida, due to the insect’s estimated economic impact in billions of dollars

(U.S. Department of Agriculture, National Agriculture Statistics Service 2017).

In 2006, the Florida Department of Agriculture and Consumer Services, Division of Plant Industry confirmed the occurrence of M. undecimpustulatus undatus in 12 southern Florida counties (Mannion et al. 2006) on 81 different plant species in 31 plant families including fruits, nuts, vegetables, and ornamentals (O’Brien et al. 2006). By

2013, this weevil was found in 27 counties feeding on over 150 host plants (Neal 2013) in 47 plant families. The adults often feed en masse on new plant foliage, potentially reducing the quality and quantity of ornamental plants and fruit production

(Thimmegowda et al. 2013). From the time when M. undecimpustulatus undatus was first recorded in Florida in 2000 (O’Brien et al. 2006), it has invaded plant nurseries, garden centers, home landscapes, fruit groves, and botanical gardens (Caldwell 2015,

Mayer and Mannion 2011). The mild Florida climate, abundance of food available year-

13 round, and lack of specific natural enemies provides ideal conditions for survival and reproduction of M. undecimpustulatus undatus.

Decisions for management of invasive insect species requires knowledge of the potential for the insect to cause harmful ecological, economic, or social impacts in areas outside their native range (Venette et al. 2010). Those species identified with this potential can be evaluated through a pest risk analysis to determine the possibility of them effectively invading an area and causing harm (Food and Agriculture Organization

2007). The ability to predict the potential movement and distribution of an invasive insect is a valuable tool within a management system.

Synthetic pesticides are the traditional method of controlling insects that attack crops and ornamentals. However, concern about environmental impacts, an increase in pesticide resistance, and the desire for effective and sustainable pest control point to the necessity of an integrated pest management approach (Leibee and Capinera 1995).

Research on the management of M. undecimpustulatus undatus has focused on synthetic chemical controls, with limited residual effect (Arévalo and Stansly 2009,

Larsen et al. 2017, Mannion et al. 2006), which is not a sustainable approach. The biological control agents that attack Myllocerus undecimpustulatus Faust (Kaur and

Bala 1988, Pruthi and Batra 1960) in their native habitat are not found in Florida. The use of botanical and biological insecticides and entomopathogenic fungi may generate effective and economic short and long-term biological control solutions of this weevil.

The study of Myllocerus species in Asia has yielded information on life cycle, host preferences, economic impacts, and successful control measures (Atwal 1976,

Josephrajkumar et al. 2011, New Pest Advisory Group 2000). However, there is little

14 information on the biology of M. undecimpustulatus undatus, and the dearth of information presents a challenge for designing pest management strategies. Developing a successful integrated approach to management essentially requires information on the biology and ecology of the weevil (Gullan and Cranston 1994). Consequently, a clear understanding of this weevil’s life cycle, the duration of each stage, biotic and/or abiotic factors that influence survival, fecundity, and population fitness is required.

Involving growers and the public by identifying M. undecimpustulatus undatus in their nurseries, landscapes, fields, and gardens and sharing practical solutions is important in promoting the adoption of best management practices. The overall goal of my research was to evaluate pest management tools for control of M. undecimpustulatus undatus in the landscape and horticulture industry. These approaches should support sustainable methodologies to manage this invasive weevil within Florida’s natural and built environment. The research focused on four objectives:

1. Design a successful rearing strategy to identify the duration of each life cycle stage of M. undecimpustulatus undatus.

2. Analyze historical data and measure cold tolerance of M. undecimpustulatus undatus to determine its potential distribution in North America.

3. Assess the mortality rates of adult M. undecimpustulatus undatus exposed to biopesticides in laboratory assays.

4. Assess the mortality rates of adult M. undecimpustulatus undatus exposed to biopesticides on peach (Prunus persica L.) leaves in field trials.

CHAPTER 2 LITERATURE REVIEW

Distribution and Characteristics of Myllocerus Species

Broad-nosed weevils in the genus Myllocerus Schoenherr are elongate beetles, more than twice longer than broad, with their dorsal surface almost convex (more pronounced in females), and the ventral surface flat (Ramamurthy and Ghai 1988).

Ramamurthy and Ghai (1988) state another characteristic feature of this genus is the vestiture in the form of scales and hair-like setae. Some subspecies of Myllocerus undecimpustulatus Faust, especially M. undecimpustulatus maculosus Debrochers des

Loges and M. undecimpustulatus marmoratus Faust (Ramamurthy and Ghai 1988), are considered serious pests of cultivated crops in India, Pakistan, and Sri Lanka.

Myllocerus undecimpustulatus undatus Marshall, commonly called the Sri

Lankan weevil, Asian gray weevil, and yellow-headed ravenous weevil (Hunsberger

2003, Malumphy and Reid 2017, Neal 2013, Thimmegowda et al. 2013), was first detected in the United States on Citrus sp. in Pompano Beach, Broward County, Florida on 15 September 2000. Three specimens were identified by Dr. Charles W. O’Brien, first as M. undecimpustulatus, a species native to southern India, and then again as M. undatus Marshall native to Sri Lanka (O’Brien et al. 2006). Department of Agriculture and Consumer Services, Division of Plant Industry inspectors collected M. undecimpustulatus undatus from multiple locations and noted some were newly emerged, evidence that the species was established in southern Florida (Thomas

2005). Myllocerus undecimpustulatus undatus was detected in 12 counties in 2006 and nine years later in 27 counties in Florida (Figure 2-1). It has not been determined how the Sri Lankan weevil arrived in southern Florida (Frank and Thomas 2009).

Description and Life Cycle

Myllocerus undecimpustulatus undatus is a small (6-8 mm long) broad-nosed weevil, whitish gray with irregular dark markings on the elytra. The head and rostrum are yellowish (Figure 2-2). The compound eyes are black and prominent

(Josephrajkumar et al. 2011). The antennal funicle articulates at an angle to the scape originating from near the apex of the rostrum (Ramanurthy and Ghai 1988) (Figure 2-2).

The elytra are much broader than the prothorax, and the humeri are distinctly angled

(Figure 2-3). The pro- and mesofemora are bidentate, and the metafemur is tridentate

(O’Brien et al. 2006) (Figure 2-4). It is similar in appearance to Artipus floridanus Horn

(Figure 2-5), a common Florida weevil, which differs by the lack of metafemoral teeth and a yellow head.

Females are larger than males, ranging 1-2 mm greater in length and twice the weight (George et al. 2015) (Figure 2-6a). Females are differentiated from males by a black-gray marking extending from the metasternum to the anterior margin of the second abdominal segment (Figure 2-6b) (George et al. 2015). Kiyanthy and Mikunthan

(2009) observed that Myllocerus species mate during the early morning and evening, with a mean copulation period estimated at 44 minutes.

Females may lay up to 360 eggs over a 24-day period, and larvae emerge in 3-5 days (Atwal 1976). Myllocerus undecimpustulatus undatus eggs (Figure 2-7a) are laid directly on organic material at the soil surface, which is common for most Myllocerus species (Atwal 1976, Mayer and Mannion 2011). Eggs are less than 0.5 mm, ovoid, and usually laid in clusters of 8-35. They are white or cream-colored at first, then gradually turn brown when they are close to hatching (Atwal 1976, Kiyanthy and Mikunthan 2009).

The larvae range in size from 1.09 ± 0.05 mm as first instars to 4.0 ± 0.05 mm as fourth 17 instars and are beige-white with a reddish brown head (Atwal 1976, Kiyanthy and

Mikunthan 2009) (Figure 2-7b). They burrow into the soil where they feed on plant roots for approximately one to two months. The larvae pupate (Figure 2-7c) in the soil for approximately one week inside an earthen cell (Atwal 1976). The complete life cycle takes approximately 62.0 ± 2.7 days at 28.9 ± 1° C (Kiyanthy and Mikunthan 2009).

Myllocerus undecimpustulatus undatus is a tropical insect originating from Sri

Lanka, which is within the Equatorial/Tropical climatic zone (Zone I). The temperature within this zone has minimal seasonal fluctuations (Walter et al. 1975) yielding an average temperature of 27° C (Merkel 2016, NOAA 2018). Walter et al. (1975) characterize the coastal climate of Florida as being a humid subtropical type, and

Osborn (2017) projects an average temperature in southern Florida of 22° C. A prediction by New Pest Advisory Group (2000) suggests that if M. undecimpustulatus undatus is limited to the equatorial/tropical climatic zone or the subtropical humid zone

(Zone II), then its range within the United States would be restricted likewise. It is unknown if M. undecimpustulatus undatus undergoes a dormancy phase in Sri Lanka,

India, or the United States.

Host Plants and Insect Damage

Myllocerus undecimpustulatus undatus is a polyphagous herbivore (George et al.

2015, Neal 2013). The adults feed on over 150 host plants that include fruit trees, palms, ornamental plants, and vegetables (Neal 2013). Several tropical fruit trees, Litchi chinensis Sonn (lychee), Dimocarpus logan Lour (longan), Mangifera indica L. (mango), and Persea americana Miller (avocado), are particularly preferred (Mannion et al. 2006,

Neal 2013). The adults consume the young, tender new plant growth. Damage ranges from irregular notching along the leaf margins to more extensive feeding inward along 18 the leaf veins and consumption of the leaf lamina (Josephrajkumar et al. 2011, Mannion et al. 2006) (Figure 2-8).

The range of larval hosts is unknown. O’Brien et al. (2006) noted that larvae burrow through the soil feeding on plant roots. Larvae have been reared in the laboratory on the roots of Ipomoea batatas L. (sweet potato), Capsicum L. (pepper),

Solanum melongena L. (eggplant), and Daucus carota sativus L. (carrot) (O’Brien et al.

2006, Neal 2013). Emergence traps placed along the dripline have captured adults emerging from under mango, lychee, longan, and Pouteria sapota (Jacq.) H.E. Moore &

Stearn (mamey sapote) trees (Epsky et al. 2009).

Management Strategies

Much of the early research to assess the efficacy of products for control of

Myllocerus species was conducted in India. Since the discovery of M. undecimpustulatus undatus in Florida, efforts to manage this pest have been ongoing.

Initial research focused on the presence of biological control agents and evaluation of chemical products.

Pruthi and Batra (1960) reported a braconid wasp, Perilitus mylloceri (Wilkinson)

Nagasawa, attacking adult M. undecimpustulatus maculosus in India. The mature larvae of Perilitus species complete their development in the weevil host, and the adults emerge by cutting a hole through the thin membrane between two segments of the abdomen, usually near the posterior end of the body (Clausen 1940). Unfortunately, there are no known parasitoids of M. undecimpustulatus undatus in Florida, and

19 attempts to find them in Sri Lanka were unsuccessful (R. D. Cave, personal communication1).

Mohandas et al. (2004) demonstrated successful control of larvae of Myllocerus subfasciatus Guerin using a Cry 3 gene toxin from Bacillus thuringiensis Berliner ssp. tenebrionis (Btt). Soil drench with Btt yielded 70% mortality of first instars (Mohandas et al. 2004). Another bioassay treating M. subfasciatus with Cry3A proteins included first instars added to potted brinjal drenched with Btt and adults fed treated brinjal leaves

(Maligeppagol et al. 2012). Of the 30 larvae per pot, only 5 survived on average compared to 26.5 in the control. More than 70% of the adults were dead within 4 days, whereas weevils in the control survived for more than two weeks (Maligeppagol et al.

2012). Another Cry3A protein experiment using leaf dip bioassays against M. undecimpustulatus undatus adults produced mortality ranging from 91 to 100% (Swamy et al. 2013).

Kaur and Bala (1988) described a gregarine parasite, Steinina lunata Kaur and

Bala, in the gut of adult and larval M. undecimpustulatus maculosus collected from cotton fields; infection was heavier during the rainy season. Valigurová (2012) suggested that gregarines are usually not lethal to insect hosts, but may reduce longevity, fecundity, and overall body size of the host. She also pointed out that, in some instances, there is no negative effect on the host or the relationship may even be symbiotic (Summer 1933). Infection of Myllocerus discolor Boheman, a highly destructive pest of Corchorus capsularis L. (jute), by a pathogenic microsporidian

1 R. D. Cave, personal communication letter, in Appendix. 20 parasite, Nosema mylloceri Ghosh, was found in the gut epithelia, fat body, and hemocytes of field-collected adults in India (Ghosh 1990).

Nagesh et al. (2016) investigated entomopathogenic nematodes by comparing three Heterorhabditis species and four Steinerema species for management of M. subfaciatus larvae on eggplant roots. All species caused >80% mortality under laboratory conditions. Gowda et al. (2016) showed that Steinerema carpocapsae

Weiser caused greater mortality of M. subfasciatus pre-pupae and third instars (20–

100% and 16-92%, respectively) than Heterorhabditis indica Poinar, Karunakar, and

David (16-92% and 12-80%, respectively) at high nematode concentrations.

Two entomopathogenic fungi, Beauveria bassiana (Balsamo) Vuillemin and

Metarhizium anisopliae (Metschnikoff) Sorokin, were evaluated in a field in India against

M. undecimpustulatus maculosus by Shanthipriya and Misra (2007). The product

Daman (B. bassiana) provided more than 62% population reduction over the control after 10 days, whereas M. anisopliae was only slightly effective at 11% population reduction over the control after 10 days. Geopalakrishnan and Narayanan (1988) found

B. bassiana naturally infecting M. subfaciatus adults on brinjal in the field in India.

Shanthipriya and Misra (2007) compared the efficacy of biopesticides and a synthetic insecticide to kill M. undecimpustulatus maculosus adults on okra seedlings in

India. Treatments were evaluated 5 and 10 days after application. The systemic neonicotinoid Padan achieved 91% and 57% reduction over the control, respectively.

Servo Agrospray (petroleum horticultural oil) and Ozoneem (neem oil) reduced the weevil population by 49% and 62%, respectively, compared to the control.

Yuma 4E (chlorpyrifos) was sprayed on the ground, the foliage, and combined ground and foliage of Citrus sinensis ’Valencia’ L. to evaluate control of M. undecimpustulatus undatus adults (Arévalo and Stansly 2009). The combined application (ground and foliage) significantly reduced the number of weevils on days 6 and 22 after application. On day 35, there was a resurgence of adult weevils, even in the combined treatment plots (Arévalo and Stansly 2009).

Figure 2-1. Distribution of Myllocerus undecimpustulatus undatus in Florida based on collection information from the Florida Department of Agriculture and Consumer Services, Division of Plant Industry from 2000 through 2015. Map created by Anita Neal using diymaps.net

Figure 2-2. Adult Myllocerus undecimpustulatus undatus with yellowish coloration of head and rostrum. The antennal funicle articulates at an angle (arrow) to the scape originating from near the apex of the rostrum. Photographs by Anita Neal.

Figure 2-3. Adult Myllocerus undecimpustulatus undatus with dark mottled elytra much broader than the prothorax and the humeri (arrow) distinctly angled. Photograph by Anita Neal.

