University of Florida Thesis Or Dissertation

BEHAVIORAL, BIOLOGICAL, AND OUTREACH APPROACHES TO THE MANAGEMENT OF Microtheca ochroloma STÅL (COLEOPTERA: CHRYSOMELIDAE)

ANGIE ALEJANDRA NIÑO BELTRAN

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

2016

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

ACKNOWLEDGMENTS

To begin with, I would like to express my sincere gratitude to my advisor Dr.

Ronald Cave who has been a great mentor and whose patience, motivation, support, and immense knowledge have provided me the guidance and encouragement needed during the research and writing stages of my dissertation.

Besides my advisor, I would like to thank the rest of my committee, Dr. Heather

McAuslane, Dr. Oscar Liburd, and Dr. Xin Zhao, for their insightful suggestions, comments, and expertise which made substantial improvements to this study. My sincere thanks also go to Dr. Pasco Avery who provided me access to his laboratory.

His experience and receptivity provided me with ideas to improve my research.

I thank the students, interns, 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 and all the fun we had at the BBQs during these last 4 years. Also, I am grateful to my classmates and friends in the Entomology and

Nematology Department.

I want to thank the Departamento de Ciencia, Tecnología e Innovación of

Colombia and Fulbright for sponsoring my doctorate studies at the University of Florida.

Similarly, thanks are due to the USDA Organic Agricultural Research and Extension

Initiative and the Florida Department of Agriculture and Consumer Services for supporting my research. I must thank Diane Cordeau for her cooperation during the execution of my experiments.

Last but certainly not least, I would like to thank my family, especially my parents, sisters, and brother, my husband, and my in-laws for supporting me spiritually throughout my studies and life in general.

TABLE OF CONTENTS

page

ACKNOWLEDGMENTS ...... 4

LIST OF TABLES ...... 7

LIST OF FIGURES ...... 8

ABSTRACT ...... 10

CHAPTER

1 INTRODUCTION ...... 12

2 LITERATURE REVIEW ...... 16

Distribution of Microtheca ochroloma Stål ...... 16 Host Plants and Insect Damage ...... 16 Description and Life Cycle ...... 17 Seasonality ...... 19 Management Strategies ...... 24

3 EFFECT OF THREE ENTOMOPATHOGENIC FUNGI ON THE FECUNDITY AND FERTILITY OF MICROTHECA OCHROLOMA STÅL (COLEOPTERA: CHRYSOMELIDAE)...... 35

Materials and Methods...... 37 Plant material ...... 37 Insect colony ...... 37 Fungal formulations ...... 38 Experiment 1. Insects dipped in fungal solutions ...... 39 Experiment 2. Insects fed leaves applied with fungal solutions ...... 41 Statistical analysis ...... 42 Results ...... 42 Experiment 1. Insects dipped in fungal solutions ...... 43 Experiment 2. Insects fed leaves applied with fungal solutions ...... 45 Discussion ...... 46

4 EFFECT OF NON-CRUCIFER EXTRACTS ON HERVIBORY BY ADULTS OF MICROTHECA OCHROLOMA STÅL (COLEOPTERA: CHRYSOMELIDAE) ...... 55

Materials and Methods...... 57 Plants and insects ...... 57 Non-crucifer essential oils ...... 57 Experiment 1. No-choice test ...... 58 Experiment 2. Paired choice test ...... 59

Experiment 3. Whole plant paired choice test ...... 59 Experiment 4. Starvation time ...... 60 Statistical analysis ...... 60 Results ...... 61 Discussion ...... 63

5 EFFECT OF HIGH TEMPERATURES AND LONG-DAY PHOTOPERIOD ON THE REPRODUCTIVE BEHAVIOR OF ADULTS OF MICROTHECA OCHROLOMA STÅL (COLEOPTERA: CHRYSOMELIDAE) ...... 71

Materials and Methods...... 73 Plants and insects ...... 73 Experimental design ...... 73 Statistical analysis ...... 74 Results ...... 74 Discussion ...... 75

6 OUTREACH TO CRUCIFEROUS CROP GROWERS ...... 82

Materials and Methods...... 83 Online Survey ...... 83 Extension Training ...... 84 EDIS publication ...... 84 Results ...... 85 Online Survey ...... 85 Extension Training ...... 86 EDIS publication ...... 87 Discussion ...... 87

7 CONCLUSIONS ...... 93

APPENDIX

A ON-LINE SURVEY QUESTIONNAIRE ...... 95

B EXTENSION TRAINING SURVEY FORMAT SPANISH VERSION ...... 102

C EXTENSION TRAINING SURVEY FORMAT ENGLISH VERSION ...... 104

D POSTER USED IN THE EXTENSION TRAINING ...... 106

E EDIS PUBLICATION ENGLISH VERSION ...... 107

F EDIS PUBLICATION SPANISH VERSION ...... 130

LIST OF REFERENCES ...... 155

BIOGRAPHICAL SKETCH ...... 165

LIST OF TABLES

Table page

3-1. Microtheca ochroloma fecundity and fertility after dipping females in three entomopathogenic fungal formulations...... 52

3-2. Net reproductive rate per treatment after dipping the insects or spraying leaves with fungal solutions...... 52

3-3. Sublethal effect of three entomopathogenic fungal formulations on Microtheca ochroloma fecundity and fertility after feeding on treated leaves...... 52

4-1. Percentage of leaf area consumed by adults of Microtheca ochroloma when insects were offered an AzaMax-treated leaf and an untreated leaf...... 69

5-1. Mean number of days that females survived under each treatment combination...... 80

LIST OF FIGURES

Figure page

2-1 States in the United States in which Microtheca ochroloma has been reported...... 32

2-1 Developmental stages of Microtheca ochroloma ...... 32

2-3 Sexual dimorphism of adult Microtheca ochroloma ...... 33

2-4 Sexual dimorphism of adults of Microtheca ochroloma ...... 33

2-5 Sexual dimorphism of pupae of Microtheca ochroloma ...... 34

3-1 Changes in reproduction and survival of Microtheca ochroloma when dipped in fungal solutions or water ...... 53

3-2 Longevity of Microtheca ochroloma females treated with three fungal formulations ...... 53

3-3 Entomopathogenic fungal growth on Microtheca ochroloma females ...... 54

3-4 Changes in reproduction and survival of Microtheca ochroloma when insects were fed leaves sprayed with fungal solutions or water ...... 54

4-1 Percentage of leaf area consumed by Microtheca ochroloma when offered a bok choy leaf sprayed with one non-crucifer essential oil or water ...... 69

4-2 Percentage of leaf area consumed by Microtheca ochroloma when offered an untreated and an AzaMax-treated bok choy plant in a paired choice test ...... 70

4-3 Percentage of leaf area consumed by Microtheca ochroloma after being deprived of food for various periods of time ...... 70

5-1 Mean number of eggs laid per Microtheca ochroloma female in each treatment combination ...... 80

5-2 Mean number of eggs laid per day by Microtheca ochroloma females at four temperatures ...... 81

6-1 Map of Florida counties where survey respondents’ farms are located...... 90

6-2 Number of respondents reporting the crucifer crops commonly planted on their farms...... 90

6-3 Frequency of insect pests on crucifer crops reported by survey respondents. ... 91

6-4 Frequency of primary and secondary pests of crucifer crops reported by survey respondents...... 91

6-5 Extension training at an organic farm ...... 92

6-6 Change in knowledge before and after extension training...... 92

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

BEHAVIORAL, BIOLOGICAL, AND OUTREACH APPROACHES TO THE MANAGEMENT OF Microtheca ochroloma STÅL (COLEOPTERA: CHRYSOMELIDAE)

Angie Alejandra Niño Beltrán

December 2016

Chair: Ronald D. Cave Major: Entomology and Nematology

The yellowmargined leaf beetle, Microtheca ochroloma Stål, causes significant damage to crucifers in organic farms in the southern United States. Separated use of various control methods has not been sufficient to manage this pest and reduce economic losses. Therefore, the goal of this study is to evaluate behavioral, biological, and outreach approaches in the search for long term management of M. ochroloma.

The sublethal effect of 3 entomopathogenic fungi, Beauveria bassiana,

Metarhizium anisopliae, and Isaria fumosorosea, on the number of eggs laid and hatch rate was evaluated during the beetle’s adult lifetime. Two methods of application, insect dipping and leaf spraying, were tested. Even though the number of eggs laid by females treated with M. anisopliae tended to be numerically less, no significant differences in the number of eggs laid and hatch rate were found among treatments and a water only control group, regardless of the method of application. Females dipped in M. anisopliae solution lived, on average, 25 d less than females in the control.

Ten non-crucifer essential oils were tested to determine repellent or antifeedant properties against adults of M. ochroloma. The most effective oils that decreased herbivory by M. ochroloma were neem, hyssop, and thyme compared to leaves sprayed

with garlic and lavender. Subsequent tests evaluating a commercially available formulation of azadirachtin showed that adults of M. ochroloma ate 54% and 68% less area of treated leaves compared to the control in a no-choice and paired choice test, respectively.

High temperatures have a detrimental effect on fertility and longevity of M. ochroloma females. The mean number of eggs was 80% lower at 30 than at 21ºC.

Longevity of females of M. ochroloma decreased between 40 and 60% at 30 compared to 21ºC.

An online survey taken by a sample (n = 14) of Florida farmers showed that the two most cultivated crops are collards and kale. The diamondback moth and aphids are considered the insects that cause the greatest amount of yield loss. After training workers on the identification and monitoring of pests on crucifers, their level of knowledge on these topics increased 59 and 55%, respectively.

CHAPTER 1 INTRODUCTION

Microtheca ochroloma Stål (Coleoptera: Chrysomelidae) is an adventive species that is native to the southern region of South America (Jolivet 1950). It was reported for the first time in the United States in Alabama in 1945 (Chamberlin and Tippins 1948), and since then it has been reported in many states in the southern and central United

States (Haeussler 1951, Woodruff 1974, Balsbaugh 1978, Staines 1999, Guillebeau

2001, Gilbert et al. 2011, Marché 2013). The larvae and adults chew holes in the leaves of plants in the family Brassicaceae (Chamberlin and Tippins 1948, Woodruff 1974,

Ameen and Story 1997c, Balusu and Fadamiro 2011) and cause aesthetic damage that renders cruciferous crops unmarketable. This insect is considered a minor pest in conventional crucifer production where plants are protected from attack by synthetic insecticides that are targeted against other key pests (Capinera 2001). However, in organic vegetable production, farmers are restricted in the use of synthetic insecticides, and the few pest management tactics available to them are not enough to successfully manage M. ochroloma (Bowers 2003, Balusu and Fadamiro 2012). According to

Bowers (2003), there are 3 main reasons why M. ochroloma has reached an important pest status in Florida. Firstly, its activity and population growth coincide with the crucifer production season in the state, so there is plenty of food available during the most active periods of the pest, which include late fall, winter, and early spring seasons.

Secondly, the mild climate in Florida provides ideal conditions for the beetle to survive and reproduce. Finally, specific natural enemies of this alien species that could reduce the populations of M. ochroloma have not been found in Florida.

From the time M. ochroloma was recorded in the United States, various studies have been performed to understand the biology and ecology of this insect and to identify eco-friendly management tactics that can both slow the selection pressure caused by the continuous use of insecticides and be implemented on organic farms.

Intercropping with non-host plants (Bowers 2003), the use of straw mulch (Manrique et al. 2010), trap cropping (Balusu et al. 2015), the use of botanical and biological insecticides (Balusu and Fadamiro 2012, 2013), and the use of generalist predators and entomopathogenic fungi (Montemayor and Cave 2011, 2012, Niño and Cave 2015,

Montemayor et al. 2016) have shown no or limited efficacy against this beetle. Trap cropping using turnips can reduce the pest population and damage in cabbage, but its efficacy is reduced when it is used with other crops such as napa cabbage and mustard

(Balusu et al. 2015). The use of spinosad and pyrethroids causes high mortality of larvae and adults of M. ochroloma (Balusu and Fadamiro 2012, 2013), but their effectivenesscan be reduced in the long term if the insect develops resistance against the active ingredients. Inundative field releases of the generalist predator Podisus maculiventris Say (Hemiptera: Pentatomidae) helped to reduce the population and damage of initial infestations of M. ochroloma, but its efficacy decreases when the population of the pest reaches high numbers (Ronald Cave, personal communication).

As a result, the isolated use of each practice appears to be insufficient to manage this pest in the long term, suggesting that only an integrated approach combining different control tactics will be useful to manage this insect.

Fundamental information on the biology and ecology of insects is crucial for the successful implementation and integration of control methods (Biever et al. 1994).

Studies of the biology and ecology of M. ochroloma have provided information about insect morphology (Chamberlin and Tippins 1948, Jolivet 1950), life cycle (Chamberlin and Tippins 1948, Oliver and Chapin 1983, Marquini et al. 2003), reproduction (Ameen and Story 1997a, b), and food preference (Ameen and Story 1997c, Balusu and

Fadamiro 2011). However, there are unknown aspects of M. ochroloma’s biology, such as dormancy during summer. It is unknown the environmental conditions that can trigger a dormancy response. Information about dormancy is important to understand insect colonization as they move from dormancy sites to the crop. Similarly, studies on the dormancy of this insect might help to predict its first appearance in the crop, based on environmental conditions, which allow farmers to be alerted and implement control methods before the insect population reaches high numbers. A better understanding, in time and space, of M. ochroloma initial infestations can increase the efficacy of trap cropping and the use of generalist predators as mentioned above.

The combination of trap cropping with other methods that could make the crop of interest unsuitable or unattractive to the insect might also increase the trap crop’s efficacy against M. ochroloma. Repellents and antifeedants have been successfully used to reduce the presence and damage caused by many insect pests (Ladd et al.

1978, Pavela 2004, Showler et al. 2004). Until now, no plant extracts have been tested against M. ochroloma.

The effect of control methods on M. ochroloma reproductive capabilities can also be exploited for the management of this pest. The efficacy of the management strategies tested against M. ochroloma has been determined by evaluating the immediate effect of each strategy on the survival of the population under study (Balusu

and Fadamiro 2012, 2013). However, some control methods, such as the use of entomopathogenic fungi, can have a detrimental effect on reproduction.

Entomopathogenic fungi can affect an insect’s fertility and fecundity and contribute to reducing the insect population and insect damage though time (Castillo et al. 2000,

Dubois et al. 2004, Quesada-Moraga et al. 2004, 2006). The manner in which entomopathogenic fungi influence M. ochroloma reproduction has not been studied.

In addition to the evaluation and development of strategies to manage M. ochroloma, the transfer of information from these studies to farmers and the general public is an essential step in promoting the adoption of any pest management program.

Constant communication and interactive dialog among farmers, extension agents, and scientists are crucial to developing and adopting new technologies that are appropriate and fit the farmer’s needs (Röling and Pretty 1997). Extension efforts should provide farmers with enough tools and choices so they can make farm-specific decisions according to their own needs (Röling and Pretty 1997).