Figure 2-4. Adult Myllocerus undecimpustulatus undatus, tridentate metafemur (arrow). Photograph by Paul Skelley.

a b

Figure 2-5. Adult Myllocerus undecimpustulatus undatus (a) compared with Artipus floridanus (b). Photographs by Paul Skelley.

a b

Figure 2-6. Adult male and female Myllocerus undecimpustulatus undatus: (a) lateral view showing the difference in size, (b) ventral view comparing black-gray marking extending from the metasternum to the second abdominal segment on the female versus only on the metasternum of the male (dashed circle). Photographs by Anita Neal (a) and Justin George (b) (George et al. 2015).

a b

c d

Figure 2-7. Life stages of Myllocerus undecimpustulatus undatus: (a) eggs, (b) larva, (c) pupa, and (d) adult. Photographs by Ronald Cave.

a b c d

e f g h

Figure 2-8. Myllocerus undecimpustulatus undatus adult feeding damage on eight host plants. Photographs by Holly Glenn (a), Susan Halbert (b), and Anita Neal (c- h).

CHAPTER 3 BIOLOGY AND REARING OF Myllocerus undecimpustulatus undatus MARSHALL (COLEOPTERA: CURCULIONIDAE)

Coleoptera is the most speciose order of insects, accounting for 40% of all described arthropod species (Zhang et al. 2018, Stork et al. 2015). The family

Curculionidae has over 60,000 species worldwide, the majority of them plant feeders

(Anderson 2002). Myllocerus undecimpustulatus undatus Marshall, a polyphagous broad-nosed weevil in the subfamily Entiminae, tribe Cyphicerini, and subtribe

Myllocerina, is commonly called the Sri Lankan weevil (Malumphy and Reid 2017, Neal

2013). Acquiring the common name from its country of origin, Sri Lanka, M. undecimpustulatus undatus is recorded in India and Pakistan as a pest of vegetables, fruits, and ornamentals (Malumphy and Reid 2017, Thimmegowda et al. 2013, Thomas

2005). Myllocerus undecimpustulatus undatus arrived in Florida in 2000, and the adults feed on over 150 different host plants (Table 3-1). The larvae fed on the roots of the adult host plant (O’Brien et al. 2006).

Little has been published in the scientific literature regarding the biology or rearing of M. undecimpustulatus undatus. The information known comes mostly from research in Asia, specifically India, with different species of Myllocerus. To develop an insect rearing program, a basic understanding of the insect’s biology, behavior, and physiology are essential (Shapiro 1992). The purpose of this study was to design a successful rearing strategy and to identify the duration of each life cycle stage of M. undecimpustulatus undatus. The rearing system is described in this chapter.

Materials and Methods

Weevil Collection and Colony.

A laboratory colony of M. undecimpustulatus undatus was established with adults collected from mature Australian pine, Casuarina equisetifolia L.

(Casuarinaceae), in Fort Pierce, FL (27° 29′ 15.43″ N, 80° 24′ 33.79″ W). The weevils were maintained in mesh-screened Bug Dorms (60 × 60 × 60 cm, BioQuip Products,

Rancho Dominguez, CA) with the temperature at 24° C ± 2°, 60% RH, and 14 h photoperiod. Three 40-dram vials with snap-on lids with a 1-cm hole cut into the lid were filled with water, and 2-3 branches of cocoplum, Chrysobalanus icaco L.

(Chrysobalanaceae), 25–30 cm long were inserted through the hole in the lid. In each

Bug Dorm, three plastic containers 13 cm high × 12 cm wide (at top) and narrowing to

10 cm (at bottom) were lined with brown paper towel for an oviposition substrate, and one vial with cocoplum was placed into a container. Two small Petri dishes (60 × 15 mm) were filled with a moistened, crumpled piece of brown paper towel to provide water to the weevils (Figure 3-1). Plants were changed weekly, and water was added to the dishes 2-3 times per week.

Rearing System.

Dr. Geetha Thimmegowda worked with M. undecimpustulatus undatus in a portion of her graduate work. My rearing system was modified from her design and is described in this chapter.

For studying the life cycle, eggs were collected 3-4 times per week for a total of eight weeks from the detritus collected on paper towels. The eggs were transferred using a camel hair brush to a Petri dish (100 × 15 mm) lined with No. 4 filter paper moistened with 800 µL of distilled water and observed daily. The dish was covered with

32 brown paper towel to simulate darkness and placed in an incubator at 25° C ± 2° and

60% RH. The number of eggs collected and hatched was recorded each week. The incubation period was based on time of egg collection and averaged for that week.

An artificial diet designed by Lapointe et al. (2008) in 30-ml plastic cups was inoculated with five neonate larvae (<24-h-old). Five cups were inoculated as a trial experiment. The diet cups were capped and placed in trays enclosed in zip-lock plastic bags and held in a dark incubator at 25° C ± 2° and 60% RH. The cups were opened 4 weeks after inoculation. No live larvae were found, and this method was discontinued.

A plant-based rearing system was designed for larval development studies.

African finger millet, Eleusine coracana (L.) Gaertn. was grown from seed (Pure Indian

Foods, Princeton Junction, NJ) in clear plastic square sealed bottom tubes (15.5 cm ×

3.8 cm, Cleartec Packaging, Park Hills, MO). Three 1.5 mm holes were drilled into the bottom of the tubes to provide drainage. A mixture of 50% potting mix and 50% play sand was combined and autoclaved at 120° C ± 5°. Ten tubes were filled with the soil mixture, 25-30 seeds spread on top, and watered thoroughly until excess water drained.

A brown paper towel was folded, wrapped around each tube, secured with a rubber band and labeled with the date. Tubes were placed in an incubator at 25° C ± 2°, 60%

RH and 14 h photoperiod. The position of the tube was kept slanted to keep root growth in one direction, and to enable observation of larval feeding (Figure 3-2). African finger millet germinated in 3-5 days. When seedlings grew to about 50 mm, a small impression was made in the soil, and 25 neonate larvae from detritus collected on paper towel were transferred to it by using a soft brush. Seedlings were watered as needed. Ten new tubes were added every two weeks. Observations of larval development were made

33 approximately every 7-10 days. Five tubes were dissected at different time intervals (2-

12 weeks) to record larval growth and to take measurements of instars and pupae. All measurements were calibrated using DinoCapture 2.0 software (AnMo Electronics

Corporation, New Taipei City, Taiwan) to 0.001 mm.

Results

Eggs were laid singly and in clusters of 3-15 eggs. The eggs were initially light tan in color, changing to a brown indicating they were close to hatching. Eggs measured on average 0.210 ± 0.002 mm wide and 0.542 ± 0.002 mm long. (Figure 3-3, Table 3-2).

Six hundred eighty eggs were collected over the eight-week period, of which 639 (94%) hatched. The incubation period ranged 6-9 days, with an average of 7.0 ± 0.4 days

(Table 3-2).

Larvae are beige-white with a reddish brown head. The sclerotization of the mandibles increased from one instar to the next. All body segments are covered with asperities and golden-tinged, fine white hairs. There are not many structural differences between instars (Figure 3-4). The duration of first through fourth instar stages averaged

39.0 ± 0.4 days (Table 3-2).

Pupae measured on average 1.864 ± 0.001 mm wide and 5.041 ± 0.001 mm long. The pupation period lasted approximately 21.5 ± 0.4 days (Table 3-2). The pupa is exarate (Figure 3-5) and initially ivory-white, gradually turning a light yellowish white.

Teneral adults emerged from the soil approximately 67.5 ± 2.7 days after eggs are laid. The newly emerged adult is light yellowish brown in color, soft-bodied, and maintains the mandibular cusps used to cut through the soil (Figure 3-6). The cusps are shed in <1 day.

Female adult weevils collected in the field measured almost 2 mm longer than males, i.e. 8.694 ± 0.002 vs 6.822 ± 0.001, respectively. Mating was frequently observed, several times a day, especially when weevils experienced stressful situations, e.g., moving them into small containers for experiments (Figure 3-7).

Discussion

This is the first laboratory study examining the biology of M. undecimpustulatus undatus using a plant-based rearing method. Kiyanthy and Mikunthan (2009) used a plant-based rearing method with four Myllocerus species (M. viridis Aurivillius, M. discolor Boheman, M. subfaciatus Guerin, and M. maculosus Desbrochers des Loges).

Larvae were fed rootlets of Vernonia cinerea L. in a sterile soil-based rearing cage.

Adults were fed leaves of Azadirachta indica L. The mean egg incubation period for the four Myllocerus species was 4.6 ± 0.5 days at 28.9° C, similar to the results of Atwal

(1976), who reported eggs hatched in 3-5 days. In my study, the mean egg incubation period for M. undecimpustulatus undatus was 7.0 ± 0.4 days at 25° C. The mean larval development period for the four Myllocerus species was 50.6 ± 1.5 days at 28.9° C, again similar to Atwal’s (1976) results of development completed in 1-2 months

(Kiyanthy and Mikunthan 2009). By comparison, the mean larval development period for

M. undecimpustulatus undatus was 39.0 ± 0.4 days at 25° C in my study. The difference in development time for my study versus previous studies could be due to the higher incubation temperature in the earlier studies.

Kiyanthy and Mikunthan (2009) found the mean pupal period lasted 7.8 ± 0.8 days at 28.9° C, similar to the one-week period reported by Atwal (1976). The mean pupal period for M. undecimpustulatus undatus in my study was 21.5 ± 0.4 days at

25° C, three times longer than for the Myllocerus species studied by Kiyanthy and 35

Mikunthan (2009). Pupation of Myllocerus species occurs within an earthen cell (Atwal

1976, Epsky et al. 2009, Kiyanthy and Mikunthan 2009, Talwar 2014), however, only one pupal cell was evident in my plant-based rearing system. The dehiscent cusps at the apex of the mandibles are prominent in pupae and present on teneral adults

(Figures 3-5 and 3-6) but usually detach as the pupal chamber is torn open for emergence (May 1966).

Sexual dimorphism in size was distinct in adults, with males generally smaller than females, confirming the same observation by George et al. (2015). In the other life stages, a size difference between male and female was not detected.

The life cycle of M. undecimpustulatus undatus was completed in 67.5 ± 2.7 days

(9.5 weeks) at 25° C, 1.5 weeks longer than the 6-8 weeks observed by Atwal (1976) and Mannion et al. (2006) but similar that of the four Myllocerus species studied by

Kiyanthy and Mikunthan (2009). In my rearing system, E. coracana, an annual grass, was planted to provide a food source for larvae. Kiyanthy and Mikunthan (2009) fed their Myllocerus larvae V. cinerea, a broadleaf plant in the family Asteraceae. Different larval host plants could affect larval development and should be explored along with possible artificial diets to identify an efficient and inexpensive rearing system that produces larger numbers of third instars for experimentation.

Table 3-1. Host plant species from which adult Myllocerus undecimpustulatus undatus were collected by Florida Department of Agriculture Consumer Services, Division of Plant Industry field inspectors and Anita Neal*.

FAMILY SCIENTIFIC NAME COMMON NAME 1 Acanthaceae Blechum pyrmidatum (Lam.) Urb. Browne's Blechum 2 Aceraceae Acer rubrum L. Red Maple 3 Anacardiaceae Anacardium occidentale L. Cashew 4 Anacardiaceae Ambrosia artemisiifolia L. Common Ragweed 5 Anacardiaceae Mangifera indica L. Mango 6 Anacardiaceae Schinus terbinthifolius Raddi Brazilian Pepper* 7 Anacardiaceae Sclerocarya birrea (A. Rich.) Hochst Marula 8 Aquifoliaceae Ilex cassine L. Dahoon Holly 9 Acecaceae Adonidia merillii Christmas Palm 10 Acecaceae Caryota mitis Lour Burmese Fishtail Palm 11 Acecaceae Cocos nucifera L. Coconut Palm 12 Acecaceae Dypsis lutesceens (H. Weendl.) Beentje & J. Dransf. Areca Palm 13 Acecaceae Livistona chinensis (Jacq.) R.BR ex Mart. Chinese Fan Palm 14 Acecaceae Phoenix roebellenii O'Brien Pygmy Date Palm 15 Acecaceae Washingtonia robusta H. Wendl. Washington Palm 16 Asteraceae Baccharis halimifolia L. Salt Bush 17 Asteraceae Bidens alba L. Common Beggarticks 18 Asteraceae Bidens pilosa L. Spanish needles 19 Asteraceae Zinna elegans L. Zinnia 20 Calophyllaceae Mammea americana L. Mamey-apple 21 Cannabaceae Humulus lupulus L. Common Hop* 22 Caricaceae Carica papaya L. Papaya 23 Casuarinaceae Casuarina equisetfolia L. Australian Pine 24 Chenopodiaceae Spinacia oleracea L. Spinach 25 Chrysobalanaceae Chrysobalanus icaco L. Cocoplum 26 Combretaceae Bucida buceras L. Black Olive 27 Combretaceae Conocarpus erectus L. Buttonwood 28 Combretaceae Conocarpus erectus v. sericeus L. Silver Buttonwood*

Table 3-1. Continued

FAMILY SCIENTIFIC NAME COMMON NAME 29 Combretaceae Laguncularia racemosa L. White Mangrove 30 Combretaceae Quisqualis indica L. Ragoon Creeper 31 Combretaceae Terminalia catappa L. Tropcal Almond 32 Convolvulaceae Ipomoea batatas (L.) Lam. Sweet Potato 33 Cupressaceae Platycladus orientalis L. Oriental Arborvitae 34 Cycadaceae Cycas sp. Sago 35 Ebenaceae Diospyros digyna Jacq. Black Sapote 36 Ebenaceae Diospyros virginiana L. Common Persimmon 37 Elaeagnaceae Elaeagnus pungens Thunb. Silverthorn* 38 Elaeagnaceae Muntingia calabura L Strawberry tree 39 Euphorbiaceae Acalypha wilkesiana Müll. Arg. Copper Leaf 40 Euphorbiaceae Cnidoscolus chayamansa McVaugh Spinach tree 41 Euphorbiaceae Euphorbia hypericifolia (L.) Graceful Sandmat 42 Euphorbiaceae Jatropha curcas L. Physic Nut 43 Euphorbiaceae Poinsettia cyathophora (Murray) Bartling Paintedleaf 44 Euphorbiaceae Ricinus communis L. Castorbean 45 Fabaceae Acacia auriculiformis A. Cunn ex Benth. Earleaf Acacia 46 Fabaceae Albizia lebbeck (L.) Benth. Woman's tongue 47 Fabaceae Bauhinia x blakeana Dunn. Hong Kong Orchid tree 48 Fabaceae Bauhinia purpurea Wall. Purple Orchid tree 49 Fabaceae Bolusanthus speciosus (Bol.) Harms. Tree Wisteria 50 Fabaceae Bombax ceiba L. Red Silk-Cotton tree 51 Fabaceae Caesalpinia bonduc (L.) Roxb. Gray Nickerbean 52 Fabaceae Calliandra emarginata Benth. Powderpuff 53 Fabaceae Calliandra haematocephala Hassk. Red Powderpuff 54 Fabaceae Dalbergia ecastaphyllum (L.) Taub. Coinvine 55 Fabaceae Desmodium floridanum Chapm. Florida Ticktrefoil (beggarweed)