The overall goal of my research was to evaluate behavioral, biological, and outreach approaches that contribute to long term management of M. ochroloma. The research focused on 4 specific objectives:

1. Evaluate the effect of 3 entomopathogenic fungi on the fecundity and fertility of M. ochroloma.

2. Assess the effect of 10 non-crucifer plant essential oils on herbivory by adults of M. ochroloma.

3. Determine how changes in temperature and photoperiod might trigger a dormancy response in adults of M. ochroloma.

4. Determine the insect pest problems of most concern to crucifer farmers in Florida and increase accessibility to information on crucifer pests through a survey and extension activities.

CHAPTER 2 LITERATURE REVIEW

Distribution of Microtheca ochroloma Stål

The genus Microtheca is limited to the Neotropical region, specifically to the southern region of Latin America from where 14 species have been described (Jolivet

1950). Microtheca ochroloma is native to Argentina, but it is also commonly found in

Brazil and Chile (Jolivet 1950). It was first intercepted in New Orleans, Louisiana in

1945 on a shipment of grapes from Argentina, and it was first reported attacking cruciferous crops in Alabama in 1947 (Chamberlin and Tippins 1948). Since then, it has been reported in many states in the southern and central United States (Chamberlin and Tippins 1948, Haeussler 1951, Rohwer et al. 1953, Balsbaugh 1978, Staines 1999,

Guillebeau 2001, Gilbert et al. 2011, Marché 2013) (Figure 2-1).

Host Plants and Insect Damage

Microtheca ochroloma is restricted to plants in the family Brassicaceae (Jolivet

1950). It is known to attack cabbage (Brassica oleracea L. var. capitata), napa cabbage

(B. pekinensis Lour.), collard (B. oleracea var. acephala), mustard (B. juncea Cosson), radish (Raphanus sativus L.), turnip (B. rapa L.), white mustard (Sinapis alba L.), and watercress (Nasturtium officinale L.) (Chamberlin and Tippins 1948, Jolivet 1950,

Ameen and Story 1997a, Balusu and Fadamiro 2011, Riquelme Virgala et al. 2014).

Microtheca ochroloma also feeds on various Asian leafy greens such as bok choy (B. rapa L. ssp. chinensis) (Niño and Cave 2015), mizuna (B. rapa L. var. kyona) and mibuna (B. rapa L. ssp. japonica) (Bowers 2003), and komatsuma (B. rapa L. var. perviridis) and tatsoi (B. rapa L. var. rosularis) (personal observations). Studies on M. ochroloma host preference indicate that this insect has a strong preference for turnip

(Chamberlin and Tippins 1948, Ameen and Story 1997c, Balusu and Fadamiro 2011), mustard (Ameen and Story 1997c), and napa cabbage (Balusu and Fadamiro 2011) over cabbage and collards. Laboratory experiments using a four-choice olfactometer have shown that host plant volatiles play an important role in host finding by M. ochroloma (Balusu and Fadamiro 2011). However, this insect might also use other clues (i.e. visual, tactic) to find, taste, and accept its host (Balusu and Fadamiro 2011).

For example, the plants most preferred for consumption and oviposition by M. ochroloma seem to have softer foliage than the less preferred hosts which have tougher, waxy leaves (Ameen and Story 1997c).

Both larvae and adults chew small holes with irregular contours in the leaves of plants (Chamberlin and Tippins 1948, Jolivet 1950, Ameen and Story 1997c). Larvae feed gregariously, consuming large areas of leaf tissue in a short time (Chamberlin and

Tippins 1948, Jolivet 1950). The insects feed initially on the leaves of plants, but once all leaf tissue has been depleted they feed on other plant tissues such stems and roots

(Rhodes and Liburd 2014).

Description and Life Cycle

Microtheca ochroloma is a small, oval insect (about 5 mm long) characterized by a dominant black or dark brown color, a pale yellow or white elytral border, and 4 rows of elytral punctures (Chamberlin and Tippins 1948, Jolivet 1950, Rohwer et al. 1953,

Woodruff 1974, Marquini et al. 2003) (Figure 2-2 d). The apex of the tibia, the 3 basal tarsomeres on each leg, and the last 2 abdominal ventrites are also yellow (Jolivet

1950). Males are smaller than females (Jolivet and Petitpierre 1981, Ameen and Story

1997b), and they can be accurately differentiated from females by the apical border of the fifth sternite; in females, the segment has a rounded apical margin with a shallow

emargination on each side, whereas in males it has a truncated apex with a distinct emargination on each side (Figure 2-3). Additionally, if the abdomen of the male is gently pressed dorsoventrally, the large eighth tergite extends from the abdomen, whereas in the female it does not (Figure 2-4). Insects can also be sexed at the pupal stage. Female pupae have 2 knob-like structures on the ventral side of the abdomen, which are absent in male pupae (Ameen 1996) (Figure 2-5).

Elongated, bright orange eggs are laid singly or in small clutches on the soil or under fallen leaves (Woodruff 1974, Bowers 2003) (Figure 2-2 a). Larvae are carabiform with a soft body that varies from gray to yellow-brown in coloration and a black, sclerotized head capsule (Marquini et al. 2003) (Figure 2-2 b). The body of the larva is covered with fine hairs. Larvae usually go through 4 instars (Ameen and Story 1997a,

Riquelme Virgala et al. 2014). However, Ameen and Story (1997a) reported that a small proportion of the population goes through a fifth instar. Prior to pupation, the larva creates a brownish cocoon made of fibrous material that is excreted through the anal opening (Woodruff 1974, Oliver and Chapin 1983) (Figure 2-2 c). The insect pupates on the undersides of dry leaves, twigs, or debris where the cocoon blends into the surroundings (Rohwer et al. 1953). Pharate adults remain in the cocoon for 1 or 2 d, until they become completely sclerotized and fully pigmented (Oliver and Chapin 1983).

Several studies on M. ochroloma developmental time and survival under various controlled environmental conditions and using different plant species as hosts have shown that the duration of each stage is significantly affected by temperature but not host type, whereas survival is affected by both (Ameen and Story 1997a, Manrique et al. 2012). The time required by M. ochroloma to develop from egg to adult decreases as

temperature increases. At 15, 20, 25, and 30°C, the insects require 57, 31, 21, and 17 d, respectively, to complete development (Manrique et al. 2012). On the other hand, survival of immature stages was reduced from 80% at 15, 20, and 25°C to 24% at 30°C

(Manrique et al. 2012). Similarly, multi-generation survivorship decreased when M. ochroloma was reared on cabbage and collard for 4 continuous generations, compared to insects reared on mustard, turnip, or radish (Ameen and Story 1997a).

Once adults of M. ochroloma emerge from the cocoon, copulation occurs within

5-8 d for prolonged periods of time, and adults copulate repeatedly during their lifetime

(Oliver and Chapin 1983, Riquelme Virgala et al. 2014). Females start laying eggs 3-7 d after copulation, and the number of eggs laid varies with host plant (Oliver and Chapin

1983, Ameen and Story 1997b). At 20°C, M. ochroloma laid more eggs on turnip followed by mustard, radish, cabbage, and collard (Ameen and Story 1997b).

Depending on the host, M. ochroloma adults can live up to 68-105 d (Ameen and Story

1997b).

Seasonality

Microtheca ochroloma is a cold season pest, whose active period in Florida lasts from fall to the end of spring (Chamberlin and Tippins 1948, Rohwer et al. 1953, Oliver and Chapin 1983, Bowers 2003). The beetles start appearing in the field in October, but are most abundant later in the season (Rohwer et al. 1953, Oliver and Chapin 1983,

Bowers 2003). In June, the M. ochroloma population decreases and limited observations indicate that this species enters a period of dormancy during the hot months (Chamberlin and Tippins 1948, Rohwer et al. 1953, Oliver and Chapin 1983,

Bowers 2003). Several authors have suggested that M. ochroloma aestivates – i.e., undergoes a summer dormancy (Chamberlin and Tippins 1948, Rohwer et al. 1953,

Oliver and Chapin 1983). But there are no studies that support and describe the aestival behavior of this beetle, including the habitats where this insect oversummers. For example, Rohwer et al. (1953) stated that, based on limited observations, M. ochroloma aestivates on weeds but no data were presented to support that hypothesis. Similar observations were reported by Balusu et al. (2015) who reported that M. ochroloma aestivates on wild hosts at the field edges. In contrast, Bowers (2003) suggested that M. ochroloma spends summer within the field, where crucifers had been grown, since she found more insects in interior plots first than in exterior plots; movement from the border to the center of the field was not evident. Additionally, she surveyed the borders of the experimental field and did not find alternative cruciferous hosts. However, she did not have records of the crops planted in the experimental plots in previous seasons and whether they were infested with M. ochroloma. Similarly, Oliver and Chapin (1983) stated that after passing through several generations, M. ochroloma leaves the host plants and aestivates; although they found few specimens in ground trash at the end of summer, they did not provide more information about possible aestivating habitats.

Adults are considered the oversummering stage (Oliver and Chapin 1983,

Bowers 2003). Previous studies showed that the survival of immature stages of M. ochroloma decreases considerably at 30°C (Manrique et al. 2012). Besides, other authors reported finding only adults during mid-summer, and the adult stage is also the first to appear in the fall (Oliver and Chapin 1983, Bowers 2003). Lastly, adults have been reported feeding on non-cruciferous food sources such as pollen of rosaceous plants (Jolivet 1950) and sugar solution (Marquini et al. 2003), which may ensure their

survival when cruciferous hosts are not present or at the end of the season when host plants have been heavily defoliated.

It is clear that M. ochroloma experiences a non-reproductive period during summer but the oversummering adults must use a strategy to survive for at least 6 months without food and under conditions that hinder reproduction. Insects have developed highly evolved mechanisms to cope with environmental changes, depending on the duration, magnitude, and predictability, which allow them to synchronize with favorable conditions and survive during unfavorable periods (Tauber and Tauber 1976,

Tauber et al. 1986). Acyclic, unpredictable changes (e.g., localized deterioration of food, lack of mates, prairie fires) are overcome by various physiological, behavioral, and genetic mechanisms such as aseasonal quiescence and aseasonal migration, which involve primarily nervous responses (Tauber et al. 1986). On the other hand, when seasonal, predictable changes in conditions occur (e.g., diurnal, lunar, tidal, or seasonal cycles), insect responses involve biological clocks or other time-measuring mechanism

(e.g., behavioral and physiological rhythms) governed by the neuroendocrine system

(Tauber et al. 1986).

Considering that M. ochroloma responds to seasonal changes that occur in a cyclic and persisting basis, I will explain 2 possible seasonal dormancy responses that have been reported in insects. The first type is a non-diapause dormancy (i.e., quiescence) in which the insect goes through a state of arrested metabolism caused by recurring unfavorable conditions that prevent growth and reproduction (Tauber et al.

1986, Danks 1987). The insect responds rapidly to changes in environmental conditions, and if favorable conditions are restored, growth and reproduction resume in

a short period of time (Tauber et al. 1986, Danks 1987). The second type is a diapause- mediated dormancy (i.e., diapause). This dormancy type involves low metabolic activity under adverse environmental conditions, but unlike quiescence, diapause occurs as a response to environmental stimuli that precede unfavorable conditions perceived by earlier developmental stages than those that undergo diapause (Tauber et al. 1986,

Danks 1987). The stimuli that trigger diapause are not, in themselves, favorable or unsuitable for development, but they announce a variation in environmental conditions

(Tauber et al. 1986). The most reliable clue is photoperiod, although temperature, humidity, and biotic factors also provide cues to future seasonal changes (Tauber et al.

1986). Another aspect that differentiates diapause from quiescence is that, under favorable conditions, the former is not terminated until after the occurrence of a specific physiological process in order to prevent undergoing premature growth and reproduction (Tauber et al. 1986).

Behavioral, physiological, and morphological characteristics differ among dormant insects and insects that develop continuously (Tauber et al. 1986, Danks

1987). In general, insects in a dormant state are less receptive to food and reproductive stimuli (Tauber et al. 1986, Danks 1987). Behavioral changes in food consumption range from a non-feeding state to continuous or sporadic feeding at a reduced rate

(Tauber et al. 1986, Danks 1987). Oviposition is stopped or reduced in dormant females and, in some species, male mating behavior is completely suppressed (Tauber et al.

1986, Danks 1987). Another common behavior of insects preparing to enter dormancy is migration to and from protective sites (Tauber et al. 1986, Danks 1987). Alterations in physiological characteristics involve the storage of lipids and other reserves as fat

content in the insect’s fat body and reduced metabolism and O2 consumption (Tauber et al. 1986, Danks 1987). Finally, morphological features such as color and size of body parts might vary in dormant insects (Tauber et al. 1986, Danks 1987). Ovarian development is reduced to small ovaries and accessory glands, and a decrease in the size of the male reproductive system is also apparent (Danks 1987).

The dormancy strategy used by M. ochroloma is unknown. Bowers (2003) proposed 3 possible dormancy strategies to explain the low populations for M. ochroloma during summer: (1) the beetle undergoes diapause; (2) the beetle experiences quiescence; or (3) M. ochroloma has multiple oversummering strategies in which some insects enter dormancy while others continue developing and ovipositing on a limited basis. Bowers’ (2003) field experiments did not clarify which strategy is used by M. ochroloma, however, she reported that she collected 12 adults on turnip plants (damage caused by M. ochroloma was not apparent) on July 1 at an organic farm southwest of Gainesville, FL and took them to the laboratory where she placed them in an environmentally controlled chamber at 25ºC and 14-h photoperiod. The adults started copulating and laying eggs 24 h after putting them in the chamber (Bowers

2003). Manrique et al. (2012) collected reproductively active adults in the field during fall/winter season and placed them in environmentally controlled chambers at either 25 or 30ºC (10-h photoperiod in both chambers). The insects at 30ºC stopped laying eggs, while those at 25ºC remained reproductively active (Manrique et al. 2012). The rapid response of M. ochroloma to changes in temperature suggests that a non-diapause dormancy (i.e., quiescence) could explain the response of this insect to changes in environmental conditions but no one has maintained adults for extended periods under

unfavorable (= summer) conditions (e.g., 30ºC and 14-h photoperiod) to know how long they would survive under these conditions and whether or not the insects can resume reproduction if placed under favorable (= winter) conditions.

The environmental cue that triggers the dormancy response in M. ochroloma is also unknown. Oliver and Chapin (1983) tested the effect of 3 photoperiods (ambient photoperiod during the summer season, 12-h photoperiod, and 14-h photoperiod) on adult oviposition behavior. They found that ambient photoperiod increased female lifespan and the number of days from emergence to oviposition compared to the 12-h photoperiod (Oliver and Chapin 1983). The total number of eggs was also higher at ambient photoperiod, but this was due to a longer lifespan of females instead of a higher daily oviposition rate (Oliver and Chapin 1983). The authors suggested that, based on successive generations developing in the laboratory at ambient photoperiod,

27ºC, and 50% RH, temperature plays an important role in M. ochroloma dormancy

(Oliver and Chapin 1983). However, no studies have tested the effect of high temperature coupled with long-day photoperiod on M. ochroloma’s dormancy response.

Management Strategies

Chemical, cultural, and biological management methods have been tested to regulate M. ochroloma populations. The availability of management methods and their efficacy depend on the type of farming system used (e.g., conventional versus organic).

So far, the most effective management methods have been the application of insecticides on a weekly basis (Rhodes and Liburd 2014). In conventional agriculture, the first pesticide used was 75% rotenone dust, which effectively killed all adults and larvae present at that time (Chamberlin and Tippins 1948). Usually, M. ochroloma is managed by insecticides applied to control other pests (Rhodes and Liburd 2014).

Webb (2013) published a list of insecticides used for the management of pests in crucifer crops.

Since organic growers are restricted in the use of synthetic insecticides, they depend on a few biopesticides and botanical insecticides approved by the Organic

Material Review Institute (OMRI) to manage M. ochroloma. Among the OMRI-accepted insecticides tested against this pest are PyGanic® (a.i. pyrethrum), Aza-Direct® EC (a.i. azadirachtin), Entrust® WP (a.i. spinosad), and Novodor® (a.i. protein crystal produced by Bacillus thuringiensis ssp. tenebrionis) (Balusu and Fadamiro 2012, 2013).

According to Balusu and Fadamiro (2012), the 2 most effective formulations in the field against larvae and adults of M. ochroloma were Entrust® WP and PyGanic®, whereas the other insecticides had inconsistent efficacy or failed to control the pest. In laboratory tests, Entrust® WP and PyGanic® caused 100% mortality within 24h. Plant extracts, such as pó-de-fumo (Nicotiana tabacum L., Solanaceae), ramo de cinamomo (Melia azedarach L., Meliaceae), and DalNeem® (a.i. azadirachtin), have been shown to cause mortality of yellowmargined leaf beetle larvae and adults in laboratory tests (Dequech et al. 2008), but field tests are needed to determine their efficacy under field conditions.

Aside from causing mortality, plant extracts can also be used as repellents and antifeedants to reduce the presence of or damage caused by insect pests. For example, extracts from 4 plants of the family Lamiaceae, marjoram (Origanum majorana L.), sage

(Salvia officinalis L.), hyssopus (Hyssopus officinalis L.), and horehound (Marrubium vulgare L.), applied on potato leaves had a strong repellent and antifeedant effect on the feeding behavior of adult Colorado potato beetles, Leptinotarsa decemlineata (Say)

(Coleoptera: Chrysomelidae) (Pavela 2004). The pollen beetle, Meligethes aeneus

Fabricius (Coleoptera: Nitidulidae), which damages oilseed rape, Brassica napus L., was strongly repelled by lavender (Lavendula angustifolia Miller) essential oil

(Mauchline et al. 2005). Papachristos and Stamopoulos (2002) reported that most of the

13 essential oils evaluated against Acanthoscelides obtectus (Say) (Coleoptera:

Chrysomelidae) caused repellency, reduced fecundity, and had negative effects on egg hatch, neonate larval survival, and offspring emergence. Azadirachtin, a compound extracted from neem trees (Azadirachta indica A. Juss), has been shown to repel the boll weevil, Anthonomus grandis grandis Boheman, the banana root borer,

Cosmopolites sordidus Germar (both Coleoptera: Curculionidae), and the Japanese beetle, Popillia japonica Newman (Coleoptera: Scarabeidae) (Ladd et al. 1978,

Musabyimana et al. 2001, Showler et al. 2004). Many products that have azadirachtin as the active ingredient are commercially available. Although Aza Direct® EC tested against M. ochroloma induced no mortality (Balusu and Fadamiro 2012), none of these or other commercially available products have been tested as repellents against this pest.

In addition to chemical control, cultural tactics including straw mulch, intercropping, and trap cropping have been tested. Covering the soil with straw mulch increased the population of M. ochroloma in small plots and reduced the number of ground predators present (Manrique et al. 2010). Similarly, intercropping host plants with non-host vegetables has had little success in the management of M. ochroloma.

When mizuna was intercropped with oak leaf lettuce (Lactuca sativa var. berenice), no difference in the number of insects present on host plants between the control and intercropped plots was found (Bowers 2003). The ability of this specialist insect to locate

its host plant through a milieu of plant volatiles (Balusu and Fadamiro 2011) likely reduces the success of intercropping. However, intercropping with aromatic non-host plants has not yet been evaluated against this pest. Planting turnips at the periphery of the crop of interest to act as a trap crop –i.e., the use of a more preferred host to lure the pest and prevent colonization of the main crop– was shown to be effective for reducing the number of adults and larvae of M. ochroloma and their damage (Balusu et al. 2015). However, the efficacy of this method varies depending on the main crop species. When the crop of interest is a less attractive crucifer crop (e.g., cabbage and collard), planting turnips as a trap crop can effectively reduce the damage caused to the main crop (Balusu et al. 2015). On the other hand, when an equally attractive crop (e.g., mustard and napa cabbage) is cultivated, then trap cropping using turnips is not sufficient to reduce damage to yield marketable produce (Balusu et al. 2015).

Studies on the biological control of M. ochroloma have focused on 2 main components: releases of generalist predators and the use of entomopathogens. Many generalist predators have been observed preying on all stages of M. ochroloma. The natural enemies include Podisus maculiventris Say (Hemiptera: Pentatomidae)

(Montemayor and Cave 2011, 2012), Chrysoperla rufilabris Burmeister (Neuroptera:

Chrysomelidae) (Niño and Cave 2015), Hippodamia convergens Say (Coleoptera:

Coccinellidae) (Montemayor and Cave 2009), Stiretrus decastigmus Herrich-Schaeffer

(Hemiptera: Pentatomidae) (Poncio et al. 2010), and Toxomerus duplicatus Wiedemann

(Diptera: Syrphidae) (Sturza et al. 2011); the first 3 of these have been observed preying on M. ochroloma in Florida (Montemayor and Cave 2009), but none of them provide significant control of the pest population. Inundative field releases of P.

maculiventris in organic farms helped to reduce the population and damage caused by initial infestations of M. ochroloma (Ronald Cave, personal communication). However, releases of P. maculiventris cannot be used as a stand-alone tactic when the population of the pest reaches high numbers.

Various species of entomopathogenic fungi (EPF) have been evaluated against

M. ochroloma. These include Beauveria bassiana (Balsamo) Vuillemin, Metarhizium anisopliae (Metschnikoff) Sorokin (both Hypocreales: Clavicipitaceae), and Isaria

(=Paecilomyces) fumosorosea Wize (Hypocreales: Cordycipitaceae) (Balusu and

Fadamiro 2012, 2013, Montemayor et al. 2016). These EPF are distributed worldwide and can be commonly found in the soil of agricultural ecosystems (Meyling and

Eilenberg 2006, 2007). They are natural enemies of arthropods and infect many species of insects within several orders (Hajek and St. Leger 1994, Shah and Pell 2003, Meyling and Eilenberg 2007, Wraight et al. 2007). Their capacity of infection is due to a synchronization of their life cycles with the host stages and environmental conditions

(Shah and Pell 2003). Pathogenesis starts with the adhesion of a conidium (spore) to the cuticle of the insect and subsequent penetration into the host’s hemocoel (Hajek and St. Leger 1994, Shah and Pell 2003, Wraight et al. 2007). In heavily sclerotized insects, fungal penetration occurs via the spiracles or arthrodial membranes (Hajek and

St. Leger 1994). Once inside, the fungus releases enzymes and toxins (e.g., destruxins, bassianolide, and leucinostatins produced by M. anisopliae, B. bassiana, and Isaria spp., respectively) and grow rapidly to overcome host defenses; host death occurs as a combination of mechanical damage, invasion of organs, depletion of nutrients, and toxicosis (Hajek and St. Leger 1994, Shah and Pell 2003, Wraight et al. 2007). After

host death, the fungus emerges from the body and sporulation and conidiogenesis produce many spores that are ready to infect other hosts (Hajek and St. Leger 1994,

Shah and Pell 2003, Wraight et al. 2007). Spores can be passively dispersed by wind, rain, and other arthropods (Hajek and St. Leger 1994, Meyling and Eilenberg 2007,

Wraight et al. 2007).

The capacity of EPF for producing spores and advances in mass production techniques have stimulated the development and commercialization of mycoinsecticides, which allows inundative augmentation of the fungus for rapid, short- term control of pests, in the same way that chemical insecticides are used (Shah and

Pell 2003, Wraight et al. 2007). In the U.S., many fungal formulations of these 3 EPF are commercially available, such as Botanigard®, Mycotrol®, Balence®, and CornGard®, each of which contains spores of a specific strain of B. bassiana that infects insects within specific orders (Faria and Wraight 2007). Isaria fumosorosea formulations labeled

PFR-97® 20% WDG and Nofly® and M. anisopliae formulations labeled Tick-EX®,

Taerain®, and Bio-Path®, among others, are available in the U.S. (Faria and Wraight

2007).

Laboratory and field studies of some commercially available fungal formulations to control M. ochroloma showed that, in general, these entomopathogenic fungi have a slow activity and the percentage of mortality is low (Balusu and Fadamiro 2012, 2013).

For instance, Botanigard® and Mycotrol® were slightly effective against larvae in laboratory tests, causing higher mortality than the control treatment 5 d after inoculation; however, mortality was less than 50% (Balusu and Fadamiro 2013). None of the fungal formulations tested caused significant adult mortality (Balusu and Fadamiro 2013). At

the same time, when those fungal formulations were tested in the field, none efficiently suppressed the pest or reduced pest damage (Balusu and Fadamiro 2012). Similarly, laboratory experiments testing the effectiveness of PFR-97® on 5 stages of M. ochroloma, egg, first and third instar, pupa, and adult, showed that the larval stage is the most susceptible to infection by this fungus, but the percentage of infection was still under 40% (Montemayor et al. 2016).

The methods studied until now have focused on mortality of the pest. However, entomopathogenic fungi can have sublethal effects that reduce an insect’s fertility, fecundity, and consumption, which can reduce the insect population and insect damage through time (Castillo et al. 2000, Dubois et al. 2004, Quesada-Moraga et al. 2004,

2006, Wraight et al. 2007). Balusu and Fadamiro (2013) observed a reduction of M. ochroloma feeding and oviposition in the treatments applied with Mycotrol® and

Botanigard®. Topical applications of I. fumosorosea (strain CECT 2705), M. anisopliae

(strains CECT 2952 and EAMa 01/58), Aspergillus ochraceus Wilhelm (strain AV1), and

B. bassiana (strain EABb 01/103) on adults of Ceratitis capitata Wiedemann (Diptera:

Tephritidae) reduced fertility by about 65, 53, 50, and 65%, respectively (Castillo et al.

2000, Quesada-Moraga et al. 2006). Beauveria bassiana and M. anisopliae also reduced egg eclosion by about 47 and 44%, respectively (Castillo et al. 2000, Quesada-

Moraga et al. 2006). German cockroaches, Blatella germanica L. (Blattodea:

Blattellidae), treated with a solution of M. anisopliae strain EAMa 01/121 had decreased oothecal production and egg hatch rate (Quesada-Moraga et al. 2004). Similarly, a reduction in oviposition occurred with exposure of adult Asian longhorned beetles,

Anoplophora glabripennis (Coleoptera: Cerambycidae), to 2 commercially available

Beauveria spp. strains (Dubois et al. 2004).

As shown above, it appears that the isolated use of each control practice is insufficient to manage M. ochroloma, suggesting that only an integrated approach combining various control tactics will be useful to manage this insect. A combination of chemical, cultural, and biological control methods should be adopted to modify the behavior of the insect and to divert the pest from the crop of interest and attract it to other resources where the insects can be retained or congregated to facilitate the implementation of management strategies and reducing costs, non-target effects to other species, and environmental contamination.

Figure 2-1. States in the United States in which Microtheca ochroloma has been reported.

Figure 2-1. Developmental stages of Microtheca ochroloma: (a) eggs, (b) larva, (c) pupae, and (d) adults. Photographs by Angie Niño.

Figure 2-3. Sexual dimorphism of adult Microtheca ochroloma. Apical border of fifth sternite in (a) female and (b) male. Photographs by Angie Niño.

Figure 2-4. Sexual dimorphism of adults of Microtheca ochroloma. Females (a) do not expose a tergite when their abdomen is pressed, whereas males (b) do. Photographs by Angie Niño.

Figure 2-5. Sexual dimorphism of pupae of Microtheca ochroloma. Females (left) have two knob-like structures on the abdomen, which are absent in males (right). Photographs by Angie Niño.

CHAPTER 3 EFFECT OF THREE ENTOMOPATHOGENIC FUNGI ON THE FECUNDITY AND FERTILITY OF MICROTHECA OCHROLOMA STÅL (COLEOPTERA: CHRYSOMELIDAE)

Microtheca ochroloma Stål, known as the yellowmargined leaf beetle, is an important pest of cruciferous crops in the southern U.S (Chamberlin and Tippins 1948,

Haeussler 1951, Rohwer et al. 1953, Staines 1999, Guillebeau 2001, Rhodes and

Liburd 2014). Larvae and adults consume the leaves of the plant until complete defoliation, after which they move to other plant tissues such as stems and roots

(Chamberlin and Tippins 1948, Jolivet 1950, Ameen and Story 1997c, Rhodes and

Liburd 2014). Microtheca ochroloma is primarily controlled through synthetic insecticides applied to manage other leaf-feeding pests (Webb 2013, Rhodes and

Liburd 2014). However, insecticide resistance, concerns about human and environmental safety, and the restricted use of synthetic insecticides in organic farming have intensified the efforts to find alternative control methods. Biological control, through the use of entomopathogenic fungi, is a potential and more ecofriendly alternative.

Entomopathogenic fungi are distributed worldwide and can be commonly found in the soil of agricultural ecosystems (Meyling and Eilenberg 2006, 2007). They are natural enemies of arthropods and infect many species of insects within several orders

(Hajek and St. Leger 1994, Shah and Pell 2003, Meyling and Eilenberg 2007, Wraight et al. 2007). Three species of entomopathogenic fungi have been evaluated against M. ochroloma. These are Beauveria bassiana (Balsamo) Vuillemin, Metarhizium anisopliae

(Metschnikoff) Sorokin (both Hypocreales: Clavicipitaceae), and Isaria (=Paecilomyces) fumosorosea Wize (Hypocreales: Cordycipitaceae) (Balusu and Fadamiro 2012, 2013,

Montemayor et al. 2016). Laboratory tests indicate that the larva of M. ochroloma is the most susceptible stage. Treating larvae with I. fumosorosea showed that this fungus caused reduction in growth, unsuccessful molting, and a mortality rate up to 39%

(Montemayor et al. 2016). Larval mortality rates up to 50% have also been reported by

Balusu and Fadamiro (2013). Unfortunately, laboratory and field studies have shown that these entomopathogenic fungi have slow activity and cause a low level of mortality of adult M. ochroloma, which reduced their effective use under field conditions (Balusu and Fadamiro 2012, Balusu and Fadamiro 2013, Montemayor et al. 2016).