Table 3-1. Continued

FAMILY SCIENTIFIC NAME COMMON NAME 56 Fabaceae Desmodium tortuosum (Sw.) DC. Dixie Ticktrefoil (beggarweed) 57 Fabaceae Erythrina sp. Erythrina 58 Fabaceae Glycine max (L.) merr Soybean 59 Fabaceae Lysiloma latisiliquum (L.) Benth. Wild Tamarind 60 Fabaceae Mucuna pruriens (L.) DC. Velvet Bean 61 Fabaceae Pithecellobium unguis-cati (L.) Benth. Catclaw Blackbeard 62 Fabaceae Pongamia pinnata (L.) Pierre Pongam 63 Fabaceae Senna Pendula (Humb & Bonpl. Ex Wild) Irwin & Barneby Climbing Cassia 64 Fabaceae Senna surattensis (Hburm.f.) Irwin & Barneby Glossy Shower 65 Fabaceae Sophora tomentosa L. Yellow Necklacepod 66 Fagaceae Quercus laurifolia Michx. Laurel oak 67 Fagaceae Quercus virginiana L. Live Oak 68 Flacourtiaceae Flacourtia indica (Burm. f.) Merr. Govenor's Plum 69 Iridaceae Neomarica gracilis (Herb.) Sprague. Walking Iris 70 Lythraceae Cuphea hyssopifolia Kunth False Mexican Heather 71 Lythraceae Lagerstroemia indica L. Crepe Myrtle 72 Malvaceae Brachychiton rupestris (T. Mitch. ex Lindl.) K. Schum Queensland Bottle tree 73 Malvaceae Ceiba speciosa (A. St.-Hil.) Ravenna Silk Floss Tree 74 Malvaceae Gossypium hirsutum L. Upland Cotton 75 Malvaceae Herissantia crispa (L.) Briz. Bladdermallow 76 Malvaceae Hibiscus rosa-sinesis L. Hibiscus 77 Malvaceae Hibiscus sabdariffa L. Roselle 78 Malvaceae Hibiscus tiliaceus L. Mahoe 79 Malvaceae Malvaviscus arboreus Dill. ex Cav. var. drummondii (Torr. & A. Gray) Schery Texas Wax Mallow, Turk's Cap 80 Malvaceae Malvaviscus penduliflorus DC. Mazapan, Turk's Cap 81 Malvaceae Pachira aquatica Aubl. Guiana Chetsnut 82 Malvaceae Pavonia bahamensis A.S. Hitchc. Pavonia 83 Malvaceae Pseudobombax ellipticum (Kunth) Dugand Shaving Brush tree 84 Malvaceae Sida acuta Burm. f. Common wireweed 85 Malvaceae Sida cordifolia L. Ilima

Table 3-1. Continued

FAMILY SCIENTIFIC NAME COMMON NAME 86 Malvaceae Sterculia sp. Sterculia 87 Malvaceae Theobroma cacao L. Cacao tree 88 Meliaceae Swietenia mahagoni (L.) Jacq. Mahogany 89 Moraceae Ficus aurea L. Strangler Fig 90 Moraceae Ficus benjamina L. Weeping Fig 91 Moraceae Ficus carica L. Common Fig 92 Moraceae Ficus microcapa L. f. Chinese Banyan 93 Moraceae Ficus tinctoria G. Forst. Dye Fig 94 Moraceae Morus alba L. White Mulberry 95 Myrsinaceae Ardesia elliptica Thunb. Shoebutton Ardisia 96 Myrtaceae Callistemon viminalis (Sol. Ex Gaertn.) G.Don. Weeping Bottlebrush 97 Myrtaceae Corymbia torelliana (F. Muell.) K.D. Hill & L.A.S Johnson Torell's Eucalyptus* 98 Myrtaceae Eugenia uniflora L. Surinam Cherry 99 Myrtaceae Morella cerifera (L.) Small Wax Myrtle* 100 Myrtaceae Myricaria cauliflora (DC.) O. Berg. Jaboticaba 101 Myrtaceae Syzgium cuminii (L.) Skeels Java Plum 102 Myrtaceae Syzgium paniculatum Gaertn. Bush Cherry 103 Nyctaginaceae Bougainvillea sp. Bougainvillea 104 Nyctaginaceae Guapira discolor (Spreng.) Little Bolly, Beefwood 105 Oleaceae Olea europaea L. Olive 106 Oleaceae Osmanthus americanus (L.) Benth. & Hook. f. ex A. Gray Wild Olive 107 Oxalidaceae Averrhoa carambola L. Carambola 108 Passifloaceae Passiflora sp. Passionflower 109 Phytolaccaceae Ribina humilis L. Rougeplant 110 Plumbaginaeae Plumbago auriculata Lam. Cape Leadwort 111 Poaceae Bambusa sp. Bamboo 112 Poaceae Dactyloctenium aegyptium (L.) Willd. Egyptian Grass 113 Polygalaceae Polygala cowellii (Britton) S.F. Blake Violet Tree 114 Polygonaceae Antigonon leptopus Hook. & Arn. Coral Vine

Table 3-1. Continued

FAMILY SCIENTIFIC NAME COMMON NAME 115 Polygonaceae Coccoloba diversifolia Jacq. Pigeon Plum 116 Polygonaceae Coccoloba uvifera (L.) L. Seagrape 117 Rhamnaceae Krugiodendron ferreum (Vahl) Urb. Black Ironwood 118 Rosaceae Eriobotrya japonica Lindl. Loquat 119 Rosaceae Malus pumila Mill. Apple 120 Rosaceae Photinia serrulata Lindl. Photinia 121 Rosaceae Prunus persica (L.) Ratsch. Peach 122 Rosaceae Rhaphiolepis umbellata (Thunberg) Makino Japanese Hawthorn 123 Rosaceae Rosa sp. Rose 124 Rubiaceae Ixora coccinea L. Scarlet Jungleflame 125 Rubiaceae Psychotria nervosa SW. Wild coffe 126 Rubiaceae Rondeletia leucophylla H. В. K. Nov. Gen. & Sp. Panama Rose 127 Rubiaceae Spermacoce verticillata L. Shrubby False Buttonweed 128 Rutaceae Aegle marmelos (L.) Corr. Serr. Indian Bael 129 Rutaceae Citrofortunella microcarpa (Bunge) Wijnands [Citrus reticulata × Fortunella japonica] Calamondin 130 Rutaceae Citrus sp. Citrus 131 Rutaceae Citrus x paradisi Macfad. Grapefruit 132 Rutaceae Murraya paniculata (L.) Jack Orange Jasmine, Mock Orange 133 Sapindaceae Blighia sapida K.D. Koenig Akee 134 Sapindaceae Cupaniopsis anacardiodes (A. Rich.) Radlk. Carrotwood 135 Sapindaceae Dimocarpus longan Lour. Logan 136 Sapindaceae Litchi chinensis Sonn. Lychee 137 Sapindaceae Melicoccus bijugatus Jacq. Spanish Lime 138 Sapotaceae Chrysophyllum oliviforme L. Satin Leaf 139 Sapotaceae Pouteria sapota (Jacq.) H.E. Moore & Stearn Mamey Sapote 140 Solanaceae Brugmansia suaveolens (Willd.) Angel Trmpet 141 Solanaceae Capsicum sp. Pepper 142 Solanaceae Solanum diphyllum L. Twoleaf Nigtshade

Table 3-1. Continued

FAMILY SCIENTIFIC NAME COMMON NAME 143 Solanaceae Solanum melongena L Eggplant 144 Turneraceae Turnera ulmifolia L. Yellow Alder 145 Ulmaceae Celtis laevigata Willd. Sugarberry, Hackberry 146 Ulmaceae Trema micrantha (L.) Blume Jamaican nettletree 147 Ulmaceae Ulmus americana L. American Elm 148 Verbenaceae Duranta erecta L. Golden Dewdrops 149 Verbenaceae Lantana camara L. Lantana 150 Verbenaceae Petrea volubilis L. Queen's Wreath 151 Vitaceae Parthenocissus quinquefolia (L.) Planch. Virginia Creeper 152 Vitaceae Tetrastigma voinierianum (Mottet) Gagnep. Chestnut Vine 153 Vitaceae Vitis rotundifolia Michx. Muscadine Grape

Figure 3-1. Mesh-screened Bug Dorm used to maintain field-collected Myllocerus undecimpustulatus undatus for experiments and observation. Photograph by Anita Neal.

Figure 3-2. Clear plastic tubes wrapped with brown paper to simulate darkness. The tubes were planted with Eleusine coracana as a food source for Myllocerus undecimpustulatus undatus larvae and angled to keep root growth in one direction. Photographs by Anita Neal.

a b

Figure 3-3. Myllocerus undecimpustulatus undatus (a) newly laid eggs and (b) a cluster of eggs. Darker color indicates close to hatching. Photographs by Anita Neal (a) and Ronald Cave (b).

a b

c d

e f

Figure 3-4. Myllocerus undecimpustulatus undatus (a) neonate larva, (b) first instar, (c) second instar, (d) third instar, (e) fourth instar, and (f) measurement of third instar (head capsule length = 0.519 mm and length of larva = 2.088 mm). Photographs by Ronald Cave (a) and Anita Neal (b-f).

a b c

Figure 3-5. (a) Ventral, (b) lateral, and (c) dorsal habitus of the pupa of Myllocerus undecimpustulatus undatus with distinct mandibular cusps (circled in (a) ventral habitus). Photographs by Ronald Cave.

Figure 3-6. Teneral adult of Myllocerus undecimpustulatus undatus with distinct mandibular cusps (arrow). Photographs by Anita Neal.

a b

Figure 3-7. Myllocerus undecimpustulatus undatus (a) adult female and male and (b) multiple males attempting to mate with a female. Photographs by Anita Neal.

Table 3-2. Mean (± SEM) and range of morphometric details and duration of life stages of Myllocerus undecimpustulatus undatus development at 25° C.

Stage of Sample Width (mm) Length (mm) Duration (Days) development size Mean Range Mean Range Mean Range Eggs 40 0.210 ± 0.002 0.190 - 0.221 0.542 ± 0.002 0.521 - 0.562 7.0 ± 0.4 6 - 8 1st instar 40 0.261 ± 0.001 0.260 - 0.270 1.104 ± 0.004 1.090 - 1.168 9.0 ± 0.4 8 - 10 2nd instar 25 0.482 ± 0.001 0.475 - 0.489 1.162 ± 0.001 1.153 - 1.168 8.0 ± 0.5 7 - 9 3rd instar 15 1.020 ± 0.010 1.014 - 1.025 2.191 ± 0.001 2.186 - 2.195 9.0 ± 0.4 7 - 10 4th instar 10 1.692 ± 0.001 1.688 - 1.696 4.282 ± 0.001 4.276 - 4.285 13.0 ± 0.4 12 - 14 Pupa 10 1.864 ± 0.001 1.857 - 1.870 5.041 ± 0.001 5.037 - 5.045 21.5 ± 0.4 21 - 23 Adult (male) 25 2.282 ± 0.002 2.279 - 2.300 6.822 ± 0.001 6.816 - 6.828 egg to adult egg to adult 67.5 ± 2.7 61 - 74 Adult (female) 25 2.572 ± 0.001 2.567 - 2.582 8.694 ± 0.002 8.680 - 8.710

CHAPTER 4 ADULT COLD TOLERANCE AND POTENTIAL NORTH AMERICAN DISTRIBUTION OF Myllocerus undecimpustulatus undatus MARSHALL (COLEOPTERA: CURCULIONIDAE)

Upon arrival to a new region, tropical insects face biotic and abiotic factors that might limit their establishment and subsequent colonization (Andrewartha and Birch

1954). Winter temperature is probably the most important abiotic factor limiting the establishment of tropical insects in subtropical and temperate regions (Singh et al.

2009). However, insects exposed to cooler temperatures may evolve morphological, physiological, and behavioral adaptations (Lalouette et al. 2007), which could potentially enable them to colonize new regions (Bale 2002). Three examples of tropical insects that overcame winter temperatures to establish in Florida are the lobate lac scale,

Paratachardina pseudolobata Kondo and Guillan (Chong et al. 2008), the West Indian drywood termite, Cryptotermes brevis Walker (Scheffrahn and Su 1999), and the papaya mealybug, Paracoccus marginatus Williams and Granara de Willink

(Amarasekare et al. 2008). Moreover, a tropical insect’s capacity to develop within a broad range of temperatures is a key adaptation to endure changing climatic conditions, which is important in predicting insect outbreaks and developing effective management strategies (Amarasekare et al. 2008, Bale 2002, Manrique et al. 2012).

One of the tools utilized to evaluate the potential spread of an insect is a species distribution model (SDM). The model employs parameters from climatic variables, such as rainfall and temperature, where a species currently exists to predict the possibility of the species survival in other areas that share similar climatic conditions (Stratman et al.

2014). Species distribution models apply geographic information systems (GIS), along with computer software packages and climate databases, to predict a particular species’

51 spread (Elith and Leathwick 2009, Franklin 2010, Stratman et al. 2014). These models have been used to predict the geographical distribution of adventive species, biological control agents, and crop pests into other regions (e.g., Lapointe et al. 2007, Stratman et al. 2014, Tognelli et al. 2009). Data from laboratory or field experiments on cold tolerance may improve the SDM projections (e.g., Diaz et al. 2008, Manrique et al.

2008, 2012).

Cold tolerance studies usually focus on the effects of a single cold exposure on insects under laboratory conditions (Lalouette et al. 2007). However, insects experience variable cold exposures in natural environments even several times within a “winter” season (Marshall and Sinclair 2012). The fluctuations in temperature between night and day may be up to 20° C, exposing them to cyclical cold stress (Marshall and Sinclair

2010). Because fluctuating thermal regimes are common in natural habitats, insects that are active year-round may take advantage of periods of favorable temperatures to feed or recover from damages caused by low temperatures (Lalouette et al. 2007). Marshall and Sinclair (2010, 2012) looked at repeated cold exposures (RCE) with Drosophila melanogaster Meigen (Diptera: Drosophilidae) crossing the chill-coma threshold (at or below 10° C) at which all muscular movement ceases. They found that repeated cold exposures improved cold tolerance when compared to insects experiencing a single, prolonged cold exposure.

Myllocerus undecimpustulatus undatus Marshall is a tropical broad-nosed weevil from Sri Lanka (Ramamurthy and Ghai 1988). Apparently, it does not undergo diapause; it is active and reproducing all year, although activity and reproduction are reduced in winter months in Florida. Despite its pest status, there is limited information

52 regarding the influence of temperature on survival and the potential geographic range of

M. undecimpustulatus undatus in North America.

The objectives of this study were to 1) determine survivorship of adult M. undecimpustulatus undatus after cold temperature exposures, 2) measure survival and food consumption by adults after repeated cold exposure, and 3) use cold tolerance data and climate niche modeling to determine the potential distribution of M. undecimpustulatus undatus in North America. My null hypothesis is that M. undecimpustulatus undatus, being a tropical insect, cannot expand its distribution outside of tropical and subtropical regions.

Materials and Methods

Weevil Collection and Maintenance. Adult M. undecimpustulatus undatus were collected from mature Australian pine, Casuarina equisetifolia L. (Casuarinaceae), in Fort Pierce, FL (27° 29′ 15.43″ N, 80° 24′ 33.79″ W) about 1 week prior to each trial.

The weevils were maintained in mesh-screened Bug Dorms at 24° C ± 2°, 60% RH, and

14 h photoperiod. They were fed cocoplum leaves, Chrysobalanus icaco L.

(Chrysobalanaceae) (as described in Chapter 3).

Cold Tolerance.