To date, the control tactics evaluated have focused on mortality of the pest.

However, entomopathogenic fungi can have sublethal effects that reduce an insect’s fertility, fecundity, and food consumption, which can reduce the insect population and damage to plants through time (Castillo et al. 2000, Dubois et al. 2004, Quesada-

Moraga et al. 2004, 2006, Wraight et al. 2007). Several studies have shown a reduction in fecundity and fertility caused by entomopathogenic fungi on a variety of insect species (Castillo et al. 2000, Dubois et al. 2004, Quesada-Moraga et al. 2004, 2006).

Balusu and Fadamiro (2013) observed a reduction of M. ochroloma feeding and oviposition when adults were treated with products containing spores of B. bassiana, but the effect of this or other entomopathogenic fungi on M. ochroloma reproduction has not yet been quantified. The purpose of this study was to evaluate the effect of 3 entomopathogenic fungi, B. bassiana, M. anisopliae, and I. fumosorosea, on the fecundity and fertility of M. ochroloma.

Materials and Methods

Plant material

Win-Win Choi F1 bok choy (Johnny’s Selected Seeds, Winslow, ME), Organic

Baby Bok choy (Sustainable Seed Company, Chico, CA) (Brassica rapa ssp. chinensis), and Purple Top White Globe turnips (Ferry-Morse Seed, Fulton, KY) (Brassica rapa var. rapa) were used to maintain the beetle colony and as a food source in the experiments.

Seeds were planted in a 72-cell tray containing sterilized soil mix (Fafard super fine germinating mix). Two-week-old seedlings were transplanted into 20- and 15-cm top diameter plastic pots that were used for rearing and experiments, respectively. The pot contained a mixture of soil mix and fertilizer (2.5 ml) (Osmocote Classic® 14N, 14P,

14K). Plants were irrigated daily and fert-irrigated once per week with Miracle Grow

Quick Start® 4N-12P-4K.

Insect colony

Field-collected larvae and adults of M. ochroloma were transported from a commercial farm in Indiantown, FL to the Hayslip Biological Control Research and

Containment Laboratory at the Indian River Research and Education Center in Fort

Pierce, FL. The larvae were confined in a Bug Dorm (60  60  60 cm; Model BD2120-

P, BugDorm Store, Taiwan) with 3 bok choy plants which were replaced as needed. The

Bug Dorm was kept near a window in a laboratory with a constant temperature of approximately 25ºC. Adults were confined in plastic boxes (18  13.5  9 cm) with a screen mesh cloth in the lid for ventilation. Two fresh bok choy leaves were provided 3 times per week. Three rolled Kimwipes® (Model S 12814, Kimberly-Clark, Pleasant

Prairie, WI) were placed in each box to mimic dried leaves that provide small spaces used by M. ochroloma females for oviposition. Rolls with eggs were changed twice per

week and placed in a 695-ml plastic container (Rubbermaid, Atlanta, GA) under laboratory conditions at approximately 25ºC. The plastic boxes, containing the adults, were stored in an environmentally controlled chamber at 21ºC, 60% RH, and 14-h photoperiod. Neonate larvae were transferred daily from the plastic containers to plants in the Bug Dorm. Fourth instars were collected 2 times per week from the Bug Dorm and placed in cylindrical plastic containers with food until they pupated. Newly emerged adults were placed in a separate plastic box; all adults emerging in the same week were placed in the same box.

All the adults used in the experiments were 5-10 d old. Both males and females were surface sterilized by dipping them for 10 s in 1% sodium hypochlorite and washing them with 3 changes of distilled water. The insects were allowed to dry before conducting the experiments.

Fungal formulations

Three commercially available entomopathogenic fungal formulations were tested:

PFR-97 TM 20% WDG (a.i. I. fumosorosea Apopka strain 97 20%, inert ingredients

80%) in the form of desiccation-tolerant granules of blastospores (Certis USA,

Columbia, MD); Botanigard® ES (a.i. spores of B. bassiana strain GHA, inert ingredients

88.7%) (Laverlam International Corporation, Butte, MT); and Met-52® 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 magnetic agitators. For PFR-97, 0.1 g was dissolved in 100 ml of water.

The beaker was placed on a stir plate to homogenize the suspension for 30 min, which was then allowed to settle for 20 min. Only the supernatant containing the blastospores was used in the tests. For Botanigard and Met-52, 100 l of the solution was dissolved

in 100 ml of deionized 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-N01 (NanoEnTek Inc.,

Seoul, Korea). They were then appropriately adjusted to 1-2  106 conidia/ml by adding either deionized water or product.

The viability of the conidia was assessed by pouring 100 µl of each fungal solution onto potato dextrose agar (PDA) in a Petri dish. The Petri dishes were sealed with Parafilm® and incubated at constant 25ºC for 7 d. The number of colony-forming units (CFU) was counted to check the quality of the fungal solutions by comparing the number of CFU obtained with the number of CFU specified in each formulation.

Experiment 1. Insects dipped in fungal solutions

Adult females were topically treated by dipping each insect in 1 of the fungal solutions or distilled water (control). A 500 l quantity of a fungal solution or water was poured into each cell in a 12-well tissue culture cell plate. One female was dipped in the solution for 1-2 min and then placed in a container until it dried.

Once the insect body was completely dry, 1 treated female and 1 untreated male were placed in a Petri dish (9.5 cm in diameter, 1.5 cm in height). Each pair was supplied with a fresh bok choy leaf every 4 d. The petiole of the leaf was inserted into a floral tube containing water to prolong the quality of the leaf. Filter paper (9 cm in diameter) was placed on the bottom of each Petri dish to prevent leaf desiccation. A twisted Kimwipe® (Kimberly-Clark®) was placed inside each Petri dish as a substrate for oviposition. Twelve pairs per treatment were kept in an environmentally controlled chamber set at 20ºC, 60% RH, and 10-h photoperiod. To measure fecundity, eggs laid

during each 96-h period were collected and counted until the female’s death. The collected eggs were transferred to another Petri dish and kept under the same environmental conditions until eclosion, and the number of emergent larvae was counted to determine the effect of each treatment on fertility. Copulation was also noted.

Pairs that did not copulate or lay eggs were excluded from the statistical analysis.

A survivorship and fertility table was constructed for adult females by determining for each age interval (x), the proportion of surviving individuals (lx) and the mean number of female progeny per surviving female (mx). Female progeny was calculated based on the sex ratio (F: M) of 0.8:1 reported by Manrique et al. (2012) when M. ochroloma was reared at 20ºC. The net reproductive rate (Ro) was calculated as the sum of the products lx * mx calculated for each age interval.

Males that died during experiments were replaced with new untreated males.

Dead females were surface sterilized to reduce contamination by saprophytic fungi, by dipping the insect in 70% ethanol for 10 s, briefly rinsing it in distilled water, and dipping it in 1% sodium hypochlorite for 60 s. After washing the female with 3 changes of water, it was placed on moistened filter paper in a Petri dish to promote fungal growth.

Confirmation of insect infection was based on the fungal phenotype unique to each entomopathogenic fungus growing from the insect.

Spore deposition on beetles was tested by dipping 6 insects in each fungal solution as described above. Each treated beetle was placed in an Eppendorf tube with

1 ml of 0.1% Triton X-100 (Sigma-Aldrich, St. Louis, MO) and vortexed for 10 s. A 100

l quantity of the vortexed solution was poured into a Petri dish containing a mixture of

PDA, dodine (1 ml), streptomycin (500 l), and chloramphenicol (500 l). The plate was

sealed with Parafilm® and placed in an environmentally controlled chamber at 25ºC. The number of CFUs was counted 10 d after insects were treated.

Experiment 2. Insects fed leaves applied with fungal solutions

The second experiment was conducted about 6 months after experiment 1.

Individuals of the same colony were used in this experiment. After preparing the fungal solutions as described above, young bok choy leaves were sprayed with approximately

3 ml of the test solutions or with water using 180-ml Nalgene™ (Rochester, NY) spray bottles. Once the leaves dried, 1 female (starved for 24 h) was enclosed with a sprayed leaf in a Petri dish (9.5 cm in diameter, 1.5 cm in height), which was then placed in an environmentally controlled chamber at 20ºC, 60% RH, and 10-h photoperiod. At the end of 48 h, females that had fed on the sprayed leaf were transferred to a 946 ml. cylindrical plastic container (Twin Pak, Québec, Canada) in which a new untreated bok choy leaf, an untreated male, and a piece of paper towel and rolled Kimwipes® were provided. Mesh cloth (11.5 cm in diameter) was used to cover the top of the cylindrical container. A total of 20 pairs of beetles per treatment were placed in the same chamber as above. Seven days later, a second untreated male was placed in the container. The eggs laid during a 72-h period were collected and counted every 3 d until the female’s death. The collected eggs were transferred to another Petri dish and kept under the same environmental conditions until eclosion, and the number of emergent larvae was counted to determine the effect of each treatment on fertility.

The proportion of surviving individuals (lx), the mean number of female progeny per surviving female (mx) and the net reproductive rate (Ro) were calculated as described above.

Dead males during the experiment were replaced with new untreated males.

Copulation was also noted. Pairs that did not copulate or lay eggs were excluded from the statistical analysis. Dead females were surface sterilized as described above.

Sterilized females from each treatment were placed on moistened filter paper in a Petri dish to promote fungal growth. Confirmation of insect infection was based on the fungal phenotype unique to each entomopathogenic fungus growing from the insect.

Spore deposition on the bok choy leaves was recorded by taking 2 leaf discs (12 mm of diameter) using a cork borer. Each disk was placed in a plastic tube with 2 ml of

0.1% Triton X-100 and vortexed for 30 s. A 100 l quantity of the vortexed solution was poured into a Petri dish containing a mixture of PDA, dodine, streptomycin, and chloramphenicol. Three plates per leaf were prepared, sealed with Parafilm®, and placed in an environmentally controlled chamber at 25ºC. The number of CFUs was counted 10 d after leaves were treated.

Statistical analysis

Data on fecundity and fertility were analyzed using an ANOVA. Treatment means were separated by the Tukey – Kramer HSD test using the agricolae package (De

Mendiburu 2015). All the tests were performed in R (R Core Team 2015) with a significance level of 5% for all statistical analyses.

Results

The viability of the conidia in all fungal formulations ranged from 80-100% at the beginning of each experiment. Spores of these 3 fungi are commonly found in the soil and on plants. However, cultivation in PDA-dodine media of sterilized insects showed that the insects did not carry any spores of the entomopathogenic fungi prior to testing.

Experiment 1. Insects dipped in fungal solutions

The average number of eggs laid and egg hatch rate per treatment from dipped beetles are presented in Table 3-1. Fecundity and fertility of M. ochroloma females dipped in the fungal solutions did not differ significantly among treatments (F = 0.49; df

= 3, 29; P = 0.69 for eggs; F = 0.54; df = 3, 29; P = 0.66 for larvae). The lowest total number of eggs oviposited by a female was 49, 47, 72, and 27 when insects were dipped in water, M. anisopliae, I. fumosorosea, and B. bassiana, respectively. The highest total number of eggs was 569, 209, 363, and 496 when applied with water, M. anisopliae, I. fumosorosea, and B. bassiana, respectively. Egg hatch rates did not differ significantly among treatments (F =3.33; df = 3, 29; P > 0.05) (Table 3-1), varying from

56-77%.

Younger females laid more eggs that successfully developed into larvae, and as time passed the number of eggs oviposited and egg hatch rate declined (Figure 3-1 a, d). Two peaks in the mean number of eggs were apparent for the females treated with water, B. bassiana, and I. fumosorosea. The first peak was on day 8, and the second peak on day 20, 24, and 16, respectively (Figure 3-1 a). Only 1 peak was observed on day 12 for M. anisopliae (Figure 3-1 a). The mean number of eggs oviposited per female treated with M. anisopliae dropped 58% in 12 d, whereas the number of eggs from females treated with water and the other 2 fungal formulations decreased gradually over time (Figure 3-1 a).

The proportion of surviving females dipped in M. anisopliae decreased at a faster rate compared with the control (Figure 3-1 b). No females dipped in M. anisopliae survived beyond 36 d after treatment, whereas in the other treatments, the proportion of surviving females reached zero 52 d or more after treatment (Figure 3-1 b). The plot of

the product of age-specific survivorship and fertility (lx*mx) shows that, in general, M. ochroloma is most productive in the first 2 weeks after emergence from the pupa

(Figure 3-1c). Females treated with fungal solutions appeared to lay more eggs in the first week than females in the control treatment, but then a reduction in survival decreased daily egg production over time (Figure 3-1c). This was most pronounced in female beetles treated with M. anisopliae.

The longevity of females dipped in M. anisopliae was significantly shorter than that of females dipped in water and I. fumosorosea (F = 11; df = 3, 40; P < 0.001)

(Figure 3-2 a). Similarly, the longevity of females dipped in B. bassiana was significantly shorter than the longevity of females dipped in water but it was not different from the other 2 fungal treatments (Figure 3-2 a). Fifty percent of the population died 20, 40, 40, and 44 d after treating beetles with M. anisopliae, B. bassiana, I. fumosorosea, and water, respectively.

The net reproductive rate (Ro), or the rate of multiplication in 1 generation, per treatment is included in Table 3-2 Females treated with M. anisopliae had a Ro value

30% lower than the control females. The Ro value of females treated with B. bassiana was only 10% lower than the control Ro, and the Ro of females treated with I. fumosorosea was 7% higher than the control Ro.

Confirmation of infection (Figure 3-3) was difficult to assess in all treatments due to the absence of entomopathogenic fungal growth or to contamination by saprophytic fungi. Only 64, 17, and 9% of dead females were confirmed as infected by B. bassiana,

M. anisopliae, and I. fumosorosea, respectively.

The average number (± SD) of CFUs per dipped insect were 137 ± 105.6 for I. fumosorosea, 411 ± 128.8 for M. anisopliae, and 654 ± 523.2 for B. bassiana. No CFUs of any of the entomopathogenic fungi tested grew from females in the control treatment.

Experiment 2. Insects fed leaves applied with fungal solutions

The average number of eggs laid and egg hatch rate per treatment of beetles fed leaves treated with fungal solutions did not differ among treatments (F = 0.76; df = 3,

55; P = 0.52 for eggs; F = 0.34; df = 3, 55; P = 0.80 for larvae) (Table 3-3). The lowest total number of eggs oviposited by a female was 5, 4, 6, and 11 when leaves were treated with water, M. anisopliae, I. fumosorosea, and B. bassiana, respectively. The highest total number of eggs was 271, 198, 269, and 439 when applied with water, M. anisopliae, I. fumosorosea, and B. bassiana, respectively. Significant differences in egg hatch rate were not detected among treatments (F = 0.50; df = 3, 55; P = 0.68) (Table 3-

3).

The number of eggs oviposited and egg hatch rates generally decreased over time (Figure 3-4 a, d). Two peaks in the mean number of eggs were observed for females treated with water and I. fumosorosea. The first peak occurred on day 9, and the second peak on day 18 and 15, respectively (Figure 3-4 a). Only 1 peak, on day 12, was observed for M. anisopliae- and B. bassiana-treated beetles (Figure 3-4 a). Some peaks were observed after day 42 for all treatments, which can be explained as oviposition by few females that escaped infection. Unlike the experiment 1, in this experiment, the number of eggs laid in all treatments followed a similar pattern and the decrease, in time, was gradual (Figure 3-4 a).

The pattern in the proportion of surviving females over time was similar among treatments (Figure 3-4 b). The plot of the product of age-specific survivorship and

fertility (lx*mx) shows that M. ochroloma females fed treated leaves were most productive in the first 3 weeks after emergence from the pupa (Figure 3-4 c).

The longevity of M. ochroloma females fed leaves differed among the treatments

(F = 2.96; df = 3, 73; P = 0.04). Females that consumed M. anisopliae lived less time than females that consumed B. bassiana (Figure 3-2 b). The longevity of females that consumed M. anisopliae did not differ from that of females that consumed I. fumosorosea or water (Figure 3-2 b). Fifty percent of the population died 33, 39, 36, and

36 d after feeding beetles the leaves treated with M. anisopliae, B. bassiana, I. fumosorosea, and water, respectively.

The net reproductive rate (Ro), or the rate of multiplication in 1 generation, per treatment is included in Table 3-2. Females treated with M. anisopliae had a Ro value

28% lower than the control females. The Ro value of females treated with I. fumosorosea was only 6% lower than the control Ro, and the Ro of females treated with

B. bassiana was 20% higher than the control Ro.

Confirmation of infection by B. bassiana, M. anisopliae, and I. fumosorosea was possible only in 10, 10, and 20% of dead females, respectively. The average number (±

SD) of CFUs per mm2 in each treatment were 0.002 ± 0.004 for I. fumosorosea, 1.000 ±

0.401 for M. anisopliae, and 3.410 ± 0.984 for B. bassiana. A very low number of I. fumosorosea colonies was obtained from the leaf discs, and no CFUs of any entomopathogenic fungus grew from leaves sprayed with water only.

Discussion

This is the first study on the effect of entomopathogenic fungi on the reproductive capacity of M. ochroloma. The results of egg productivity and egg hatch rate indicate that the fecundity and fertility of M. ochroloma are apparently not affected by the

entomopathogenic fungi tested (Tables 3-1 and 3-2). However, the net reproductive rate parameters indicate that M. ochroloma females applied with M. anisopliae might multiply less per generation than untreated beetles or beetles treated with I. fumosorosea and B. bassiana (Table 3-3). A reduction of fecundity caused by these entomopathogenic fungi has been reported in other insects, e.g., the western corn rootworm, Diabrotica virgifera virgifera LeConte (Coleoptera: Chrysomelidae) (Mulock and Chandler 2001), the

Colorado potato beetle (Fargues et al. 1991), the sweetpotato weevil, Cylas puncticollis

Boheman (Coleoptera: Brentidae) (Ondiaka et al. 2008), the legume flower thrips,

Megalurothrips sjostedti Trybom (Thysanoptera: Thripidae) (Ekesi and Maniania 2000), the German cockroach (Quesada-Moraga et al. 2004), and the Mediterranean fruit fly

(Castillo et al. 2000). In my study, 2 application techniques were tested, insect dipping and leaf spraying, but no effect was found on the number of eggs oviposited and hatch rate of M. ochroloma regardless of the method used. Two studies have shown that B. bassiana (BotaniGard formulation) and I. fumosorosea (PFR-97 formulation) have low virulence in M. ochroloma. Balusu and Fadamiro (2013) observed that BotaniGard was more efficacious against M. ochroloma larvae (~50% mortality), whereas very low efficacy was observed against adults. Similarly, Montemayor et al. (2016) found that I. fumosorosea caused reduced growth, unsuccessful molting, and subsequent mortality of first and third instar of M. ochroloma, whereas no effect was observed in adults.

There are no reports of the virulence of M. anisopliae against the immatures stages and adults of M. ochroloma.

The poor efficacy of these fungal formulations to affect the reproductive capacity of female M. ochroloma could be related to various factors. One cause contributing to

the lack of infectivity of adults of M. ochroloma is the presence of a hard cuticle in adult beetles (Balusu and Fadamiro 2013, Montemayor et al. 2016). The cuticle is the first line of defense that insects have against pathogens and other natural enemies (Nation

2008), and its composition varies among the developmental stages of the insect. For instance, larvae of the red flour beetle, Tribolium castaneum Herbst (Coleoptera:

Tenebronidae), exhibit a higher percentage (80%) of a simpler blend of mid-chain aliphatic hydrocarbons (HC) (source of carbon for conidial germination) than the adults, whose cuticle is composed of only 40% HC (Pedrini et al. 2007). Additionally, HC content and the secretion of antimicrobial compounds (cuticular lipids, small molecule toxins, and proteins) 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, Otti et al. 2014). For example, T. castaneum secretes into their immediate environment quinones that have antimicrobial properties (Joop et al. 2014).

The southern green stink bug, Nezara viridula L (Hemiptera: Pentatomidae), produces aldehydes that can inhibit conidial germination and germ tube development of M. anisopliae (Sosa-Gomez et al. 1997). Leaf beetle larvae of Chrysomela spp. and

Phratora vitellinae (Linnaeus) (both Coleoptera: Chrysomelidae) produce secretions of which the main component is salicylaldehyde that strongly inhibits germination of M. anisopliae (Gross et al. 2002, 2008). Besides the cuticle and external secretions, insects count on humoral and cellular defense mechanisms to prevent infection by viruses, bacteria, fungi, and other parasites (Hajek and St. Leger 1994, Nation 2008,

Otti et al. 2014).

Even though, the cuticle provides a good defense against many parasites, spores can penetrate through intersegmental membranes. In heavily sclerotized insects, fungal penetration occurs via the spiracles or arthrodial membranes (Hajek and St.

Leger 1994). The results observed in Experiment 2 when insects were fed treated leaves indicate that the spores are not able to germinate or penetrate the digestive membrane of the insects or the insects did not ingest sufficient spores to become infected. This may explain differences between dipped and fed beetles.

Some studies have also found that sublethal effects of these entomopathogenic fungi on fecundity and fertility were apparent in adults that emerged from surviving larvae that had been treated with entomopathogenic fungi. Adults of M. sjostedti that survived infection by M. anisopliae as larvae showed a significant reduction in fecundity, fertility, and longevity (Ekesi and Maniania 2000). Fertility and fecundity of adults of the

Colorado potato beetle were reduced when treated in the fourth instar with B. bassiana and reared at 22ºC (Fargues et al. 1991). Sikura et al. (1972) reported histological and cytological injuries to the ovaries of Colorado potato beetles that survived treatment with

B. bassiana during the larval stage. In my study, the adult stage received the treatment, therefore, it is necessary to determine if sublethal concentrations of entomopathogenic fungi applied to the larval stage would have an effect on fecundity and fertility.

The efficacy of entomopathogenic fungi might also be affected by the strain of the fungi used. Infection levels, germination rates, and optimum developmental temperature can vary among fungal isolates (Shah and Pell 2003, Cherry et al. 2005).

Some species of insects are sensitive only to entomopathogenic fungi isolated from individuals belonging to their species or until the insect of interest has been infected

(under artificial conditions) and the fungus has been isolated from it (Zimmermann

2007a, Ortiz-Urquiza and Keyhani 2013). For example, M. anisopliae strain F52 tested against 9 species of Scarabaeidae showed that most insects were susceptible only to fungal strains isolated from the same species (Ferron et al. 1972). Beauveria bassiana isolates with distinct origins might or may not be virulent to target pests (Zimmermann

2007b). Six isolates of B. bassiana were able to infect the Russian wheat aphid,

Diuraphis noxia Mordvilko (Hemiptera: Aphididae), but their virulence varied with the isolate regardless of the source of isolation (hemipteran or non-hemipteran host) (Feng and Johnson 1990). Fargues and Robert (1983) reported that 2 strains of M. anisopliae increased their infection potential factor from 10 to 100 after a single in vivo passage through the host. Therefore, it is necessary to screen the virulence of different strains against M. ochroloma to select the most virulent or to increase the infection factor.

Metarhizium anisopliae significantly reduced the longevity of dipped females compared to females dipped in water only. Females treated with water lived, on average, 27 d longer than females treated with M. anisopliae. Similarly, Ekesi and

Maniania (2000) reported M. sjostedti longevity was 6–9 d shorter when treated with M. anisopliae. However, the cause of death in my study cannot be attributed to M. anisopliae with certainty due to the inability of confirming the presence of the entomopathogenic fungi. When females ate leaves applied with M. anisopliae, no significant differences in longevity, compared to the control, were observed but there was a tendency of a shorter longevity for females treated with M. anisopliae. Quesada-

Moranga et al. (2004) attributed a reduction in the number of oothecae laid by B. germanica treated with M. anisopliae to a reduction in adult longevity. However, a

reduction in longevity of M. ochroloma did not affect the total number of eggs oviposited because females treated with M. anisopliae had the highest oviposition rates during the first 2 weeks of the experiment before they were killed by the fungus (Figure 3-1 a).

Oviposition in the first 2 weeks was 2 times higher than in the control treatment, which compensated for the reduction in longevity, thus causing no differences in fecundity among the treatments.

The low mortality caused by M. anisopliae strain F52, B. bassiana strain GHA, and I. fumosorosea strain 97 (Balusu and Fadamiro 2013) and the lack of effect on fecundity or fertility of M. ochroloma adults do not suggest that this pest will be well managed using commercially available entomopathogenic fungi. Therefore, the integration with other methods of control is necessary. However, it is important to point out that oviposition rate declined more sharply in females treated with M. anisopliae and they died at a faster rate than in other treatments (Figure 3-1 a-c), which likely led to the notably lower Ro for Met females. This indicates more potential with M. anisopliae versus the other fungi, and this potential should be investigated with more potent species-specific strains. Additionally, the potential effect of M. anisopliae on the immature stages and the effect on fecundity and fertility of M. ochroloma adults after being treated in the larval stage with sublethal doses of these entomopathogenic fungi should be further tested. Spraying is the most common method of application of these fungal formulation under field conditions. Therefore, additional tests spraying insects instead of dipping them in fungal solution should be done.

Table 3-1. Microtheca ochroloma fecundity and fertility after dipping females in three entomopathogenic fungal formulations. Treatment n Eggs laid Emergent larvae % emergence Control 9 200.2 ± 51.0 116.2 ± 37.6 55.6 ± 6.8 Metarhizium anisopliae 8 139.1 ± 36.3 111.9 ± 31.3 76.5 ± 5.6 Beauveria bassiana 7 178.1 ± 64.8 108.3 ± 36.5 62.8 ± 5.4 Isaria fumosorosea 9 215.1 ± 37.5 160.0 ± 28.2 75.0 ± 3.8 Data are means ± SEM. Means within columns are not significantly different (P = 0.69 for eggs and P = 0.66 for larvae).

Table 3-2. Net reproductive rate per treatment after dipping the insects or spraying leaves with fungal solutions. Treatments Ro (dipped insects) Ro (sprayed leaves) Control 79.78 37.20 Metarhizium anisopliae 55.65 26.58 Beauveria bassiana 71.71 46.41 Isaria fumosorosea 86.09 34.83

Table 3-3. Sublethal effect of three entomopathogenic fungal formulations on Microtheca ochroloma fecundity and fertility after feeding on treated leaves. Treatments n Eggs laid Emergent larvae % emergence Control 16 81.9 ± 19.6 45.7 ± 14.3 45.7 ± 14.3 Metarhizium anisopliae 11 62.5 ± 19.7 39.0 ± 14.1 39.0 ± 14.2 Beauveria bassiana 17 111.5 ± 27.8 63.5 ± 23.0 63.5 ± 23.0 Isaria fumosorosea 15 84.4 ± 18.6 51.5 ± 13.5 51.5 ± 13.5 Data are means ± SEM. Means within columns are not significantly different (P = 0.52 for eggs and P = 0.80 for larvae).

Figure 3-1. Changes in reproduction and survival of Microtheca ochroloma when dipped in fungal solutions or water. (a) Mean number of eggs oviposited per female during lifetime, (b) proportion of surviving females, (c) adult age- specific survivorship*fertility, and (d) proportion of egg emergence.

Figure 3-2. Longevity of Microtheca ochroloma females treated with three fungal formulations. (a) Insects dipped in the solution and (b) insects fed with leaves sprayed with the solution. Treatments with the same letter in each graph are not significantly different (P < 0.001 for dipped insects and P = 0.04 for insects fed treated leaves). Tukey – Kramer HSD test was used to compare treatment means.

Figure 3-3. Entomopathogenic fungal growth on Microtheca ochroloma females: (a) Beauveria bassiana, (b) Metarhizium anisopliae, and (c) Isaria fumosorosea.

Figure 3-4. Changes in reproduction and survival of Microtheca ochroloma when insects were fed leaves sprayed with fungal solutions or water. (a) Mean number of eggs oviposited per female during lifetime, (b) proportion of surviving females, (c) adult age-specific survivorship*fertility, and (d) proportion of egg emergence.

CHAPTER 4 EFFECT OF NON-CRUCIFER EXTRACTS ON HERVIBORY BY ADULTS OF MICROTHECA OCHROLOMA STÅL (COLEOPTERA: CHRYSOMELIDAE)

Crucifer crops in the southern U.S. are threatened by the presence of the introduced beetle Microtheca ochroloma Stål (Chamberlin and Tippins 1948, Haeussler

1951, Rohwer et al. 1953, Staines 1999, Guillebeau 2001, Rhodes and Liburd 2014).

This cool-season pest is active from October to late April in Florida, which coincides with the crucifer production season in the state (Bowers 2003). During summer months, adults enter aestivation (Chamberlin and Tippins 1948, Oliver and Chapin 1983).

Microtheca ochroloma has multiple generations during its active season (Ameen and

Story 1997a). Larvae and adults feed exclusively on crucifers, chewing holes in the leaves (Jolivet 1950). Among crucifers, M. ochroloma has a strong preference for turnip

(Chamberlin and Tippins 1948, Ameen and Story 1997c, Balusu and Fadamiro 2011), mustard (Ameen and Story 1997c), and napa cabbage (Balusu and Fadamiro 2011) over cabbage and collards.

The use of synthetic, broad-spectrum insecticides has been the primary and most effective tactic to manage M. ochroloma (Webb 2013, Rhodes and Liburd 2014).