Two experiments were conducted to test adult cold tolerance with three trials each (June, July, and August) with a minimum temperature of 0° C or -5° C. In all trials, the plastic containers (42 × 29 × 9 cm Rubbermaid® Newell Brands, Atlanta, GA) used to contain adults were lined with brown paper towel. An 18 × 13-cm piece of the container’s lid was replaced with fine mesh screen for ventilation. Two florist vials, each inserted with one 35-38 cm long branch of cocoplum with leaves, was placed in each container, along with a small Petri dish (60 × 15 mm) filled with a crumpled piece of

53 brown paper towel that was moistened to provide water to the weevils (Figure 4-1). All containers initially housed 25 field-collected adult weevils of unknown age and sex, held at 20° C for 24 h, and then used for exposure to cold temperatures in each experiment.

Three control containers with 25 weevils each were held at 20° C throughout each experiment.

In one experiment, the weevils under cold temperature treatments were gradually acclimated to lower temperatures from 20° C to a final test temperature of 0° C at intervals of 5° C per 24 h in environmentally controlled chambers with 60% RH and 14 h photoperiod. At the end of each exposure interval of 0.5, 1, 2, 4, and 8 d at 0° C, five weevils were removed from each container (15 weevils/trial, 45 weevils per minimum temperature exposure interval across three trials) and placed in a Petri dish (35 × 10 mm) lined with Whatman® grade No.1 filter paper moistened with 200 μl of distilled water. The Petri dishes were held at room temperature (23-26° C) for 24 h, and then dead and live weevils were counted. A weevil was scored alive if leg or antennal movement was observed initially or after gentle prodding. The same procedure was used for the three trials of the experiment with minimum temperature at -5° C, except that adults were exposed to 0° C for 24 h before exposure to -5° C for 0.5, 1, 2, 4, and 8 d. At the same time cold treatment weevils were sampled in both experiments, five control weevils were sampled from each container at 20° C and evaluated for survival in a manner similar to those under cold treatments.

Percentages of adult survival were compared among different exposure intervals within each cold tolerance experiment, including survival without cold exposure. The

Shapiro-Wilk W test and normality plots validated the assumptions of parametric

54 analysis using the Univariate procedure (SAS Institute, 2009). Data were subjected to

ANOVA using the GLM procedure to evaluate treatment effects on the weevil, and treatment means were separated using LSD contingent on a significant treatment effect

(P < 0.0001). The effect of temperature and exposure time was also analyzed by a logistic model using the software JMP and the inverse prediction function (SAS Institute

2017, JMP®PRO 13.1, Cary, NC). This analysis estimated the times required to kill 50%

(LT50) and 90% (LT90) of the adult population at a specific temperature.

Sustained Cold Exposure (SCE) Versus Repeated Cold Exposure (RCE).

Adult weevils used in the experiments were collected from Australian pine on

August 17, 2014 (summer) and January 17, 2015 (winter). Approximately one week prior to weevil collections, temperatures at the collection site ranged 20-34° C in August and 6-26° C in January (Florida Automated Weather Network 2017). Cocoplum branches with leaves of similar age and size (25 cm) were washed with water and allowed to air dry. Each treatment/exposure time utilized five replicated plastic containers (27 × 16 × 8 cm, Great Value® Wal-Mart Stores, Inc., Bentonville, AR). Ten weevils from the same collecting event were added to each container along with one florist tube filled with water supporting one cocoplum branch (sample size/treatment =

50 weevils) (Figure 4-2).

All experiments were conducted in environmental chambers with 60% RH and a

14 h photoperiod. Weevils in the sustained cold exposure treatments were gradually acclimated from 20° to -5° C as described for the cold tolerance experiments, at which time they were held at -5° C for 10 h and returned to 20° C. Weevils in the repeated cold exposure treatments were initially gradually acclimated to -5° C and then held for one

(I), two (II), three (III), or four (IV) 2-h exposure periods at -5° C, each exposure period 55 separated by 22 h at 20° C (Table 4-1). Twenty-four h after the end of the final low temperature exposure period, adult survival was confirmed by leg or antennal movement. Control weevils were maintained at 20° C and evaluated for survival the same as those in the sustained cold exposure and repeated cold exposure experiments.

Survivors remained in their original containers and received new food and water. Leaf consumption was determined four days later by quantifying the unconsumed area of each leaf lamina using ImageJ software (Rasband 2014). This Java-based application for analyzing images re-creates the original leaf shape by filling in areas lost to herbivory. The amount of herbivory was calculated as the difference between pre- and post-herbivory surface areas.

Data were analyzed for effects of cold treatment (sustained cold exposure and repeated cold exposure) and time of year (summer versus winter) on adult survival and herbivory. Normality of the survival data was checked in JMP Pro using the Shapiro-

Wilk Test. The p-value was <0.0001 to reject the null hypothesis and indicate the data does not come from a normal distribution. Survival and leaf are consumption were compared with a one-way ANOVA, and means were separated with Tukey’s HSD test

(α = 0.05) (SAS Institute 2009).

Niche Modeling.

Historical collection data of M. undecimpustulatus undatus from reports in Florida were obtained from Michael Thomas of the Florida Department of Agriculture and

Consumer Services, Division of Plant Industry, Gainesville, FL. These data started at the weevil’s first known occurrence in September 2000 and proceeded through January

2015. The geographic coordinates of each collection location were imported into DIVA-

GIS 7.5 (Hijmans et al. 2012). Bioclimatic variables (average for years 1970-2000) were 56 downloaded from WorldClim 2.0 (Fick and Hijmans 2017) containing a set of global climate layers with a spatial resolution of approximately 1 km2. Temperature and precipitation parameters were selected using the BIOCLIM tool (Busby 1991) within this program. The geographic coordinates (data points) were used to identify the potential distribution in Florida based on mean annual temperature and precipitation. Utilizing the same data points, potential distribution was projected on a North American map by using the BIOCLIM tool and selecting isothermality and annual precipitation parameters.

In addition to the statistical model projections, a model was developed with the biological data from the cold tolerance experiment. This model looked at areas in North

America where there were at least four consecutive days with average daily air temperatures at or below 0° C (LT90). Maps were imported to ArcGis 10.2, and an isothermal line was created that predicts the northern limit suitable for M. undecimpustulatus undatus establishment.

Results

Cold Tolerance of Adults. A significant negative effect of low temperature on M. undecimpustulatus undatus adult survival was observed for both 0° and -5° C treatments (P < 0.0001, Table 4-2). The trend of reduced survival with increasing exposure time to low temperature was evident in both experiments. Weevil survival was always 100% in the control treatments at constant 20° C and until the insects were exposed to the minimum temperatures in both experiments. After 2 d of exposure, survival was 40-73% (mean 60%) at 0° C and 13-20% (mean 18%) at -5° C. After 4 d of exposure, 7-13% (mean 11%) and 0-7% (mean 4%) of weevils survived at 0° and -5° C, respectively. No weevils survived 8 d of exposure to the minimum temperatures.

A logistic fit of weevil response to temperature by exposure time (days) for each of the three trials of each experiment showed no significant difference between logistic fits for cold exposures at 0° or -5° C. Therefore, data for each experiment were combined for calculating the LT50 and LT90. At 0° C, the LT50 was 2.3 d, and the LT90 was 4.0 d (Figure 4-3a). At -5° C, the LT50 was 1.0 d, and the LT90 was 2.3 d (Figure 4-

3b).

Sustained Cold Exposure (SCE) / Repeated Cold Exposure (RCE). Survival of weevils collected in summer was significantly reduced in the sustained cold exposure treatment compared to the control and repeated cold exposure treatments (F = 7.1, df =

5, 25, P < 0.001) (Figure 4-4). Survival was reduced at least 3.4-times in the sustained cold exposure treatment compared to all other treatments. There was no significant effect of sustained cold exposure or repeated cold exposure treatments on survival of weevils collected in winter (F = 2.6, df = 5, 29; P > 0.05) (Figure 4-4).

Leaf consumption by summer weevils was not significantly different among the control and repeated cold exposure treatments, whereas weevils under sustained cold exposure consumed significantly less than weevils in the repeated cold exposure III and repeated cold exposure IV treatments (F = 7.2; df = 5, 29; P < 0.0001) (Figure 4-5). Leaf consumption by weevils collected in winter and exposed to repeated cold exposure and sustained cold exposure treatments was significantly less than that of control weevils (F

= 15.1; df = 5, 29; P < 0.0001) (Figure 4-5). Leaf consumption by winter weevils decreased as the number of repeated cold exposure periods increased and was least in the sustained cold exposure treatment. Average leaf area consumed by winter collected weevils was greater than that consumed by summer collected weevils, being 4.0-times

58 greater in the control treatment (1.53 cm2 vs. 0.37 cm2, respectively) and 2.0-times greater in the sustained cold exposure treatment (0.3 cm2 vs. 0.15 cm2, respectively).

Niche Modeling. An isothermal line based on cold exposure survival data at

0° C predicts that climatic conditions are suitable for M. undecimpustulatus undatus in all of Florida and Louisiana, the southern portions of South Carolina, Georgia, Alabama,

Mississippi, Arkansas, Texas, New Mexico, Arizona, and Nevada, a majority of

California, and western Oregon (Figure 4-6). A niche model based on voucher collections data predicted the ability of M. undecimpustulatus undatus to survive in

South Carolina, Georgia, Alabama, Louisiana, and portions of southern North Carolina,

Mississippi, southeastern Texas, northern California, western Oregon and Washington, eastern Mexico on the Gulf Coast, and the Bahamas (Figure 4-6). The highest probability areas (red and orange) are in Florida, southern Georgia, northern California, western Oregon, along with eastern Mexico and the Bahamas. An area from the Florida panhandle westward along the Gulf Coast was defined as not suitable for this weevil in the niche model.

Discussion

Some weevils exposed to -5° C for 4 d survived, suggesting that M. undecimpustulatus undatus can acclimate to colder temperatures not experienced in their native range. Insects that demonstrate adaptations to cooler temperatures have evolved different physiological mechanisms to survive (Renault et al. 2004). However, adult M. undecimpustulatus undatus may have suffered sublethal damage or tradeoffs, such as have been observed in the development, reproduction, and tissues and cells of other insects (Marshall and Sinclair 2010). For example, Rueda and Axtell (1996) found that reproduction and development in an adult tropical beetle, Alphitobius diaperinus 59

Panzer (Coleoptera: Tenebrionidae), was inhibited at temperatures below 17° C. There could also be a possibility of tradeoffs based on the immediate thermal history of the beetles, thus compromising their tolerance to the other extremes such as winter collected weevils for tolerance of high summer temperatures and vice versa.

In natural environments in subtropical and temperate regions, insects are subjected to repeated cold exposures throughout their lifetime (Marshall and Sinclair

2012). The impact of temperature fluctuations on adult M. undecimpustulatus undatus collected in summer and winter was observed in their mortality and consumption rates, both of which were generally greater in winter-collected weevils exposed to repeated cold exposure and sustained cold exposure treatments. Renault et al. (2004) surmised returning to higher temperatures may provide for the repair of injuries sustained when A. diaperinus experienced 5° and 0° C for 22 h/day and then returned to 20° C for 2 h.

They also found that the beetles exposed to repeated cold exposure had greatly improved survival over beetles kept continuously at lower temperatures. Marshall and

Sinclair (2012) concluded that D. melanogaster’s response to repeated cold exposure versus sustained cold exposure always resulted in higher survival, which could be attributed to the cumulative effect of the physiological impact of repeated cold and warm cycles. Anderson et al. (2017) found improved chill coma recovery, cellular survival, and cold tolerance in the migratory locust, Locusta migratoria Linnaeus, after brief periods of cold exposure.

Seasonality appears to be a factor in cold tolerance by adult M. undecimpustulatus undatus. Winter weevils’ survival was two times greater than that of summer weevils in the sustained cold exposure treatments. Similarly, Russell et al.

(2017) found survival of salvinia weevils, Cyrtobagous salviniae Calder and Sands, from

South America was greater and chill coma recovery was faster in weevils that experienced longer durations of cold temperatures before being exposed to 0° C treatments. Russell (2017) found similar results with summer and winter-collected salvinia weevils from Louisiana; specifically, survival increased 1.8 times and chill coma recovery was 1.5 times faster in the winter-collected weevils compared to those collected in the summer.

Leaf consumption after cold exposure was higher for M. undecimpustulatus undatus weevils harvested in winter. Winter weevils experienced natural cooling events preceding collection, when temperatures dipped to about 4° C over a period of weeks.

This may have provided natural repeated cold exposure effects, thereby increasing their ability to withstand the cold exposures experienced during the experiment. This suggests that M. undecimpustulatus undatus has the potential to expand its distribution northward of subtropical southern Florida and survive the modulating winter temperatures in the southern and western regions of temperate North America.

Identifying climatic factors correlated to the native distribution of M. undecimpustulatus undatus are important in projecting its spread within the United

States. Sri Lanka has an annual mean temperature of 27° C and annual rainfall of 1,337 mm compared to 22° C and 1,314 mm in southern Florida (Merkel 2016, Osborn 2017).

The niche model showed an area from the Florida Panhandle westward along the Gulf

Coast as not suitable for this weevil. This area receives more than 1,524-1,778 mm of annual rainfall (Daly 2017) and has predominantly aquic or udic soil moisture regimes

(U. S. Department of Agriculture 2017). This leads to the suggestion that it is too wet for

61 weevil larvae to survive along the Gulf Coast from the western Florida Panhandle to

Texas.

The spatial coincidence of the isothermal line based on cold exposure and the climate niche model increases assurance in the prediction of where M. undecimpustulatus undatus has the possibility of surviving. The niche model and the isothermal line provide projections that M. undecimpustulatus undatus could spread north and west within the United States. The most likely method of movement is through accidental transport by commerce, specifically potted plant materials (O’Brien 2006) grown in southern Florida and distributed to other areas below the isothermal line for survival. Adult weevils tend to hide on the undersides of leaves out of direct view and drop to the soil if disturbed. Weevil eggs and larvae may be simply carried to other destinations in the potting media. Shippers of potted plant materials should first inspect foliage for chewing damage from adults and treat infested plant material and soil appropriately before distribution per Florida Statue 581.161 (FLA. STAT. § 2018).

California has intercepted this weevil six times between 2000 and 2017, typically on nursery stock from Florida (Iqbal 2017). The California Department of Food and

Agriculture currently has a proposal to rank M. undecimpustulatus undatus as an “A”, which means there is the expectation of significant economic and environmental impacts if it were to establish within the state (Iqbal 2017).

a b

. Figure 4-1. Myllocerus undecimpustulatus undatus adults in (a) plastic containers with Chrysobalanus icaco leaves and a Petri dish with moistened paper towel, b) closed plastic containers with screened windows. Photographs by Anita Neal.

Table 4-1. Details of sustained cold exposure (SCE) and repeated cold exposure (RCE) experiments on Myllocerus undecimpustulatus undatus adults collected from the field in August 2014 and January 2015. Five containers with ten weevils each were used for each treatment.