However, concerns about pesticide resistance, human and environmental safety, and insecticide restrictions in organic farming have intensified the efforts to find alternative control methods. Trap cropping using turnips, which takes advantage of M. ochroloma’s host preference, was evaluated by Balusu et al. (2015) as a feasible method to manage this beetle. Planting turnips at the perimeter of the main crop to lure and prevent insect colonization reduced the number of adults and larvae of M. ochroloma and their damage. However, this tactic was only successful when the main crop was a less attractive crucifer (e.g., cabbage and collard). When mustard and napa cabbage, which

are equal to turnip in attractiveness, were the main crops, trap cropping using turnips was not sufficient to reduce the damage caused by M. ochroloma. Balusu et al. (2015) suggested that planting turnips at the periphery of mustard and napa cabbage should not be used as a stand-alone tactic to control M. ochroloma.

The efficacy of trap cropping might be increased by reducing the attractiveness of the main crop. Synthetic repellent and non-host volatiles have been successfully used to drive away insects and reduce the attraction and palatability of the crop of interest (Cook et al. 2006). For example, leaves of sassafras (Sassafras albidum Nutt.) and soybeans (Glycine max L.) treated with extracts from seeds of the neem tree

(Azadirachta indica A. Juss) remained untouched or suffered slight damage by the

Japanese beetle, Popillia japonica Newman (Coleoptera: Scarabeidae) in both laboratory and field tests (Ladd et al. 1978). Similarly, extracts from 4 plants of the

Lamiaceae family, marjoram (Origanum majorana L.), sage (Salvia officinalis L.), hyssop (Hyssopus officinalis L.), and horehound (Marrubium vulgare L.), applied on potato leaves had a strong repellent and antifeedant effect on the feeding behavior of the Colorado potato beetle, Leptinotarsa decemlineata Say (Coleoptera:

Chrysomelidae) adults (Pavela 2004). The pollen beetle, Meligethes aeneus Fabricius

(Coleoptera: Nitidulidae), which damages oilseed rape, Brassica napus L., was strongly repelled by lavender (Lavendula angustifolia Miller) essential oil (Mauchline et al. 2005).

Papachristos and Stamopoulos (2002) reported that most of the thirteen essential oils evaluated against Acanthoscelides obtectus Say (Coleoptera: Chrysomelidae) caused repellency, reduced fecundity, and negatively affected egg hatchability and neonate larval survival. The use of non-host extracts that have shown good results against

various pests has not yet been evaluated against M. ochroloma. Therefore, the purpose of my study was to evaluate the effect of 10 non-crucifer essential oils on herbivory by adults of M. ochroloma.

Materials and Methods

Plants and insects

Organic Baby Bok choy (Sustainable Seed Company, Chico, CA) (Brassica rapa ssp. chinensis) and Purple Top White Globe turnips (Ferry-Morse Seed, Fulton, KY)

(Brassica rapa var. rapa) were used to maintain the beetle colony and as a food source in the experiments. Plants were grown as described in Chapter 3.

Field-collected larvae and adults of M. ochroloma were used to establish a laboratory colony as described in Chapter 3. All the adults used in the experiments were laboratory-reared and 5-10 d old as adults. The beetles were starved for 24 h prior to experimentation. During the starvation period, only distilled water was provided via a saturated cotton ball.

Non-crucifer essential oils

Essential oils of the following non-crucifer plant extracts were evaluated: marjoram, clary sage (Salvia sclarea L.), hyssop, thyme (Thymus vulgaris L.), oregano

(Origanum vulgare L.), black pepper (Piper nigrum L.), neem, lavender, eucalyptus

(Eucalyptus citriodora Hooker), and garlic (Allium sativum L.). The essential oils were purchased from Bulk Apothecary (Streetsboro, OH).

The commercially available, organically accepted product AzaMax® (Parrys

America Inc., Irving, TX) was also used in the no-choice and paired choice tests. The highest recommended concentration, 1.70% vol/vol, was prepared by diluting the product with distilled water. The same concentration was used for all tests.

Experiment 1. No-choice test

The effect of the non-crucifer essential oils on leaf consumption by adults of M. ochroloma was evaluated by providing a bok choy leaf sprayed with 1 of the non- crucifer essential oils. For the no-choice test, a 2% concentration of each essential oil was prepared by dilution in 0.1% Tween 20 (Sigma-Aldrich, St. Louis, MO). The solutions were vortexed for 10 seconds before application. A bok choy leaf, trimmed to an area of 26 ± 2 cm2, was sprayed with 1 ml of 1 of the test solutions or 0.1% Tween

20 (control) using a 2-oz plastic perfume atomizer (Sample Container Store,

Champaign, IL). The leaves were allowed to dry for an hour before starting the experiment. One treated trimmed leaf and a 10-d-old adult (mixed sexes) were contained in a Petri dish (10 cm in diameter) with white filter paper placed at the bottom to prevent water condensation on the walls. The leaf petiole was inserted in a floral tube containing water to prevent leaf desiccation. The number of repetitions varied from 11 to

13 per treatment. The area consumed by the beetles within 24 h was measured by photographing the leaf and processing the image through the software ImageJ

(Schneider et al. 2012). The percentage of area consumed was calculated as consumed area / total leaf area x 100.

Based on the results obtained from the non-choice test of the 10 non-crucifer essential oils, neem oil was chosen for follow-up experimentation. There are several commercially available products that contain azadirachtin, which is considered the main component of neem seeds responsible for the antifeedant and toxic effect in insects

(Mordue and Nisbet 2000). The formulation AzaMax was used instead of the neem essential oil since AzaMax is low-cost, user-friendly, and accessible. Bok choy leaves were sprayed with either AzaMax or distilled water and offered to M. ochroloma adults

following the methodology described above. A trial with 21 repetitions per treatment was conducted.

Experiment 2. Paired choice test

In the paired choice tests, bok choy leaves were sprayed with 1 ml of AzaMax or distilled water. In the AzaMax treatment, 2 bok choy leaves, 1 applied with AzaMax and the other applied with water, were placed in a 350-ml rectangular plastic container

(Rosti Mepal, The Netherlands). In a control treatment, 2 bok choy leaves applied with water were offered to a pair of beetles to determine if differences in consumption were related to feeding behavior of the beetles when 2 bok choy leaves were available instead of the effect of the repellent. An adult pair (male and female) was placed in each container and allowed to feed for 48 h. At the end of 48 h, the leaves were photographed and the area consumed was measured using the software ImageJ.

Twenty pairs per choice test per treatment were evaluated.

Considering that the goal of using non-crucifer extracts would be to reduce the attractiveness of the main crop when turnip is used as a trap crop, a second paired choice test was performed in which adults were offered a treated bok choy leaf (sprayed with AzaMax) and an untreated turnip leaf (sprayed with water). In the control, untreated bok choy and turnip leaves were offered to a M. ochroloma pair. At the end of 48 h, the leaves were photographed, and the area consumed was measured using the software

ImageJ. Twenty pairs per choice test per treatment were evaluated.

Experiment 3. Whole plant paired choice test

Two 2-week-old bok choy plants were applied with AzaMax or distilled water until the leaves were completely wet. The plants were allowed to dry and then placed in a rectangular mesh cage (25  25  45 cm). The plants were approximately 30 cm apart.

Twenty adults (10 males and 10 females) were placed in the middle of the cage. The number of insects on each plant was counted once a day for 2 d. At the end of 48 h, all the leaves of each plant were photographed and the consumed area was measured using the software ImageJ. Two trials, the first with 10 cages and the second with 5 cages, were conducted. For the second trial, the plants were taken out of the cage and the leaves were photographed 48 h after starting the experiment. A Petri dish was used to flat the leaves without removing them from the plants. The plants were returned to their respective cage and the insects were allowed to feed for 5 more days. At the end of 7 d, the leaves were removed from the plant and photographed again. The leaf area consumed at 48 and 168 h was measured using the software ImageJ.

Experiment 4. Starvation time

Due to high variation in leaf consumption in the control treatments, a test was conducted to quantify the variation in consumption when adults were starved for different time intervals. Four starvation times, 12, 24, 36, and 48 h, were evaluated. The insects were provided with distilled water via moistened cotton during the starvation period. A 26-cm2 piece of bok choy leaf was offered to a pair of adults for 48 h. At the end of the feeding period, the consumed leaf area was measured using the software

ImageJ. Fourteen pairs per starvation time were tested.

Statistical analysis

Data on the percentage of consumed leaf area in the no-choice test of the 10 non-crucifer essential oils were analyzed using ANOVA. Non-choice test testing of

AzaMax against the control was analyzed using the nonparametric Wilcoxon rank-sum test. The paired choice tests data were analyzed using a paired t-test. Repeated measures ANOVA was used to analyze data in the second trial of the whole plant

paired choice test. Starvation time data were subjected to linear regression analysis.

Data transformation of the percentage of consumed leaf area using arcsine or probit transformation was done when needed to meet the parametric test assumptions.

Transformed data were back transformed to be presented in tables and figures.

Treatment means were separated by the Tukey–Kramer HSD test using the agricolae package in R (De Mendiburu 2015). All tests were performed in R (R Core Team 2015) with a significance level of 5% for all statistical analyses.

Results

The initial no-choice bioassay showed that herbivory can be affected by the presence of non-crucifer essential oils on the leaves. Significant differences (F =4.90; df

= 9, 110; P < 0.001) in leaf consumption were found among the 10 non-crucifer essential oils tested. Leaf area consumed was reduced when leaves were sprayed with neem, hyssop, and thyme essential oils compared to leaves sprayed with garlic and lavender (Figure 4.1). The leaf area consumed in the control was not significantly different from the other treatments. However, herbivory tended to numerically decrease when leaves were applied with neem, hyssop, and thyme.

Leaf area consumed by adults was significantly reduced (W = 88; P < 0.001) when the leaves were treated with AzaMax compared to untreated leaves. The percentage of feeding on leaves sprayed with AzaMax was 54% less compared to the control.

A significantly higher (t = 3.07; df = 19; P < 0.001) percentage of foliar area of the untreated leaf was consumed by adults when they were provided with 2 bok choy leaves, 1 treated with AzaMax and the other untreated (Table 4.1). The percentage of feeding reduction was, on average, 68% when AzaMax was sprayed on the leaves.

When 2 untreated bok choy leaves were offered to the beetles, no significant difference

(t =-1.66; df = 19; P = 0.11) between the percentages of leaf area consumed per leaf was found.

Similarly, when adults of M. ochroloma were offered a treated bok choy leaf and an untreated turnip leaf, the insects consumed a significantly higher percentage of leaf area of the untreated turnip leaf than the treated bok choy leaf (t =3.56; df = 19; P =

0.002) (Table 4.1). The percentage of feeding reduction was, on average, 77% when the AzaMax was sprayed on the bok choy leaves. Beetles given a choice of untreated bok choy and turnip leaves did not show a preference for either species (t =1.58; df =

19; P = 0.13).

The paired choice test using whole plants did not show differences in percentage of leaf area consumed in the first trial (t = 0.78; df = 9; P = 0.46). However, the mean percentage of consumption was slightly lower on treated plants (0.21%) than on untreated plants (0.33%). In the second trial, significant differences were only detected among treatments on day 7 (F = 9.16; df = 1, 15; P = 0.008) and, in the control treatment, significant differences were found among sampling dates (F = 8.06; df = 1,

15; P = 0.01). The beetles consumed more leaf tissue in plants treated with water than plants treated with AzaMax (Figure 4.2). No significant difference (t = -0.81; df = 1, 18; P

= 0.43) was detected between the mean number of insects on treated (3.4 ± 2.2) and untreated (4.5 ± 3.7) bok choy plants at the end of day 1. Neither were there significant differences (t = -0.36; df = 1, 18; P = 0.72) between the mean number of insects on treated (2.2 ± 1.7) and untreated (2.5 ± 2.0) bok choy plants at the end of day 2. The number of dead insects at 48 h varied from 1 to 3 per cage for an average of 1.5 dead

insects per cage. The number of dead insects at 168 h varied from 2 to 11 per cage for an average of 6.8 dead insects per cage.

The percentage of consumed leaf area varied significantly (F = 3.57; df = 3, 52, P

= 0.02) among starvation times. The percentage of leaf area consumed by M. ochroloma adults decreased with increasing starvation time as shown in Figure 4-3.

Consumed leaf area after a 12-h starvation was significantly higher than at 48 h, while it was not different from the area consumed by beetles starved for 24 and 36 h.

Discussion

This is the first study on the effect of non-crucifer extracts on the feeding behavior of adults of M. ochroloma. From the 10 non-crucifer essential oils tested, high concentrations of origanum, sage, and eucalyptus caused phytotoxicity to the leaves.

Among 10 non-crucifer essential oils that have shown repellent and antifeedant properties against other insect pests, neem, hyssop, and thyme might potentially reduce leaf damage by M. ochroloma (Figure 4.1). Extracts of these plants have been reported to deter feeding by other beetle pests. For example, Pavela (2004) reported a strong repellent and antifeedant effect on adults of the Colorado potato beetle generated by essential oils extracted from hyssop and thyme plants. Unlike the Colorado potato beetle, the antifeedant effect on adults of M. ochroloma was not strong enough to cause significant differences in consumption compared to the control treatment. However,

Pavela (2004) reported that the effect caused by hyssop and thyme extracts changed with time. The antifeedant and repellent effect of hyssop declined while the effect of thyme was better after 24 h (Pavela 2004). Therefore, the inconsistent effect of the essential oils through time could have affected the results in my study. Once the effect of the essential oils decreased, insects could have resumed feeding or, in the case of

thyme, more time would have been needed to see a higher reduction in feeding by adults of M. ochroloma.

Neem has long been recognized for its antifeedant and toxic properties against insects (Mordue and Nisbet 2000). Neem has been used successfully to manage various insect pests because it not only deters feeding, but it also affects insect growth, molting, and fecundity (Mordue and Nisbet 2000). Neem deters feeding by other chrysomelids such as the cucurbit beetle, Diabrotica speciosa (Germar) (Ventura and

Ito 2000), the Colorado potato beetle (Zehnder and Warthen 1988), the striped cucumber beetle, Acalymma vittatum (Fabricius), and the spotted cucumber beetle,

Diabrotica undecimpunctata howardi (Barber). Neem’s active ingredient that has been isolated and is considered responsible for the adverse effects on insects is the secondary compound azadirachtin (Isman et al. 1990). The neem product used in my study, in the initial no-choice test, did not significantly reduce the percentage of leaf area consumed by beetles compared to the control. A possible factor could be differences in the amount of azadirachtin in the neem essential oil used in this test compared to the neem extracts used in other studies. Isman et al. (1990) reported that azadirachtin content varies greatly among neem extracts, and the effect of neem oils is correlated with their azadirachtin content, the higher the concentration of azadirachtin, the stronger is the effect. Composition and concentration of plant extracts within the same species can vary due to chemotypes, plant stage, plant tissue used, cultivation practices, environmental conditions, and extraction methods (Regnault-Roger et al.

2011). This high variation in plant extract components has become a challenge to manufacturing standardized products, which is necessary for regulatory and marketing

purposes (Regnault-Roger et al. 2011). Companies have opted for isolating azadirachtin from neem extracts to produce commercial products with a determined concentration of this secondary compound. In this study, AzaMax, which contains 1.2% azadirachtin, caused a significant reduction in the percentage of leaf area consumed compared to the control in a no-choice test, whereas the neem essential oil did not. Therefore, as suggested by Isman et al. (1990), the azadirachtin content of neem extracts should be quantified if its intended use is pest management.

In the paired choice tests, in which the insects were presented with an AzaMax- treated and an untreated leaf, M. ochroloma adults consumed less of the AzaMax- treated leaves. When the beetles were provided with 2 untreated bok choy leaves, they ate similar amounts of leaf area from both leaves offered. This demonstrates that the beetle’s decreased feeding on AzaMax-treated leaves was not random. A similar trend was observed in the paired choice test offering a treated bok choy leaf and an untreated turnip leaf.

Trap cropping using turnip has been successful to reduce the number of insects and the damage caused by M. ochroloma on cabbage crops (Balusu et al. 2015).

However, turnip as a trap crop was not successful in preventing insect damage on equally attractive crops such as mustard (Balusu et al. 2015). In my study, when adults of M. ochroloma were offered untreated leaves of turnip and bok choy, the insects did not show a preference for either species since they were both equally attractive in this assay. However, when the beetles were offered an AzaMax-treated bok choy leaf and an untreated turnip leaf, they avoided the treated leaf and consumed a significantly higher percentage of leaf area of the untreated turnip leaf. In a more complex cage

setting offering insects an AzaMax-treated and an untreated bok choy plant, no significant differences in the percent of leaf area consumed was observed in the first 48 h, whereas significant differences were observed 7 d after starting the experiment. The herbivory on the AzaMax-treated plants was lower than that on untreated plants (Figure

4-2). Differences in damage between evaluation dates can be due to initial testing of the leaves by the beetles. The product on the leaves might only be perceived by the beetles after contact. Once the beetles tasted the plants, they might have avoided them and concentrated feeding in the control plant. Olfactometer tests are needed to evaluate the repellent effect of AzaMax on M. ochroloma.

Unlike the paired choice tests in the small containers where only a pair was tested, in the whole plant paired choice test a group of 20 beetles was introduced in each cage. Testing groups of insects can introduce other variables that can influence the location and feeding behavior of an insect. For example, male flea beetles,

Phyllotreta cruciferae Goeze (Coleoptera: Chrysomelidae), produce an aggregation pheromone that attracts other beetles and affects host-plant location (Peng and Weiss

1992). Similarly, Dickens et al. (2002) reported a male-produced aggregation pheromone in the Colorado potato beetle, and Smyth and Hoffmann (2003) found that males of the striped cucumber beetle also produce an aggregation pheromone that facilitates host location at the beginning of the colonization process. Considering the observation that similar numbers of insects were found on treated and untreated plants, azadirachtin might only be detected by the insect after feeding on the treated plant. In that case, an aggregation pheromone could attract more beetles to the treated plant before they are affected by the antifeedant properties of azadirachtin. There are no

studies about possible aggregation pheromones present in M. ochroloma. More trials with a higher number of replicated treatments may be needed to determine the effect of

AzaMax when applied to whole plants and offered to a group of adult M. ochroloma.

High variation in consumption in the control treatments of the no-choice tests and the lack of a stronger effect of non-crucifer essential oils, which are known to have an antifeedant effect on other pests, prompted me to test the effect of various starvation times on the consumption of adults of M. ochroloma. The percentage of leaf area consumption in 48 h was similar when beetles were starved for 12, 24, or 36 h, but leaf consumption was significantly reduced when beetles were starved for 48 h. The coefficient of determination (R2) was low for the linear regression as a result of a high variation within treatments and the narrow range of starvation times (Figure 4-2). Most studies testing consumption rates or feeding behavior of insects require putting the insect through a period of starvation to promote an insect response during experimentation. Besides increasing the responsiveness of the insect to the host, starvation can also affect insect searching behavior, locomotion rate, oriented movement, and acceptance of a previously unacceptable host (Bell 1990). My test showed that M. ochroloma feeding behavior is negatively affected by the amount of time the beetle is starved. However, the insects in my tests of plant essential oils were deprived for only 24 h, so other factors might have affected consumption in the control treatments.

Based on my results, applications of AzaMax might be integrated with trap cropping to reduce insect damage when a crucifer crop that is equally attractive as turnip is grown as the main crop. However, further studies using whole plants and field

testing are needed to assess the potential of integrating AzaMax, as an antifeedant, with trap cropping using turnip.

Table 4-1. Percentage of leaf area consumed by adults of Microtheca ochroloma when insects were offered an AzaMax-treated leaf and an untreated leaf in paired choice tests. % of consumed leaf area Choice test Treated bok choy Untreated bok choy Untreated turnip Treated vs untreated bok 0.72 ± 0.21 a 2.04 ± 0.39 b NT choy Treated bok choy vs 0.48 ± 0.13 a NT 2.22 ± 0.43 b untreated turnip NT = not tested. Data are means ± SEM. Means within rows with the same letters are significantly different (P < 0.001 for treated vs untreated bok choy and P = 0.002 for treated bok choy vs untreated turnip). Tukey – Kramer HSD test was used to compare treatment means.

Figure 4-1. Percentage of leaf area consumed by Microtheca ochroloma when offered a bok choy leaf sprayed with one non-crucifer essential oil or water (control) in a no-choice test. Bars are means ± SEM. Bars with the same letter are not significantly different (P < 0.001). Tukey – Kramer HSD test was used to compare treatment means.

Figure 4-2. Percentage of leaf area consumed by Microtheca ochroloma when offered an untreated and an AzaMax-treated bok choy plant in a paired choice test. Bars are means ± SEM. Bars with the same lowercase letter are not significant different (P = 0.46 for 48h evaluation and P = 0.008 for 168h evaluation). Tukey – Kramer HSD test was used to compare treatment means.

Figure 4-3. Percentage of leaf area consumed by Microtheca ochroloma after being deprived of food for various periods of time. Treatments with the same letter are not significantly different (P = 0.02). Tukey–Kramer HSD test was used to compare treatment means.

CHAPTER 5 EFFECT OF HIGH TEMPERATURES AND LONG-DAY PHOTOPERIOD ON THE REPRODUCTIVE BEHAVIOR OF ADULTS OF MICROTHECA OCHROLOMA STÅL (COLEOPTERA: CHRYSOMELIDAE)

Cruciferous crops in the southeastern U.S. are being attacked by the yellowmargined leaf beetle, Microtheca ochroloma Stål (Coleoptera: Chrysomelidae), since its unintended introduction from Argentina (Chamberlin and Tippins 1948, Jolivet

1950). Larvae and adults of this beetle chew holes in the leaves of the plant, causing complete defoliation when the beetles are present in high numbers (Chamberlin and

Tippins 1948, Jolivet 1950). This beetle is strongly attracted to economically important crops such as turnips, mustard, napa cabbage, and Asian leafy greens (Ameen and

Story 1997, Bowers 2003, Balusu and Fadamiro 2011). It is a cool-season pest

(Chamberlin and Tippins 1948, Rohwer et al. 1953, Oliver and Chapin 1983). In Florida, adult beetles start appearing in the field in October, at the beginning of the cruciferous crop season, and by early spring their population reaches high numbers that cause economic losses (Rohwer et al. 1953, Oliver and Chapin 1983, Bowers 2003). At the end of spring and in summer, the beetles become inactive (Chamberlin and Tippins

1948, Rohwer et al. 1953, Oliver and Chapin 1983, Bowers 2003). Environmental conditions during hot months are detrimental to the development of the larvae

(Manrique et al. 2012). Therefore, the beetles require a strategy to survive long periods of time without food and under conditions that prevent reproduction. In more northern latitudes (e.g., Alabama), M. ochroloma overwinters as an adult from late December to mid-March (Balusu et al. 2015) but winter dormancy has not been observed in Florida due to the mild weather.

Various mechanisms have been developed by insects to cope with environmental changes that allow them to synchronize with favorable conditions and survive during unfavorable periods (Tauber and Tauber 1976, Tauber et al. 1986). The responses of insects to seasonal changes might vary within a range of dormancy types limited by 2 seasonal dormancy responses: quiescence or diapause (Tauber and

Tauber 1976, Danks 1987). Quiescence is a state of arrested metabolism in which the insect responds rapidly to changes in environmental conditions, and if favorable conditions are restored, growth and reproduction resume in a short period of time

(Tauber et al. 1986, Danks 1987). On the other hand, diapause, which also involves a low metabolic activity, occurs as a response to environmental stimuli that precede unfavorable conditions; these stimuli are perceived by earlier developmental stages than those that undergo diapause (Tauber et al. 1986, Danks 1987). Cues to seasonal changes are provided by many factors such as photoperiod, temperature, humidity, and biotic factors (Tauber et al. 1986).

Microtheca ochroloma adults are considered the oversummering stage, and anecdotal information suggests that the beetles enter a period of dormancy in refuges surrounding the crop areas (Oliver and Chapin 1983, Bowers 2003, Balusu et al. 2015).

However, crucial information about this beetle’s dormancy is unknown. There is no published information about the environmental factors that induce dormancy or the type of dormant strategy that M. ochroloma experiences. Information about M. ochroloma dormancy is valuable to increase the effectiveness of control tactics. For example, the first appearance of the insects can be predicted or detected in a timely manner to apply control methods before the beetles cause significant damage.

The goal of this study is to provide information on the effect of high temperatures and long photoperiod on the consumption and reproductive behavior of adults of M. ochroloma. This information may serve as a starting point for future research on M. ochroloma dormancy.

Materials and Methods

Plants and insects

Organic Baby Bok choy (Sustainable Seed Company, Chico, CA) (B. rapa ssp. chinensis) was used to maintain the beetle colony and as a food source in the experiments. Plants were grown as described in Chapter 3.

Field-collected larvae and adults of M. ochroloma were used to establish a laboratory colony as described in Chapter 3. Newly emerged adults (24–48 h old) were kept in a plastic container until they started copulating. Copulating pairs were isolated in a separate container and kept under the same conditions (21ºC, 60% RH, and 10L: 14D photoperiod) until the female started laying eggs.

Experimental design

Combined regimens of 4 temperatures (21, 24, 27, or 30ºC) with 2 photoperiods

(10h or 14-h photoperiod) were tested. Relative humidity was consistently 60% for all treatments. Approximately 24 h after first oviposition, the container holding a copulating pair was moved from the rearing chamber (21ºC and 10-h photoperiod) to an environmentally controlled chamber set at 1 of the 8 temperatures/photoperiod combination treatments. Between 4 and 5 mating pairs (replicates) were subjected to each treatment.

The insects were fed a fresh bok choy leaf every 3 d, and leaf consumption was indexed by using the following scale: 1 = < 10% herbivory, 2 = 11-30% herbivory, 3 =

31-50% herbivory, 4 = 51-70% herbivory, 5 = 71-90% herbivory, and 6 = > 90% herbivory. The number of eggs laid per female was counted daily. If feeding or oviposition stopped for a day, the date was recorded, and the pair was kept under their current conditions for 2 weeks more to observe whether or not these behavioral changes remained. If no halt in consumption or oviposition was observed, the beetles were kept under experimental conditions for 3 weeks. Longevity was measured by recording the date that females died. Dead females were identified by having very dry body parts that broke easily when touched and their ovipositor extruded from the abdomen. Longevity of males was not recorded.

Statistical analysis

Data on the total number of eggs laid per female were analyzed using a two-way

ANOVA with 2 factors, temperature and photoperiod. The data were squared root transformed to meet the parametric test assumptions. Data presented in tables and figures were back-transformed. Treatment means were separated by the Tukey–Kramer

HSD test using the agricolae package in R (De Mendiburu 2015). All tests were performed in R (R Core Team 2015) with a significance level of 5% for all statistical analyses.

Results

Data on the number of eggs per treatment, laid per mating pair during 3 weeks after first oviposition, are included in Figure 5-1. The number of eggs was significantly affected by temperature (F = 4.57; df = 3, 24; P = 0.01) whereas the effect of photoperiod was not significant (F = 0.00; df = 1, 24; P = 0.99). No significant interaction effect was found (F = 0.18; df = 3, 24; P = 0.90). The mean number of eggs was 80% lower at 30 than at 21ºC (Figure 5-1). Even though the mean number of eggs did not

differ among 21, 24, and 27ºC, it tended to decrease as temperature increased (Figure

5-1).

A decrease in the number of eggs laid per day was apparent after moving the mating pairs from 21 to 27 and 30ºC at the 10-h photoperiod and 30ºC at the 14-h photoperiod (Figure 5-2). A complete cessation of feeding or ovipositing was not apparent. Seventy-eight percent of the mating pairs stopped laying eggs on average 3,

2, 4, 3 d before dying at 21, 24, 27, and 30ºC (range 1–10). The beetles ate leaf tissue throughout the experiment. The percentage of damage recorded, every 3 d, varied from

11 to 30% regardless of temperature or the photoperiod.

Statistically significant differences in female longevity were found among temperatures (F = 34.668; df = 3, 24; P < 0.001), but not between photoperiods (F =

2.43; df = 1, 24; P = 0.13). There was a temperature * photoperiod interaction effect (F =

8.69; df = 3, 24; P < 0.001) on female longevity. Analysis of simple main effects showed that for females subjected to a 10-h photoperiod, high temperatures (27 and 30ºC) reduced significantly (F = 89.3; df = 3, 13; P < 0.001) the number of days they lived

(Table 5-1). For females under a 14-h photoperiod, only 30ºC caused a significant reduction in longevity (F = 5.7; df = 3, 11; P = 0.01) (Table 5-1).

Discussion

High temperatures have a detrimental effect on the longevity (Table 5-1) and fertility (Figure 5-1) of females of M. ochroloma. Longevity of females was shorter at 27 and 30ºC under the 10-h photoperiod and at 30ºC under the 14-h photoperiod (Table 5-

1). Even though photoperiod seems to cause no effect on longevity, significant differences were observed between the short and long photoperiod at 27ºC. The difference could be explained by variation in the conditions in the environmental

chamber used for the 27ºC – 14-h photoperiod treatment. The average temperature in the chamber was 26.7 ± 4.3 ºC, which could have masked the effect of temperature on female longevity at the longer photoperiod. Oliver and Chapin (1983) reported no effect on M. ochroloma female life span when beetles were subjected to 2 controlled photoperiods (12 and 14 h) at 27ºC.

Manrique et al. (2012) reported detrimental effects of high temperatures on the development and survival of larvae. Larvae reared at 30ºC suffered high mortality, and the surviving larvae developed into smaller females. Manrique et al. (2012) also reported a reduction in herbivory when larvae were reared at 30ºC, but in my study a reduction in feeding by adults was not observed.