Treatment Temperature Frequency Duration of No. of of exposure/ of exposures/ exposures recovery (° C) exposure recovery time

Control 20 / 20 Continual 8 days N/A

SCE -5 / 20 Daily for 1 day 10 h/14 h + 4 days 1

RCE I -5 / 20 Daily for 1 day 2 h/22 h + 4 days 1

RCE II -5 / 20 Daily for 2 days 2 h/22 h + 4 days 2

RCE III -5 / 20 Daily for 3 days 2 h/22 h + 4 days 3

RCE IV -5 / 20 Daily for 4 days 2 h/22 h + 4 days 4

Table 4-2. Mean percentage survival of field-collected Myllocerus undecimpustulatus undatus adults following exposure to 0° C or -5° C for 0.5, 1, 2, 4, or 8 d during experiments conducted in three consecutive months. Means with the same letters within each column are not significantly different (ANOVA, LSD, P < 0.05).

June July August Overall mean

Exposure Temperature Temperature Temperature Temperature Time (d) 0° -5° 0° -5° 0° -5° 0° -5° 0 100a 100a 100a 100a 100a 100a 100a 100a 0.5 86.7ab 60.0b 80.0ab 66.7b 93.3a 73.3b 82.2b 66.7b 1 80.0ab 33.3c 73.3b 33.3c 80.0ab 26.7c 82.2b 31.1c 2 73.3b 20.0d 66.7b 13.3d 40.0bc 20.0cd 60.0c 17.8d 4 6.7c 6.7e 13.3c 0.0d 13.3c 6.7de 11.1d 4.4e 8 0.0c 0.0e 0.0c 0.0d 0.0c 0.0e 0.0d 0.0e

LT50

LT90

LT50

LT90

Figure 4-2. (a) Survival response models of adult Myllocerus undecimpustulatus undatus over time at a) 0° C and b) -5° C. The green line represents observed data. LT50 and LT90 are the estimated number of days to kill 50% and 90%, respectively, of the population at a given temperature; arrows extending each side of the LT points indicate the mean ± SE.

Summer 5 Winter b

2 a a a a Mean Mean Number of a 1 a a a

Dead Adults/Container Dead a a a 0 Control RCE II RCE III RCE IV RCE V SCE Treatment

Figure 4-3. Mean (+ SE) number of dead summer- and winter-collected adult Myllocerus undecimpustulatus undatus after repeated cold exposure (RCE) and sustained cold exposures (SCE). Weevils in the RCE treatments were held for one (I), two (II), three (III), or four (IV) 2-h exposures. Columns for summer-collected weevils and columns for winter-collected weevils with different letters are significantly different (Tukey's HSD test, P < 0.05).

1.8 a Summer 1.6 Winter 1.4 ab 1.2 bc bc 1.0 0.8

) per Adult ) per a cd 2 0.6 ab ab a 0.4 ab d (cm b

0.2 Leaf Area Area Consumed Leaf 0.0 Control RCE I RCE II RCE III RCE IV SCE Treatment

Figure 4-4. Mean (+ SE) leaf area consumed by summer- and winter-collected adult Myllocerus undecimpustulatus undatus after repeated cold exposure (RCE) and sustained cold exposures (SCE). Weevils in the RCE treatments were held for one (I), two (II), three (III), or four (IV) 2-h exposures. Columns for summer-collected weevils and columns for winter-collected weevils with different letters are significantly different (Tukey's HSD test, P < 0.05).

Isothermal line

Figure 4-5. Isothermal line (dashed) predicting the northern limits of Myllocerus undecimpustulatus undatus distribution based on daily air temperatures of 0° C for at least 4 d (LT90). Colored areas in North America where M. undecimpustulatus undatus is predicted to occur by DIVA-GIS BIOCLIM utilizing two climate variables, isothermal temperature, and annual mean precipitation. Color is indicative of the probability of occurrence from low (dark green) to high (red). Inset: Peninsular Florida with blue dots representing collection locations by Florida Department of Agriculture and Consumer Services, Division of Plant Industry.

CHAPTER 5 MORTALITY AND FEEDING BEHAVIOR OF ADULT Myllocerus undecimpustulatus undatus MARSHALL (COLEOPTERA: CURCULIONIDAE) EXPOSED TO BIOPESTICIDES IN LABORATORY ASSAYS

Management of Myllocerus undecimpustulatus undatus Marshall through the use of synthetic insecticides has proven to be unsustainable (Arévalo and Stansly 2009).

These pesticides are the traditional method of controlling insects that attack crops and ornamental plants. However, concern for environmental impacts, pesticide resistance, and the desire for effective and sustainable pest control point to the necessity of an

Integrated Pest Management (IPM) approach (Leibee and Capinera 1995).

Implementing IPM through combining different strategies together to overcome the limitations of specific practices should include the use of biopesticides (Chandler et al.

1998).

Biopesticides are compounds containing living organisms or substances derived from natural materials like animals, plants, fungi, bacteria, or certain minerals (Sporleder and Lacey 2013). Biopesticides are categorized according to their active ingredient into one of three types: microorganisms; biochemicals; and semiochemicals (Chandler et al.

1998). Microbial pesticides, e.g., entomopathogenic fungi, are found naturally occurring in soils worldwide, and they may infect a wide range of insect hosts (Hajek and Leger

1994, Meyling and Eilenberg 2007, Senthil-Nathan 2015). The biochemical pesticides that discourage herbivores from feeding are made from plants that produce secondary metabolites like pyrethrins produced by Chrysanthemum cinerariaefolium Vis.

(pyrethrum) and phenolic acids extracted from the seeds of Azadirachta indica A. Juss

(neem) (Chandler et al. 1998, Senthil-Nathan 2015). Another biochemical pesticide is spinosad, a mixture of natural metabolites (spinosads A and D) produced under aerial

70 fermentation conditions by a soil actinomycete, Saccharopolyspora spinosa (Mertz and

Yao 1990, Chandler et al. 1998).

Shanthipriya and Misra (2007) evaluated nine pesticide treatments against adult

M. maculosus on field grown okra. The number of adult weevils per plant were recorded prior to treatment and five and ten days after treatment application. Padan

(neonicotinoid), spinosad, Daman (Beauveria bassiana (Balsamo) Vuillemin), and

Ozoneem (A. indica) reduced weevil numbers at day 5 by 91%, 48%, 30%, and 48%, respectively. At day 10, B. bassiana showed a further reduction in the number of weevils per plant to 62%.

To date, studies on control strategies for M. undecimpustulatus undatus are limited. The purpose of this study was to evaluate survival and mortality of adult M. undecimpustulatus undatus exposed to biopesticides in laboratory assays. My null hypothesis is that biopesticides have an equal effect on M. undecimpustulatus undatus mortality and feeding behavior.

Materials and Methods

Petri Dish Plant-based Bioassay.

A Petri dish bioassay system was designed to evaluate seven commercial products (treatments) on cocoplum leaves for toxicity to the adults of M. undecimpustulatus undatus. The bioassay system was modified from one developed by

McKenzie et al. (2001). Each bioassay cage consisted of a conical tube inserted into a vented Petri dish into which plant and insect material were added and the dish sealed with Parafilm® (Figure 5-1).

Polystyrene conical centrifuge tubes (15 ml, 17 × 120 mm) held the stem of the plant material. A hole was drilled near the top to accommodate adding water as needed

71 throughout the duration of the bioassay. A small piece of 1 cm diameter wooden dowel was inserted into the conical tube to prevent breaking or cracking the tube during drilling. A small drill bit was used to make a starter hole into the conical tube, and a larger drill bit was used to bore out the hole to accommodate a squirt bottle tip.

Polystyrene Petri dishes (150 × 20 mm) were used to contain the leaves and insects during the bioassay. A hole was drilled through the side of a closed Petri dish using a step drill bit (the exact size of the conical tube). Another hole, approximately 120 mm in diameter, was made using a hole-saw bit through the center of the small plate of the

Petri dish (for ventilation). A piece of thrips-proof screen slightly larger than the hole was attached to the Petri dish by a clear, fast-curing solvent cement for joining acrylic (Weld-

On Acrylic Cement-IPS Corporation, www.ipscorp.com). This provided ventilation for the plant and insects contained within the cage during the bioassay.

Plant and Insect Preparation.

Cocoplum leaves on stems were used as a food source for the experiments as the leaves fit inside the Petri dish without needing to trim them. Twenty-four cocoplum branches cut to 25 cm in length each with 4-6 leaves were washed and kept hydrated until use. Two hundred forty field-collected M. undecimpustulatus undatus adults from the laboratory colony (described in Chapter 3) were removed from the colony three days prior to use and placed into a Bug Dorm cage with a water source but no food.

Fungal Formulations.

Three commercially available entomopathogenic fungal formulations were tested:

BotaniGard® ES (a.i. spores of B. bassiana strain GHA, inert ingredients 88.7%)

(Laverlam International Corporation, Butte, MT); PFR-97™ 20% WDG (a.i. Isaria fumosorosea Apopka strain 97 20%, inert ingredients 80%) in the form of desiccation-

72 tolerant granules of blastospores (Certis USA, Columbia, MD); and Met52® EC (a.i. spores of M. anisopliae strain F52, inert ingredients 89%) (Novozymes Biologicals Inc.,

Salem, VA). Fungal solutions were prepared by mixing the fungal products with distilled water in a beaker with a magnetic agitator. For PFR-97, 0.66 g were dissolved in 300 ml of distilled water. The beaker was placed on a stir plate for 30 min, and the solution then allowed to settle for 20 min. This allowed the inert ingredient to precipitate out, thereby leaving the supernatant containing the blastospores. For BotaniGard, 750 µl of the product were dissolved in 300 ml of distilled water and mixed on a stir plate for 10 min.

For Met52, 960 µl were dissolved in 300 ml of distilled water and mixed on a stir plate for 10 min. The concentrations of all fungal solutions were determined by counting the number of spores per ml using a disposable plastic Neubauer hemocytometer, C-

Chip™ DHC-N012 (Incyto Co., Ltd., Chungnam-do, Korea). Each solution was then appropriately adjusted to 6.0 x 107 blastosores/ml for PFR 97, 2.0 x 1010 conidia/ml for

BotaniGard, and 5.0 x 1010 conidia/ml for MET52 by adding either distilled water or product according to the formula in Avery et al. (2018), remixing and recounting spore concentrations. The viability of the blastospores and conidia was assessed by spreading 100 μl of each fungal solution onto potato dextrose agar in a Petri dish. The

Petri dishes were sealed with Parafilm® and incubated at constant temperature 25ºC for

7 d. Colony-forming units (CFU) were counted to check the quality of the fungal solutions by comparing the number of CFU obtained with the number of CFU specified for each formulation.

Biochemical Formulations.

Three products approved for organic production by the Organic Materials Review

Institute (OMRI) were tested: AzaMax™ (a.i. azadirachtin, inert ingredients 98.8%) 73

(Parry America, Inc. Sacramento, CA); Entrust® (a.i. spinosad (a mixture of spinosad A and spinosad D, inert ingredients 20%) (Dow AgroSciences L.L.C. Indianapolis, IN);

® PyGanic EC 1.4II (a.i. pyrethrins, inert ingredients 98.6%) (McLaughlin Gormley King

Co. Minneapolis, MN). Solutions were prepared by adding product amounts (AzaMax,

90 µL (equivalent rate 5 ml / L); Entrust, 0.26 g (equivalent rate 240 g / L); PyGanic, 480

µL (equivalent rate 19.8 ml / L) to 300 ml of distilled water in a beaker with a magnetic agitator and mixing on a stir plate for 10 min.

A conventional insecticide product, Sevin® SL (a.i. carbaryl, inert ingredients

57%) (Bayer Environmental Science Research Triangle Park, NC), was chosen for its broad-spectrum activity and because it is a popular choice for pest control by growers and homeowners. This pesticide was prepared with 90 µl of product mixed with 300 ml of distilled water (equivalent rate 2.5 ml / L) in a beaker with a magnetic agitator placed on a stir plate for 10 min. Distilled water only was the control.

Treatment Applications.

The centrifuge tubes were filled with approximately 35 ml of water 5 ml below the watering hole and were covered with a square (2.5 × 2.5 cm) of Parafilm that was secured to the sides of each tube to prevent insects from crawling in and drowning and to also provide support to the plant stem. Plant stems were inserted through the

Parafilm, as far as possible, keeping only foliage above. Another square of Parafilm was added around the stem and secured to the tube.

All treatments were applied using pressurizable 180-ml Nalgene® aerosol spray bottles (Nalgene Nunc International, Rochester, NY) and fine mist nozzles to deliver consistent spray coverage. Each bottle was labeled with one of eight treatments and filled with 80 ml of the corresponding liquid. The bottle was pressurized with eight to ten 74 strokes of the pump mechanism prior to each application. Each treatment was sprayed to the point of runoff on both leaf surfaces of cocoplum leaves, with three tubes per treatment. After treatment application, the tubes were set in a wire tube rack to air dry.

The tube was then placed into a hole in the lip of the bottom half of a Petri dish, ten weevils were added, and the Petri dish was immediately closed with the top half and secured with a long strip (10 × 2.5 cm) of Parafilm (Figure 5-1a). Bioassay cages were individually covered with a liter-size (17.7 × 18.8 cm) plastic bag (Ziploc® S. C. Johnson

& Son, Inc., Racine, WI) to increase humidity around the cage and prevent entomopathogenic fungal treatments from spreading among one another. Each bag was labeled with treatment and replicate number (Figure 5-1b). Six cages were randomized

(www.randomizer.org) in one of four Styrofoam racks. Racks were placed in a controlled environmental chamber (Percival Scientific, Inc., Perry, IA) at 25° C ± 1° with 20% RH and a 14 h photoperiod for the duration of the experiment (Figure 5-1c).

Weevils were checked daily for mortality for 15 d or until water only control weevils had eliminated their food source. Dead insects were surfaced sterilized and then placed in Petri dishes (100 × 15 mm) lined with No. 4 filter paper moistened with

800 µL of distilled water and observed daily until mycosis of the fungal pathogen was detected (7-14 days). Percentage mycosis was determined from the number of cadavers showing infection. Percentage leaf damage was estimated on day 15 using a rating scale of 1 = 0 - 10%, 2 = 11 - 25%, 3 = 26 - 50%, 4 = 51 - 75%, 5 = >76% (Figure

5-2) that was modified after Maletta et al. (2004).

This experiment was repeated four times (each treatment with 3 replicates) for a total of five trials. Trials were conducted in June, July, August, and October 2014.

Statistical Analysis.

Data on survival of M. undecimpustulatus undatus from each trial with entomopathogenic fungi and insecticide was evaluated. The median survival time in days (ST50) was compared for each trial using Kaplan-Meier survival analysis (α = 0.05) followed by a log rank test (SAS Institute, Inc. 2017, JMP®PRO 13.1, Cary, NC). Data from the five trials were compared to determine significance among trials using

Duncan’s Multiple Range Test. The combined mean survival time in days was subjected to ANOVA, and treatment means were separated and compared using Tukey’s HSD test (α = 0.05). To determine mortality, combined data comparing treatments were arcsine transformed and subjected to ANOVA; treatment means were separated and compared using Tukey’s HSD test (α = 0.05). Percentage leaf damage was subjected to ANOVA and treatment means were separated and compared using Tukey’s HSD test

(α = 0.05). All tests were performed using PROC GLM in SAS v. 9.2 (SAS Institute

2009).

Results

Survival rates varied significantly among treatments in each trial: Trial 1: log rank

X2 = 33.2, P < 0.0001, df = 7; Trial 2: log rank X2 = 76.6, P < 0.0001, df = 7; Trial 3: log rank X2 = 192.9, P < 0.0001, df = 7; Trial 4: log rank X2 = 89.9, P < 0.0001, df = 7; and

Trial 5: log rank X2 = 157.3, P < 0.0001, df = 7 (Figure 5-3). Trials were compared and not found to be significantly different (F = 1.64; df = 4, 1015: P = 0.169); therefore, the results from all trials were pooled for analysis.

Kaplan-Meier analysis (censored at day 15) revealed that the ST50 was significantly longer in the control, PFR-97, Met52, and PyGanic treatments (each at 14 days) compared to AzaMax and Sevin (13 days), BotaniGard (12 days), and Entrust (9

76 days). The ST50 for BotaniGard and Entrust were significantly shorter than the ST50 for

AzaMax and Sevin, and the ST50 for Entrust was significantly shorter than that for

BotaniGard (F = 134.21; df = 7, 1043; P < 0.0001) (Figure 5-4).

Adult weevils had significantly greater mortality 15 d after leaves were treated with Entrust and BotaniGard compared to the other treatments (F = 28.39; df = 7, 98; P

< 0.0001) (Figure 5-5). The mortality rate with the Entrust and BotaniGard treatments was about two-fold higher than with the other product treatments.

Dead weevils exposed to BotaniGard exhibited significant mycelial growth emerging from intersegmental regions (Figures 5-6a and 5-6b). Dead weevils collected from BotaniGard treatments typically had mycelial growth prior to surface sterilization.

Dead weevils from the other fungal treatments failed to show any mycelial growth prior to surface sterilization. In the BotaniGard treatments, mycosis averaged 90% ± 0.15 across the five trials. Dead weevils in the PFR 97 and Met 52 treatments averaged 8%

± 0.04 and 5% ± 0.03 mycosis, respectively, across the five trials. No mycosis in dead weevils was seen in the non-fungal treatments, thus indicating no cross-contamination.

Significant differences in leaf consumption rates were recorded among treatments (F = 124.71; df = 7, 9; P < 0.0001) (Figure 5-7). The highest mean damage rating occurred in the control (4.9), followed in order by BotaniGard (3.3), PFR 97 (2.7),

Met 52 (2.5), Sevin (1.6), AzaMax (1.4), Entrust (1.0), and PyGanic (1.0) treatments.

The control weevils ate significantly more than weevils exposed to BotaniGard treated leaves, which in turn ate significantly more than weevils exposed to leaves treated with

PFR-97 and Met52. Weevils on leaves treated with Entrust, PyGanic, AzaMax, and

Sevin consumed significantly less than with other treatments.

Discussion

This is the first laboratory study to identify effective biochemical pesticides for managing adult M. undecimpustulatus undatus. Of the seven pesticides tested, Entrust consistently performed well in killing M. undecimpustulatus undatus adults and reducing plant damage (Figures 5-5 and 5-7). The mode of action of spinosad, the active ingredient in Entrust, is by contact and ingestion (Eger and Lindenberg 1998). This attests to the high mortality rate (83%) and low plant damage rate (1.0). Balusu and

Fadamiro (2012) obtained similar results with Entrust treatments on the yellowmargined leaf beetle, Microtheca ochroloma Stål (Coleoptera: Chrysomelidae). Entrust consistently achieved high efficacy among eight botanical and microbial insecticides tested compared to the control. This pesticide also reduced plant damage (1 = <10%

Plant Damage Rating Index) by the yellowmargined leaf beetle.

My bioassay is the first investigation of a spinosad product tested against M. undecimpustulatus undatus. Entrust is labeled for chrysomelids but not for curculionids.

The rate used in my bioassay, 27 g ai/ha (.025 lb ai/A) is one fifth the highest recommended rate of 140 g ai/ha (0.125 lb ai/A). This amount was chosen to determine efficacy of a lower recommended rate. McLeod and Rashid (2010) reported 100% mortality of eggplant flea beetles, Epitrix fuscula Crotch, at the highest recommended rate of Entrust. Reduced feeding started the first day. Bažok et al. (2016), using 72 g ai/ha (0.07 lb ai/A) compared contact and ingestion activity of spinosad against adult sugar beet weevils, Bothynoderes punctiventris Germar. Five days after treatment, mortality rate was three times higher through ingestion (95%) than through contact

(39%).

My bioassay is the first investigation of entomopathogenic fungi tested against M. undecimpustulatus undatus. Of the three entomopathogenic fungi tested, BotaniGard consistently resulted in infection. BotaniGard was twice as effective as PFR-97 and Met

52. Observation of fungal outgrowth on insect cadavers is used to verify death by fungal infection (Ondiaka et al. 2008). Mycosis by B. bassiana was observed prior to and after surface sterilization. The effectiveness of B. bassiana has been described with other insects. For example, the red palm weevil, Rhychophorus ferrugineus Olivier

(Coleoptera: Curculionidae), showed signs of infection 3-5 days after inoculation

(Güerri-Agulló et al. 2010). The pine weevil, Hylobius abietis Linnaeus (Coleoptera:

Curculionidae), had 100% mortality 6-7 days after exposure (Martin et al. 2015). The

Japanese beetle, Popillia japonica Newman (Coleoptera: Scarabaeidae), had 79% mortality 9 days after exposure (Lacey et al. 1994). And the sweet potato weevil, Cylas formicarius Fabricius (Coleoptera: Brentidae), had 93% mortality 7-8 days after exposure (Leng and Reddy 2012).

The poor efficacy of PFR-97 and Met 52 could be related to various factors. One possible cause contributing to the lack of infectivity is an insect’s first line of defense, the cuticle (Ortiz-Urquiza and Keyhani 2013). BotaniGard and Met 52 contain hydrophobic conidia, whereas I. fumosorosea was applied as blastospores, which are hydrophilic in nature (Dunlap et al. 2005). Entomopathogenic fungi spores are suspended when mixed with water, however, formulations of them in products are dissimilar. PFR-97 is a dry desiccation-tolerant blastospore powder with no surfactants or emulsifiers. BotaniGard and Met 52 contain a mixture of surfactants and emulsifiers suspended in oil (Avery et al. 2018). A number of studies indicate that conidia

79 suspended in oil have superior infectivity to a pure aqueous application (e.g., Bateman et al. 1993, Prior et al. 1988).

Another possibility for the low efficacy of PFR-97 and Met 52 is the plant cuticle.

Cocoplum is a drought-, salt-, and wind-tolerant plant species (Brown and Frank 2018) usually identified as having a thick waxy cuticular layer (Barrick 1978). The plant cuticle is a hydrophobic layer composed of cutin and waxes serving as a primary barrier to water loss and interactions with organisms and the environment (Yeats and Rose

2013). Plant surface chemistry, i.e., plant exudates, affecting conidia directly, herbivore- induced plant volatiles affecting sporulation or germination, and plant cuticle altering spore persistence (Cory and Ericsson 2010) are possibilities for reduction in infectivity of both pathogens.

Genomic data revealed that even closely related fungal pathogens, i.e., B. bassiana and M. anisopliae, have different molecular mechanisms that mediate virulence (Xiao et al. 2012). Beauveria bassiana has many more bacteria-like toxins and

Cry-like delta endotoxins than M. anisopliae and other fungi, suggesting the possibility of greater oral toxicity than other fungi (Xiao et al. 2012). There is evidence that B. bassiana may infect insects per os, especially in insects with chewing mouthparts (Feng et al. 1994). Recent studies are investigating B. bassiana as a fungal endophyte in plant defense against the coffee berry borer, Hypothenemus hamperi Ferrari (Coleoptera:

Curculionidae) (Vega et al. 2008), and the banana weevil, Cosmopolities sordidus

Germar (Coleoptera: Curculionidae) (Akello et al. 2008). Further investigation is needed to determine if the higher incidence of B. bassiana infection is due to entry per os.

PyGanic and AzaMax were less effective than Entrust and BotaniGard, both producing less than 50% mortality. Although both products are contact insecticides,

AzaMax also controls by ingestion (Barry et al. 2005). The repellency action by PyGanic and AzaMax might explain the lower feeding damage ratings of 1.0 and 1.4, respectively. Other reports recognized the reduced effectiveness of azadirachtin-based insecticides with adult insects, e.g., the Colorado potato beetle, Leptinotarsa decemlineata Say (Coleoptera: Chrysomelidae) (Trdan et al. 2007) and grain beetles,

Sitophilus oryzae Linnaeus (Coleoptera: Curculionidae) and Tribolium confusum

Jacquelin du Val (Coleoptera: Tenebrionidae) (Kavallieratos et al. 2007). PyGanic was recognized as ineffective against adult apple flea weevils, Orchestes pallicornis Day

(Coleoptera: Curculionidae) (Neilson et al. 2012) and adult alfalfa weevils, Hypera postica (Gyllenhal) (Coleoptera: Curculionidae) (Long et al. 2009).

In conclusion, the results of the laboratory assays determined a high mortality caused by Entrust and BotaniGard, suggesting that M. undecimpustulatus undatus adults might be well managed using these two commercially available biopesticides.

However, it is important to point out the effectiveness of the products was tested under controlled conditions in the laboratory. Therefore, additional testing should be carried out in the field where the environmental conditions vary and the products may yield different results.

a b

Figure 5-1. Petri dish cages (a) modified from McKenzie et al. (2001), (b) with ten adult weevils on Chrysobalanus icaco and enclosed within a plastic bag, (c) arranged in a randomized block design in an environmentally controlled chamber at 25° C ± 1°. Photographs by Anita Neal.

PDRI - 1 PDRI - 2 PDRI - 3

PDRI - 4 PDRI - 5

Figure 5-2. Plant Damage Rating Index (PDRI) examples of adult Myllocerus undecimpustulatus undatus feeding on treated Chrysobalanus icaco leaves. Percentage leaf damage was estimated at day 15 using a rating scale of 1 = 0 - 10%, 2 = 11 - 25%, 3 = 26 - 50%, 4 = 51 - 75%, 5 = >76% for trials 1 through 5. Photographs by Anita Neal.

Trial 1: 6/6/2014 Trial 2: 7/25/2014

Trial 3: 8/22/2014 Trial 4: 10/9/2014

Trial 5: 10/30/2014 Combined Trials

Figure 5-3. Comparison of Myllocerus undecimpustulatus undatus survival curves for each treatment trial and combined treatment trials using Kaplan-Meier survival analysis (α = 0.05). Treatments included PFR-97™ 20% WDG (PFR), BotaniGard® ES (BG), Met 52® EC (MET), Entrust® SC (ET), PyGanic® EC ™ ® 1.4II (PY), Aza-Max (AZ), Sevin SL (SV), and water (C).

16 a a a a 14 b b c 12 d 10 8 6 4

2 Mean Survival Time (Days) Time Survival Mean 0 PFR BG MET ET PY AZ SV C

Treatment

Figure 5-4. Comparison of mean survival time in days of adult Myllocerus undecimpustulatus undatus on treated Chrysobalanus icaco leaves. Treatments included PFR-97™ 20% WDG (PFR), BotaniGard® ES (BG), Met 52® EC (MET), Entrust® SC (ET), PyGanic® EC 1.4II (PY), Aza-Max™ (AZ), Sevin® SL (SV), and water (C). Bars are means ± SEM of five trials (three replicates per treatment per trial). Treatments not followed by the same letter above the bars are significantly different (P < 0.0001 Tukey – Kramer HSD test).

100 a 90 a 80 70 60 b 50 bcd 40 bcd

Mortality (%) Mortality bcd 30 cd d 20 10 0 PFR BG MT ET PY AZ SV C

Treatment

Figure 5-5. Mortality of adult Myllocerus undecimpustulatus undatus 15 d after treatment. Treatments included PFR-97™ 20% WDG (PFR), BotaniGard® ES ® ® ® (BG), Met 52 EC (MET), Entrust SC (ET), PyGanic EC 1.4II (PY), Aza- Max™ (AZ), Sevin® SL (SV), and water (C). Bars are means ± SEM. Treatments not followed by the same letter above the bars are significantly different (P < 0.0001 Tukey – Kramer HSD test).

a b

Figure 5-6. Myllocerus undecimpustulatus undatus adults (a) mycosing after infection with Beauveria bassiana, (b) close up of B. bassiana conidia on the weevil’s tibia and tarsus. Photographs by Anita Neal.

6 a 5

4 b c 3

2 de d e e

SE) Plant Damage Rating Damage Plant SE) 1 ±

PFR BG MET ET PY AZ SV C Mean ( Mean

Treatment

Figure 5-7. Plant damage ratings of treated Chrysobalanus icaco leaves fed on by adult Myllocerus undecimpustulatus undatus 15 days after treatment application. Treatments included PFR-97™ 20% WDG (PFR), BotaniGard® ES (BG), Met ® ® ® ™ 52 EC (MET), Entrust SC (ET), PyGanic EC 1.4II (PY), Aza-Max (AZ), Sevin® SL (SV), and water (C). Data are for five combined trials. Treatments not followed by the same letter above the bars are significantly different (P < 0.0001 Tukey-Kramer HSD test).

CHAPTER 6 COMPARISON OF MORTALITY RATES AND FEEDING BEHAVIOR OF ADULT Myllocerus undecimpustulatus undatus MARSHALL (COLEOPTERA: CURCULIONIDAE) EXPOSED TO BIOPESTICIDES ON PEACH FOLIAGE IN A FIELD EXPERIMENT

The presence of Myllocerus undecimpustulatus undatus Marshall in Florida and its potential spread to other areas in the southern United States has led growers and homeowners to search for control measures that are effective and sustainable. Peach leaves, Prunus persica L. (Rosaceae), are one of the pest’s favored adult food sources

(George et al. 2015) (Figure 6-1). Growing peaches in Florida is a practical option for citrus growers who are looking to new crops when citrus greening pressure has made citrus production unprofitable. Moreover, the Florida crop fills a market gap between production from South America (ending in March) until other states like Georgia,

California, and South Carolina start harvesting in May (Olmstead et al. 2016). Annual peach production in Florida is about 4.5 million pounds valued at $6 million. The unique market window (April/May) allows Florida peaches to be marketed at a premium price

(Morgan and Olmstead 2013). Myllocerus undecimpustulatus undatus is a threat to peach and other fruit crop production in Florida and, as such, necessitates the development of a sustainable management strategy.

Applications of synthetic, broad-spectrum insecticides are the current primary method of control. Management with these insecticides can kill beneficial organisms and may lead to pesticide resistance, intensifying the search for viable alternative solutions (Chandler et al. 1998). Entomopathogenic fungi have a rich diversity and cosmopolitan distribution and are widely used as biological control agents against invertebrate pests in agriculture (Gul et al. 2014, Herrero et al. 2012, Meyling and

Eilenberg 2007). The most important entomopathogenic fungi commercially produced 89 are Beauveria bassiana, Metarhizium anisopliae, and Isaria fumosorosea (Gul et al.

2014). The effective use of entomopathogenic fungi requires consideration of high relative humidity, temperature, and sunlight along with an understanding that pest populations will not be destroyed but maintained below economic thresholds (Gul et al.

2014, Shah and Pell 2003).

Azadirachtin, a botanical biopesticide with modes of action as a repellant, antifeedant, and insect growth regulator, has been shown to be effective against

Coleoptera. Musabyimana et al. (2001) reported that adults of the banana root borer,

Cosmopolities sordidus Germar (Coleoptera: Curculionidae), were repelled and oviposition reduced on azadirachtin-treated banana corms. Azadirachtin applied to okra reduced populations of adult Myllocerus maculosus Desbrochers des Loges at day 5 by

48% (Shanthipriya and Misra 2007). Azadirachtin sprayed on jute, Corchorus olitorius

L., reduced the number of adult Myllocerus discolor by 50% on day 3 (Rahman and

Khan 2011).

To date, only one study on control strategies for M. undecimpustulatus undatus has been performed on crops grown in the field (Arévalo and Stansly 2009). The purpose of my study was to use sleeve cages to evaluate mortality rates and feeding behavior of adult M. undecimpustulatus undatus exposed to biopesticides on peach foliage in the field. My null hypothesis is that biopesticides have an equal effect on M. undecimpustulatus undatus mortality rates and feeding behavior.

Materials and Methods

Sleeve Cages.

Each sleeve cage consisted of a 2-L soda bottle approximately 30 mm tall with eight windows, 7.0 × 5.5 mm with a 2-mm strip between, cut out to allow air movement

90 and four windows were cut in both the bottom and top half of the bottle (Figure 6-2a).

The neck of the bottle was removed 6 mm from the top. Three 1.5 mm holes were drilled in the base of the bottle to provide drainage during rain events. Nylon screen (two

® Léggs Knee Highs, Hanesbrands, Inc. Winston-Salem, NC) were placed over the cage with a portion extended just beyond the bottle opening.

Plants and Insects.

Peach trees at the Florida Research Center for Agriculture Sustainability in Vero

Beach, FL (29° 36′ 56.42″ N, 82° 21′ 57.3″ W) were used because local growers identified peach as a popular host plant for M. undecimpustulatus undatus. Twenty-five trees were randomly selected from a row of 50 trees located in Row 16 E, south of 33rd

Street for Site 1, and 25 trees were randomly selected from a row of 50 trees in Row E, north of 33rd Street for Site 2 (Figure 6-3). Tree variety was ‘Tropic Beauty’, and trees were planted in 2011. No insecticides or fungicides were applied to the trees 10 years prior to or during Site 1 trials. Prior to Site 2, the fungicide Nexicor™ Xemium® (a.i. fluxapyroxad, pyraclostrobin, and propiconazole, BASF Corporation, Research Triangle

Park, NC) was applied at the label rate on April 4 and 19, 2017. The last application was made one month before the second trial of Site 2 began. On June 7, 2017, the fungicide

Pristine (a.i. pyraclostrobin and boscalid, BASF Corporation) was applied at the label rate to nearby (±18 m) citrus trees two weeks prior to the third trial of Site 2.

Adult M. undecimpustulatus undatus were collected from mature Australian pine,

Casuarina equisetifolia L. (Casuarinaceae), in Fort Pierce, FL (27° 29′ 15.43″ N, 80° 24′

33.79″ W) about 1 week prior to each trial. The weevils were maintained in mesh- screened Bug Dorms at 24° C ± 2°, 60% RH, and 14 h photoperiod. They were fed cocoplum leaves as described in Chapter 3. Three days prior to use, 125 adults were 91 removed from the laboratory colony and placed in a Bug Dorm where feeding was discontinued but a water source provided.

Fungal and Biochemical Formulations.

Three commercially available entomopathogenic fungi were tested: BotaniGard®

ES (a.i. spores of B. bassiana strain GHA, Laverlam International Corporation, Butte,

MT); PFR-97™ 20% WDG (a.i. I. fumosorosea Apopka strain 97 20% in the form of desiccation-tolerant granules of blastospores, Certis USA, Columbia, MD); and Met52®

EC (a.i. spores of M. anisopliae strain F52, Novozymes Biologicals Inc., Salem, VA).

The fungal solutions were prepared and their concentrations quantified as described in

Chapter five.

One Organic Materials Review Institute product was tested: AzaMax™ (a.i. azadirachtin, Parry America, Inc. Sacramento, CA). The AzaMaz solution was prepared as described in Chapter 5. The control treatment was distilled water only.

The three fungal products were chosen to assess their performance in the field after comparing their effectiveness in the laboratory. AzaMax was selected due to its repellent effect on feeding behavior (Pavela 2014).

Experimental Design.

A randomized design was used on both sites. The three trials for each site used the same 25 trees, but different branches were selected for each trial. Small branches with 6-8 leaves showing no evidence of herbivory were selected prior to treatment application. A laminated tag identifying treatment, tree number, and sleeved or unsleeved was attached with a twist tie to each of two branches per tree (Figure 6-2b).

The branch labeled sleeved had the sleeve cage placed over it after treatment application, whereas the branch labeled unsleeved remained uncaged. The purpose of

92 the uncaged branch was to assess the presence of resident entomopathogenic spores and any that might be acquired from spray drift.

Prior to treatment application, one leaf was removed from each tagged branch labeled unsleeved. The leaf was placed in a pre-labeled sandwich-size (16.5 × 14.9 cm), resealable plastic bag (Ziploc® S. C. Johnson & Son, Inc., Racine, WI). Bags were kept in the shade until transported to the laboratory where they were stored in a refrigerator for up to two weeks until processed (see below).

Treatments were applied in late afternoon one hour before dusk. A pressurizable

180-ml Nalgene® aerosol spray bottle (Nalge Nunc International, Rochester, NY) with a fine mist nozzle was used to deliver consistent spray coverage. Each bottle was labeled with one of the five treatments and filled with 80 ml of the corresponding solution. The bottle was pressurized with 8-10 strokes of the pump mechanism for each application.

Each treatment was sprayed to the point of runoff on both surfaces of the peach leaves on the sleeve-labeled branch and allowed to air dry. Spray applications were made in a manner to minimize drift to other parts of the tree. One sprayed leaf was then removed and placed in a pre-labeled sandwich-size plastic bag. Bags were kept in the shade until transported to the laboratory where they were stored in a refrigerator for up to two weeks until processed (see below). A sleeve cage was placed over the sprayed branch, five weevils were released inside the cage, and the cage was closed with a twist tie

(Figure 6-2c).

Applications for trials in Site 1 were made on March 18, May 5, and August 12,

2016. Applications for trials in Site 2 were made on October 19, 2016, May 18, and

June 22, 2017.

The sleeve cage was removed 15 days later, and dead and live weevils were counted. All weevils from each cage were placed in appropriately labeled sandwich-size plastic bags and taken to the laboratory. The dead weevils were surface-sterilized by dipping weevils successively in 65-70% ethanol, distilled water, 1% sodium hypochlorite, and distilled water, and then placed in Petri dishes (100 × 15 mm) lined with No. 4 filter paper moistened with 800 uL of distilled water. Weevils were observed daily for 7-10 days for mycosis (as described in Chapter 5). Live weevils were destroyed in the laboratory.

Herbivory on caged leaves was visually assessed 15 days post-application by using the Plant Damage Rating Index (Figure 6-4) described in Chapter 5. One leaf was removed from each uncaged and caged branch, placed in separate pre-labeled sandwich-size plastic bags, and stored in a refrigerator for up to two weeks until processed (see below).

Leaf Sample Processing.

Leaves were processed with the purpose of identifying the incidence of entomopathogenic fungal spores on caged and uncaged leaves pre- and post- application and at the end of the 15-day weevil-feeding period. A 1-cm brass cork borer was used to cut two leaf discs centered on the abaxial midrib (Figure 6-5a) from each leaf. The two discs were placed in a labeled Fisherbrand™ 15- ml polystyrene conical centrifuge tube (Fisher Scientific Co. LLC, Pittsburgh, PA). Three ml of 0.01% Triton X-

100 were added to each tube (Figure 6-5b), and vortexed for 15 seconds. One hundred

μl was removed from each tube and spread across potato dextrose agar in a Petri dish

(100 x 15 mm) labeled with the appropriate treatment, tree number, caged or uncaged, and collection date. Dishes were sealed with Parafilm. The 50 Petri dishes for each 94 collection date were randomized in a complete block design (five blocks, each block included all five treatments) on two trays (one for caged leaves and one for uncaged leaves) and placed in an environmental chamber at 22° C, 77% RH, and 14 h photoperiod for 7–10 days.

Statistical Analysis.

Data from the three trials within each site were compared to determine significance among trials using Duncan’s Multiple Range Test. Percentage mortality rates were arcsine transformed and subjected to ANOVA, and treatment means were compared using Tukey’s test (α = 0.05). Means of leaf damage ratings were subjected to ANOVA, and treatment means were compared using Tukey’s HSD test (α = 0.05). All tests were performed using PROC GLM as implemented in SAS v. 9.2 (SAS Institute

2009).

Results

Data from the three trials in each site were not significantly different among trials

(Site 1: F = 0.16; df = 2, 48: P = 0.85; Site 2: F = 0.47; df = 2, 48: P = 0.64). Since no interaction among trials and treatments was detected, results from trials in Site 1 were combined and results from trials for Site 2 were combined for analyses.

Weevil mortality rates in the sleeved BotaniGard treatment in both locations were significantly higher (Site 1: F = 44.35; df = 4, 56; P < 0.0001; Site 2: F = 25.09; df = 4,

66; P < 0.0001) compared to the other treatments (Figure 6-6). The mortality rate with the BotaniGard treatment was about fourfold higher than the other product treatments in both sites. The mortality rates of weevils with the PFR-97, Met52, and AzaMax treatments were not significantly different from the control.

There were significant differences in mean plant damage rating indices among treatments in both locations (Site 1: F = 42.58; df = 4, 56; P < 0.0001; Site 2: F = 65.47; df = 4, 56; P < 0.0001) (Figure 6-7). The mean damage rating index was greatest in the control treatment in both sites, followed in decreasing order by the BotaniGard, PFR-97,

Met52, and AzaMax treatments. Damage in the Met52 and AzaMax treatments in both sites was significantly less than in the BotaniGard, PFR-97, and control treatments, except there was no difference between the Met52 and PFR-97 treatments in site 2.

Dead weevils exposed to BotaniGard displayed substantial conidiophore growth emerging from intersegmental regions (Figure 6-8). These weevils typically exhibited mycosis prior to surface sterilization. In the BotaniGard treatment, weevils averaged

78% ± 0.03 mycosis across both sites. Dead weevils from the other treatments did not show mycosis prior to or after surface sterilization.

Comparing the number of CFUs from leaves sprayed with B. bassiana versus unsprayed leaves in all trials, it is apparent the spore population on leaves in this treatment was increased substantially by the application of the entomopathogenic fungus. Contamination by B. bassiana was found on many unsprayed leaves and also on leaves sprayed with other treatments (Tables 6-1 and 6-2). However, the spore population on leaves not treated with B. bassiana did not approach the population observed on B. bassiana-treated leaves. Spore populations on leaves sprayed with I. fumosorosea and M. anisopliae were found mainly on day 1 of the first trial in each site after the respective treatments were applied (Table 6-2).

The spore population of B. bassiana found on unsprayed leaves was usually greater on leaves collected on day 1 than on day 15, except for Site 1 trial 1, and Site 2

96 trials 1 and 2 (Table 6-1). The spore population on B. bassiana-treated leaves decreased from day 1 to day 15 in all trials. No rainfall events were recorded during the

15 days of trial 1 in both locations, and the spore reduction observed in these trials was not substantial (25 and 27%). Relatively little rainfall occurred during trial 2 of Site 2, and as in the trials that experienced no rainfall, spore reduction during the 15 days was not high (32%). However, in the three trials that experienced substantial rainfall (>60 mm per trial period), there was greater reduction (74-90%) in the B. bassiana spore population from day 1 to day 15 (Table 6-1).

Discussion

This is the first study to evaluate biochemical pesticides for managing adult M. undecimpustulatus undatus on peach leaves in the field. Of the four pesticides tested,

BotaniGard consistently performed well in killing M. undecimpustulatus undatus adults in both sites (Figure 6-6). BotaniGard was four times more efficacious than PFR-97 and

Met52. Similar effectiveness of B. bassiana in the field experiment has been described with other weevils. An application of B. bassiana sprayed on the trunks of pecan trees, against the weevil Curculio caryae Horn, caused significantly greater mortality (≥ 80%) than in the control (≤ 33%) (Shapiro-Ilan et al. 2009). Damage by the sweetpotato weevil, Cylas formicarius (Fabricius), was reduced 100% four days after a treatment combining B. bassiana and Metarhizium brunneum Petch (Reddy et al. 2014). Five species of the black vine weevil, Otiorhynchus spp., in a semi-field application of B. bassiana showed the first signs of infection within the first 14 days after treatment

(Hirsch and Reineke 2014).

Observation of fungal outgrowth on insect cadavers can be used to verify death by fungal infection (Ondiaka et al. 2008). Mycosis of B. bassiana was observed prior to 97 and after surface sterilization. This same observation was noted in the laboratory bioassay in Chapter 5.

The poor efficacy of PFR-97 and Met52 to affect adult survival could be related to various factors as noted in the discussion of the laboratory bioassays (Chapter 5). Isaria fumosorosea was applied as blastospores, and germination takes longer due to the hydrophobic layer of both the insect cuticle and plant leaf cuticle (Avery et al. 2018,

Yeats and Rose 2013). The insect cuticle’s aliphatic hydrocarbons and the secretion of antimicrobial compounds can have an important effect on fungal pathogenesis and might be considered an external immune defense (Hajek and St. Leger 1994, Ortiz-

Urquiza and Keyhani 2013).

The shaded environment provided by the sleeve cage may have protected the entomopathogenic fungi from ultraviolet (UV) radiation, the most detrimental environmental factor affecting entomopathogenic fungal spore viability (Fernandes et al.

2015). Most entomopathogenic fungi can survive only a few hours of direct exposure to solar UV radiation. Exposure to UV radiation can cause a delay of conidial germination, reducing fungal development (Fernandes et al. 2015). The sleeve cage may have also increased humidity around the leaves, another important requirement for pathogenesis

(Hajek and St. Leger 1994).

Rainfall events occurring on both sites may have contributed to a reduction in B. bassiana spore abundance on sprayed leaves (Table 6-1) by washing the spores off the leaves. Inglis et al. (2000) found that simulated rain removed 89-95% of B. bassiana spores from potato leaves. They also found that a substantial number of spores remained after rain exposure. In my study, it is apparent that some B. bassiana spores

98 remained on the peach leaves after rainfall events. Conidia deposited on the abaxial surface of the leaves may have been protected from the direct impact of rain. Although the rain likely removed a significant number of spores from the peach leaves, 10-75% of them seemed to have remained after experiencing 25-88 mm of rainfall (Table 6-1).

Beauveria bassiana, one of the best known entomopathogenic fungi, is commonly found in the soil of agricultural ecosystems worldwide (Meyling and Elienberg

2007, Xiao et al. 2012). Besides naturally occurring in soil, this fungus can also be found on leaf surfaces after being deposited by air or water (Ortiz-Urquiza and Keyhani

2013). The contamination by B. bassiana on peach leaves treated by other biopesticides or water only (Table 6-1) did not apparently affect weevil mortality or plant feeding damage. In both sites, there were significant differences in mortality rates between the B. bassiana treatments (69-73%) and the other treatments (≤20%) (Figure

6-6). Significant differences in mean plant damage rating in both sites were also detected between the B. bassiana treatments and nearly all other treatments (Figure 6-

7). Therefore, the relatively low level of incidence of B. bassiana spores on the leaves in the PFR-97, Met52, AzaMax, and control treatments did not cause sufficient mortality, if any, to mask differences among treatments.

AzaMax in both sites produced low mortality (16% and 19%), but it was more effective in suppressing plant damage (Figure 6-7). These results are similar to those obtained in the laboratory bioassay (Chapter 5), in which AzaMax caused 43% mortality but a mean PDRI of 1.4. Azadirachtin is well known for its ability to suppress herbivory by insects (Schmutterer 1990). For example, adults of the boll weevil, Anthonomus grandis grandis Boheman, exposed to azadirachtin in choice and no-choice assays

99 were continually repelled for >90 minutes and made significantly fewer feeding and oviposition punctures after 24 hours (Showler at al. 2004).

High mortality caused by the B. bassiana in BotaniGard and reduced feeding caused by AzaMax suggest that M. undecimpustulatus undatus may be well managed in the field using these two products in combination. Hernandez et al. (2012) confirmed a clear synergy combining B. bassiana and azadirachtin against the twospotted spider mite, Tetranychus urticae Koch (Acari: Tetranychidae). Combined treatments of B. bassiana and azadirachtin caused 100% mortality of C. formicarius 72-144 h after treatment; B. bassiana alone required 168-192 h to cause 100% mortality (Reddy et al.

2014). Combined treatments of spinosad and B. bassiana also caused 100% mortality of C. formicarius 48 h post-treatment (Reddy et al. 2014). The high mortality and reduced feeding of weevils exposed to spinosad-sprayed leaves in my laboratory bioassay (Chapter 5) may provide similar efficacy in the field. Products containing spinosad should be investigated as another potential environmentally friendly pesticide for the control of M. undecimpustulatus undatus adults in the field.

Finally, although this was a field experiment, the design assumed a more controlled environment because of the sleeves. The results could be considerably different if sleeved cages were not employed.

100

Figure 6-1. Peach leaves damaged by adult Myllocerus undecimpustulatus undatus. Photographs by Anita Neal.

101

a b c

Figure 6-2. Sleeve cage (a) a 2-L soda bottle with 8 square windows cut into it, (b) laminated tag identifying treatment and tree number, (c) cage with nylon sleeve enclosing sprayed plant material and secured with a tagged twist tie. Photographs by Anita Neal.

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Row 16 E Row E

Figure 6-3. Peach trees at two locations: Row 16E located on the south side and Row E located on the north side of 33rd Street at the Florida Research Center for Agriculture Sustainability in Vero Beach, FL. Photographs by Anita Neal.

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PDRI - 1 PDRI - 2 PDRI - 3

PDRI - 4 PDRI - 5

Figure 6-4. Plant Damage Rating Index (PRDI) examples of adult Myllocerus undecimpustulatus undatus feeding on treated peach leaves. Percentage leaf damage was estimated at day 15 using a rating scale of 1 = 0 - 10%, 2 = 11 - 25%, 3 = 26 - 50%, 4 = 51 - 75%, 5 = >76% for both sites for a total of six different treatment periods. Photographs by Anita Neal.

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a b

Figure 6-5. Peach leaf discs (a) taken from leaf samples, (b) in labeled centrifuge tube prior to vortexing. Photographs by Anita Neal.

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100 Site 1 90 80 a 70 60 50

40 Mortality (%) Mortality 30 b b 20 b b 10 0 PFR BG MET AZ C

Treatment

100 Site 2 90 80 a 70 60 50

Mortality (%) Mortality 40 30 b b 20 b b 10 0 PFR BG MET AZ C

Treatment

Figure 6-6. Adult Myllocerus undecimpustulatus undatus mortality rates 15 days after treatment applications. Treatments were PFR 97™ 20% WDG (PFR), BotaniGard® ES (BG), Met52® EC (MET), AzaMax™ (AZ), and water (C). Bars are means ± SEM of three trials combined within each site. Treatments not followed by the same letter above the bars in each graph are significantly different (P < 0.0001, Tukey – Kramer HSD test).

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Site 1

5 a

3 b b c 2 c

1 Mean plant damage rating damage plant Mean

0 PFR BG MET AZ Control

Treatment

Site 2 5 a

4 b 3 c 2 cd d

1 Mean plant damage plant rating Mean

0 PFR BG MET AZ Control

Treatment

Figure 6-7. Plant damage ratings of treated peach leaves exposed to adult Myllocerus undecimpustulatus undatus for 15 days after treatment application. Treatments are PFR-97™ 20% WDG (PFR), BotaniGard® ES (BG), Met52® EC (MET), AzaMax™ (AZ), and water (C). Bars are means ± SEM of three trials combined within each site. Treatments not followed by the same letter above the bars in each graph are significantly different (P < 0.0001, Tukey- Kramer HSD test).

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a b

Figure 6-8. Myllocerus undecimpustulatus undatus adult (a) exhibiting mycosis after infection with Beauveria bassiana, (b) with conidiophores emerging from the tarsus. Photographs by Anita Neal.

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Table 6-1. Mean ± SEM number of Beauveria bassiana CFUs/mm2 from sprayed and unsprayed peach leaves. Leaves were collected on the day of treatment application (d1) and 15 days later (d15). Bold numbers indicate presence of Beauveria bassiana on leaves not treated with the fungus. Sprayed is synonymous with treated sleeved or caged leaves. Unsprayed is synonymous with nontreated unsleeved or uncaged leaves. Collection BotaniGard® ES Met 52® EC PFR-97™ 20% AzaMax™ Control Rainfall (day) (CFU’s/mm2) (CFU’s/mm2) (CFU’s/mm2) (CFU’s/mm2) (CFU’s/mm2) (mm) Sprayed Unsprayed Sprayed Unsprayed Sprayed Unsprayed Sprayed Unsprayed Sprayed Unsprayed Site 1 Trial 1 (d1) 24 ± 2.3 0 0 0 0 0 0 0 0 0 0.0 Trial 1 (d15) 18 ± 1.7 2 ± 0.2 0 0 0 0 0 0 0 0 Trial 2 (d1) 205 ±19.6 97 ± 9.0 15 ± 1.4 3 ± 0.3 3 ± 0.3 1 ± 0.2 1 ± 0.2 1 ± 0.2 0 1 ± 0.2 88.9 Trial 2 (d15) 52 ± 5.0 8 ± 0.8 4 ± 0.4 0 3 ± 0.3 0 0 0 0 1 ± 0.2 Trial 3 (d1) 25 ± 2.3 1 ± 0.2 3 ± 0.3 1 ± 0.2 1 ± 0.2 0 1 ± 0.2 1 ± 0.2 1 ± 0.2 1 ± 0.2 63.5 Trial 3 (d15) 9 ± 0.9 0 0 0 0 0 0 0 0 0 Site 2 Trial 1 (d1) 134 ±12.8 0 0 0 0 0 0 0 0 0 0.0 Trial 1 (d15) 98 ± 9.4 3 ± 0.3 1 ± 0.2 0 2 ± 0.3 0 0 1 ± 0.2 0 1 ± 0.2 Trial 2 (d1) 25 ± 2.3 1 ± 0.2 0 0 0 0 0 0 0 1 ± 0.2 25.4 Trial 2 (d15) 17 ± 1.6 16 ± 1.5 22 ± 2.1 17 ± 1.6 13 ± 1.2 4 ± 0.4 1 ± 0.2 1 ± 0.2 1 ± 0.2 2 ± 0.2 Trial 3 (d1) 21 ± 2.0 2 ± 0.2 4 ± 0.4 3 ± 0.3 2 ± 0.2 1 ± 0.2 0 2 ± 0.2 1 ± 0.2 2 ± 0.2 88.9 Trial 3 (d15) 2 ± 0.2 1 ± 0.2 0.5 ± 0.04 0.75 ± 0.07 0.5 ± 0.04 0 0 0.5 ± 0.04 0 0

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Table 6-2. Mean ± SEM number of Metarhizium anisopliae and Isaria fumosorosea CFUs/mm2 from sprayed and unsprayed Prunus persica, peach leaves. Leaves were collected on the day of treatment application (d1) and 15 days later (d15). Dashed lines indicate contamination by Beauveria bassiana CFUs that masked the evidence of other entomopathogenic fungi CFUs.

Collection Met 52® EC PFR-97™ 20% (day) CFU’s/mm2) CFU’s/mm2) Sprayed Unsprayed Sprayed Unsprayed Site 1 Trial 1 (d1) 7 ± 0.7 0 1 ± 0.2 0 Trial 1 (d15) 1 ± 0.2 0 0 0 Trial 2 (d1) ------Trial 2 (d15) ------0 0 0 Trial 3 (d1) ------0 Trial 3 (d15) 0 0 0 0 Site 2 Trial 1 (d1) 12 ± 1.1 0 1 ± 0.2 0

Trial 1 (d15) ------0 ------0

Trial 2 (d1) 1 ± 0.2 0 1 ± 0.2 0

Trial 2 (d15) ------Trial 3 (d1) ------Trial 3 (d15) ------0

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CHAPTER 7 CONCLUSIONS

The overall goal of my research was to evaluate pest management tools for control of Myllocerus undecimpustulatus undatus in the landscape and horticulture industry. These approaches should support sustainable methodologies to manage this invasive weevil within Florida’s natural and built environment. I focused on the following objectives: 1) Design a successful rearing method to identify the duration of each developmental interval in the life cycle of M. undecimpustulatus undatus; 2) Analyze historical data and measure cold tolerance of M. undecimpustulatus undatus to determine its potential distribution in North America; 3) Assess the survival and mortality of adult M. undecimpustulatus undatus exposed to biopesticides in laboratory assays; and 4) Assess the survival and mortality of adult M. undecimpustulatus undatus exposed to biopesticides on peach leaves in the field.

Successful rearing of M. undecimpustulatus undatus was partially achieved through a plant-based system that allowed observation and examination of the developmental stages from egg to adult. The clear tubes provided surveillance of larvae consuming roots of Eleusine coracana L. This is the first known record of larval root consumption. This rearing method required a diligent watering schedule. If soil dried out, the grass would die; if the soil was overwatered, the larvae would drown. The larval stage of this weevil is not only sensitive to excess moisture but also to overcrowding.

Each tube started with 20-25 neonates, but by the end of 30-40 days, only 2-4 larvae survived. Observing larvae through the tube, I saw that if one touched another, the larger larva would respond aggressively. Suggested modifications to this rearing system would be to 1) add neonates to a Petri dish with soil and roots, transferring them in 1

111 week to the tubes, 2) place only five 1st instars in each tube, and 3) use a modified plastic pipette to direct the water to the roots in the tubes. Another suggestion would be to experiment with different artificial diets. The goal would be to devise a rearing system to produce large numbers of 3rd instars available for experimentation.

The cold tolerance study indicates an acclimation of M. undecimpustulatus undatus adults to winter conditions. In addition, it provides a greater understanding of the temperature requirements that influence the population dynamics and dispersal of

M. undecimpustulatus undatus in a global landscape. A reasonable prediction of the extent of a northern distribution of M. undecimpustulatus undatus is that it will be limited by cold temperatures at 0° C for more than four days. This information is a critical component in developing an effective management plan for this weevil’s current and potential distribution. The movement of plant material from Florida across state lines requires inspection, usually of the growth above the soil. It is recommended that an appropriate chemical treatment be applied to the soil prior to movement of plant materials. Plants with feeding damage should be avoided unless treated.

A comprehensive determination of the effect of cooler temperatures should include the influence of low temperature on fecundity and longevity of this weevil. The immature stages of M. undecimpustulatus undatus live in the soil, which may provide a buffer to cooler air temperatures. The manner of which low soil temperatures affect larval and pupal survivorship should be studied.

Two biopesticides in the laboratory study, Entrust and BotaniGard, reduced survival time and consistently performed well in killing M. undecimpustulatus undatus adults compared to the other treatments (PyGanic, AzaMax, PFR-97, Met52, Sevin and

112 water only). Both Entrust and AzaMax are effective in reducing plant damage.

BotaniGard (Beauveria bassiana) is twice more efficacious than PFR-97 (Isaria fumosorosea) and Met 52 (Metarhizium anisopliae). Entrust provides effective control of adult weevils and potentially could be integrated with other treatments. The effectiveness of BotaniGard tested under controlled conditions in the laboratory may yield different results in the field.

BotaniGard consistently performs well in killing adult weevils on peach leaves in the field and is four times more efficacious than PFR-97 and Met52. AzaMax produces a low mortality rate but is more effective in suppressing plant damage, similar to the results in the laboratory bioassay. High mortality caused by the B. bassiana in

BotaniGard and reduced feeding caused by AzaMax suggest that M. undecimpustulatus undatus adults may be well managed in the field using these two commercially available products in combination. The high mortality and reduced feeding of weevils exposed to

Entrust make this biopesticide another option for the control of M. undecimpustulatus undatus adults.

Management of M. undecimpustulatus undatus adults should be with a combination of biopesticides to increase effectiveness and reduce pesticide resistance.

Since the immature stages of this weevil are in the soil, further investigation of the effect of soil temperature and biopesticide treatments needs to be investigated.

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APPENDIX PERSONNAL COMMUNICATION

Indian River Research and Education Center 2199 South Rock Road Fort Pierce, FL 34945-3138 Tel. (772) 468-3922

Fax (772) 468-5668 Internet: www.irrec.ifas.ufl.edu

July 23, 2018

Anita Neal Entomology & Nematology Department University of Florida Gainesville, FL

Dear Anita:

The letter confirms that indeed I provided to you direct communication that my attempts to find biological control agents for Myllocerus undecimpustulatus undatus in Sri Lanka were unsuccessful. You may communicate this information in your dissertation and indicate me as the source.

Best regards,

Ronald D. Cave Professor and Center Director 772-577-7378

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BIOGRAPHICAL SKETCH

Anita Neal’s interest in entomology was evident at a young age. The first association with an ant farm led to a Bachelor of Arts degree in biology from the State

University of New York in 1979. Anita gained professional experience working for the

University of Florida as an Environmental Horticulture agent for Saint Lucie County

Extension from 1995 to 2016. Anita taught horticulture, entomology, and integrated pest management to young children, volunteers, homeowners, and clientele in the ornamental and turf industries. In 2002, Anita was promoted to County Extension

Director after completing her Master of Agriculture degree in agricultural education and communication from the University of Florida.

Anita enrolled at the University of Florida and began her Doctor of Philosophy degree program in entomology and nematology in the summer of 2012. In 2016, she accepted a position as the UF / IFAS Southeast District Extension Director with responsibilities for over 70 faculty in 12 counties and the Seminole Tribe. Anita earned her Ph.D. in the summer of 2018.

Anita is a member of the Entomological Society of America, the Gamma Sigma

Delta Honor Society of Agriculture, and Epsilon Sigma Phi. She has professional certifications from the Florida Natural Resources Leadership Institute, the Florida

Department of Environmental Protection, and the University of Florida Green Industries

Best Management Practices. Anita received the Distinguished Service Award for

Excellence in Extension Education programs, an award for Outstanding Environmental

Service, and three awards from the Fort Pierce Garden Club.

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