A decline in the oviposition rate was observed a couple of days after the beetles were transferred to the environmental chambers set at 30ºC at either short or long photoperiod (Figure 5-2). Similarly, Manrique et al. (2012) observed that adults collected in the field during the winter season and placed at 30ºC stopped laying eggs, while those held at 25ºC continued to be reproductively active. Therefore, a reduction in the number of eggs laid might not only be due to shorter longevity but also a direct effect of temperature on the beetle’s reproductive behavior. The mean total number of eggs per female was reduced at 30ºC, but no effect caused by photoperiod was observed (Figure

5-1). Similar observations were reported by Oliver and Chapin (1983), who found that the total number of eggs laid by female M. ochroloma at 27ºC under 12- and 14-h photoperiods were not different.

The detrimental effect of high temperature on adults and larvae of M. ochroloma indicates that this insect cannot survive under field conditions in Florida during hot

months, unless they can move to other areas where temperatures are maintained below

30ºC. In my study, the combination of high temperature and long photoperiod did not induce changes in the behavior of adults that suggest the beetle entering a dormancy state. Insects in a dormant state are less receptive to food, and their consumption might vary from non- feeding to continuous or sporadic (Tauber et al. 1986, Danks 1987).

Oviposition is stopped or reduced in dormant females (Danks 1987). However, M. ochroloma females fed and oviposited during most of their adult life in my study.

Considering that detrimental temperatures for the development of larvae and adults of

M. ochroloma last about 5 months and the host plant is not cultivated during summer season, this beetle must go through a dormancy state which allows them to survive until optimal conditions are present.

The adult stage might not be able to respond to quick changes in environmental conditions by entering a dormancy state or quiescence. Rather, it must perceive environmental cues earlier in development to enter diapause. According to Danks

(1987), in common cases of adult dormancy, adults with a developed reproductive system cannot enter diapause, and the dormant stage is always the non-parous adults that perceived environmental cues a few days after emergence. However, in other chrysomelids both the larval and adult stages are sensitive to induction of reproductive dormancy. For example, in the Colorado potato beetle, Leptinotarsa decemlineata (Say)

(Coleoptera: Chrysomelidae), the environmental cues that induce diapause are perceived mainly by adults, but the larval stage has some influence (Wilde et al. 1959).

In the cabbage beetle, Colaphellus bowringi (Baly) (Coleoptera: Chrysomelidae), the larva is the sensitive stage for summer diapause (Xue et al. 2002).

Bowers (2003) proposed 3 possible dormancy strategies to explain the low populations of M. ochroloma during summer: quiescence, diapause, or multiple oversummering strategies in which some beetles enter dormancy while others continue developing and ovipositing on a limited basis. Quiescence could still be a possible option if eggs, larvae, or pupae are the dormant stage. However, a quick response to environmental changes could be detrimental if the insects become active before their host plant is present since immature stages cannot travel long distances. However,

Danks (1987) reported that the most common diapause stage in Chrysomelidae is the adult, but a few species and genera diapause as eggs or larvae.

On the other hand, aestivation (i.e., summer dormancy) is also a sound alternative, and it has been reported in some crucifer leaf beetles like C. bowringi and the brassica leaf beetle, Phaedon brassicae (Baly) (Coleoptera:Chrysomelidae), which are important pests of crucifers in China and go through diapause in winter and summer seasons (Xue et al. 2002, Wang et al. 2007). In both species, long photoperiods experienced by larvae or newly emerged adults induced aestivation in the adult stage

(Xue et al. 2002, Wang et al. 2004, 2007).

Even though high temperature was most influential on M. ochroloma longevity and oviposition, both factors, temperature and photoperiod, can still play an important role in M. ochroloma diapause. Photoperiod has been considered a very reliable environmental cue, and many insects use it to signal diapause initiation, maintenance, and termination (Masaki 1980, Tauber et al. 1986, Danks 1987). For example, aestivation by C. bowringi and P. brassicae is induced by long photoperiods, whereas short photoperiods and low temperatures induce diapause in winter (Xue et al. 2002,

Wang et al. 2007). I consider that due to the detrimental effect of high temperature on larvae and adults, M. ochroloma might rely on photoperiod before environmental conditions become unfavorable.

Further studies of M. ochroloma diapause should be focused on determining the stage sensitive for induction of diapause and the environmental factors involved. The effect of high temperature and long photoperiod should be evaluated in larvae, pupae, and newly emerged adults. Changes in behavior such as burrowing in the soil, reduced consumption, cessation of oviposition, and poor development of the reproductive system should be taken into account to determine initiation of diapause. Information on the dormancy behavior of M. ochroloma will provide more tools to organic growers for the management of this pest.

Table 5-1. Mean number of days that females survived under each treatment combination. Photoperiod (h) Temperature (°C) 10 14 21 21.0 ± 0.0 a 20.0 ± 1.0 a 24 21.0 ± 0.0 a 17.3 ± 2.2 a 27 13.7 ± 0.6 b 20.3 ± 0.4 a 30 8.3 ± 1.2 c 12.0 ± 1.7 c Data are means ± SEM. Means within columns and rows with the same letter are not significantly different (P < 0.001). Tukey – Kramer HSD test was used to compare treatment means.

Figure 5-1. Mean number of eggs laid per Microtheca ochroloma female in each treatment combination. Bars are means ± SEM. Temperatures with the same letter are not significantly different (P < 0.001). No significant differences were found between photoperiods at each temperature (P value in black rectangle). Tukey–Kramer HSD test was used to compare treatment means.

Figure 5-2. Mean number of eggs laid per day by Microtheca ochroloma females at four temperatures at (a) 10-h photoperiods and (b) 14-h photoperiod.

CHAPTER 6 OUTREACH TO CRUCIFEROUS CROP GROWERS

Agricultural extension involves a non-formal educational process to promote the diffusion and application of scientific knowledge of agricultural practices (Birkhaeuser et al. 1991, Swanson et al. 1997). One of the goals of agricultural extension is to bridge the gap between academic research and the farmers by promoting a two-way flow of information: technology transference from research centers to farmers and communication of the farmer’s concerns to researchers (Birkhaeuser et al. 1991). The traditional agricultural extension model focuses on a top-down process in which scientific knowledge and information are linearly transferred to farmers through extension agencies that provide advice in relation to specific problems (Birkhaeuser et al. 1991). However, after years of implementing the traditional diffusion/ adoption model, several flaws became apparent (Vanclay and Lawrence 1994). First, products and new technologies obtained from agricultural research were considered “improvements” without considering the appropriateness of them to farmers (Vanclay and Lawrence

1994). There has also been an unequal distribution of impacts and benefits and, in many cases, farmers’ indigenous technical knowledge has been ignored and marginalized (Vanclay and Lawrence 1994). Many of the faults in the traditional model have become a barrier for a farmer’s adoption of new technology.

Farms are complex environments in which interactions among various trophic levels and the environment are in constant change. Therefore, a single technology that may be applicable and appropriate for a farm might not be for other farms, or the technology may need to be adjusted to work in each particular situation (Röling and

Pretty 1997). Routine, calendar-based activities are difficult for farmers to adopt,

therefore, efforts should focus on facilitating learning instead of transferring technology so that farmers are provided with tools to overcome major problems arising in their farms (Röling and Pretty 1997).

To promote a participatory role of farmers in research, facilitate learning, and provide equal accessibility to information by farmers and the farm workers in Florida, an on-line survey about major insect pests in crucifers was conducted. Extension training about monitoring and identification of cruciferous crop pests was conducted in Spanish at a vegetable farm in Florida because most of the workers were Latin-Americans and their proficiency in English was limited. Additionally, an EDIS publication on management of insect pests in cruciferous crops was updated and translated into

Spanish (Appendix E).

Materials and Methods

Online Survey

An online survey titled: “Pests in cole crops: a survey of Florida farmers” was conducted with the objective of determining the importance of insect pests on crucifer crops in Florida. The survey (Appendix A) had 3 blocks of questions: demographic information; farm practices; and pest management. The demographic information included age, educational level, farm tenure status, and location of the farm. The other 2 blocks requested information about the crops planted and the pest management practices used by growers to control pests. Prior to launching the survey, the UF

Institutional Review Board approved the execution of the online survey (IRB#:

IRB201601474).

Telephone or email information of growers in Florida was obtained from the

USDA Organic Integrity database. The company names were searched online to find

other contact information if email address were missing. For those growers whose telephone information only was available, calls were made to obtain their email address.

Additionally, other growers were contacted through the St. Lucie County Extension

Office to obtain their email address. Both organic and conventional growers were surveyed. A total of 50 surveys were distributed, of which only 14 were returned. All data were collected, stored, and analyzed through Qualtrics (Qualtrics, Provo, UT), which is a software for creation and delivery of web-based surveys.

Extension Training

An on-farm extension training activity in Spanish was conducted with the goal of teaching Spanish-speaking workers about monitoring and identification of pests in cruciferous crops. The training was conducted on 2 separate dates at an organic farm located in Indiantown, Florida. Six people attended the first presentation, and 7 attended the second; 3 people attended both trainings. A poster containing pictures and common names, in English and Spanish, of cruciferous crop pests was made, and an oral presentation was delivered to the farm crew. The workers learned about morphological and behavioral characteristics that could be used to identify the insects directly in the field without using specialized equipment. They were also taught about methods of monitoring and reporting insect pests in crucifers. Ten workers participated in the training. At the end of the training activity, a survey (Appendix B) was delivered to each participant to receive feedback about the extension activity. An English version of the survey is included in Appendix C.

EDIS publication

The EDIS publication ENY-464 titled: “Insect Management for Crucifers (Cole

Crops) (Broccoli, Cabbage, Cauliflower, Collards, Kale, Mustard, Radishes, Turnips)”,

published originally by Webb (2013), was updated. Information about other pests of crucifer crops was included and pesticide tables were revised. Additionally, the updated version of the EDIS publication was translated into Spanish.

Results

Online Survey

Demographic information and farm practices. All respondents were 25 years old or older. Half of the respondents was over 65 years old. The survey was answered mainly by owners and operators (79%) who have at least 6 years of experience in farming. The area under cultivation ranged from 1 to 950 acres. Two of the farms were located in North Florida, 6 in Central Florida, and 6 in South Florida (Figure 6-1).

Conventional farming is the most common system used by the respondents

(43%), followed by organic farming (36%). Most farmers grow crucifers in open field areas. However, 30% of the respondents reported growing crucifers in both open areas or in greenhouses or shade-protected areas. The 5 most cultivated crucifers are collards, kale, broccoli, arugula, and turnips (Figure 6-2). Besides the crucifers listed in the questionnaire, radish is also cultivated by some respondents.

Pest management. The 3 most common insect pests reported are aphids

(Hemiptera: Aphididae), the diamondback moth, Plutella xylostella (Linnaeus)

(Lepidoptera: Plutellidae), and the cabbage looper, Trichoplusia ni (Hübner)

(Lepidoptera: Noctuidae) (Figure 6-3). However, the insects that cause the greatest amount of yield loss (primary pests) are the diamondback moth and aphids (Figure 6-4).

Aphids are also considered the most important secondary pest, followed by the great southern white, Ascia monuste (Linnaeus) (Lepidoptera: Pieridae), and the cabbage looper (Figure 6-4). Other insect pests reported but not listed in the questionnaire are

the cross-striped cabbageworm, Hellula rogatalis (Hulst) (Lepidoptera: Crambidae), the broad mite, Polyphagotarsonemus latus (Banks) (Arachnida: Acari: Tarsonemidae), and mole crickets (Orthoptera: Gryllotalpidae).

The preferred methods for managing the primary pests are the use of synthetic insecticides (50%) and biological control (21%). One responder reported using synthetic insecticides to control the diamondback moth without success. Secondary pests are controlled with synthetic insecticides (43%) and botanical or microbial products (36%).

Only 5 respondents observed Microtheca ochroloma Stål (Coleoptera:

Chrysomelidae) in their farms, and only 1 of them considered this beetle a primary pest.

The farms where M. ochroloma was reported are located in central Florida, specifically in Polk, Martin, Sarasota, and Hillsborough counties. Three of the respondents use organic farming systems, the other 2 use bio-dynamic or conservation farming systems.

In all these systems, there is a restricted or reduced use of synthetic insecticides.

However, the preferred methods used to control M. ochroloma are the application of botanical and microbial insecticides allowed in organic production.

Extension Training

The poster in Appendix D was used to train 10 workers on insect monitoring and identification. The trainings were conducted on June 3 and 10 of 2016. Each oral presentation lasted 1 h. The pictures in the poster were used to describe the pests

(Figure 6-5 a, b). Additionally, some specimens of M. ochroloma adults, larvae, and A. monuste adults were shown to the participants.

Ten training attendees completed a survey at the end of the training activity

(Figure 6-5 c), in which they reported an increase in knowledge of 66, 33, 66, and 66% about the importance of monitoring, monitoring steps, pest and damage identification,

and reporting pests, respectively (Figure 6-6). The attendees considered the information taught during the activity was relevant to their professional needs and met their expectations. Eighty percent of the attendees identified themselves as Hispanic/Latino.

EDIS publication

The EDIS manuscripts, English and Spanish versions, are found in Appendices E and F, respectively. In the English version, 2 new crucifer pests were added: brown stink bugs, Euschistus obscurus (Palisot de Beauvois) and Euschistus servus (Say)

(Hemiptera: Pentatomidae), and the great southern white. Information on recognition, biology, and damage to plants was included. Additionally, a table with recommended management practices for M. ochroloma was added. All insect illustrations were replaced with pictures obtained from the Entomology and Nematology Insect

Photography Database. The table of insecticides was updated according to the

Handbook of Vegetables 2015-2016. Some insecticides whose registration was cancelled or are no longer sold or distributed, such as thiocarb and flubendiamide, were removed. The tables of insecticides, in the English and Spanish versions, are not included in this dissertation document but they are included in the EDIS publications.

Discussion

The insect pest reported by the survey respondents as the most serious pest that attacks their crucifer crops is the diamondback moth. Webb (2013) also reported the diamondback moth as the most important pest of crucifers in Florida. This species is a highly invasive, broadly distributed lepidopteran (Talekar and Shelton 1993). The management of this insect is problematic because populations have developed resistance to many synthetic pesticides (Shelton et al. 1993, Zhao et al. 2006) and even to commercial products of the bacterium Bacillus thuringiensis Berliner (Tabashnik et al.

1990). However, the preferred management method used by the respondents for this pest is synthetic insecticides, but 1 of the respondents reported using various pesticides without successful control. The respondents who use organic farming to cultivate crucifers reported using cultural, physical, and biological control tactics but no information about the specific tactics and success of those methods was gathered through the survey. With the failure of many pesticides to control the diamondback moth, several studies on cultural methods have been done, from which intercropping

(Talekar et al. 1986, Åsman et al. 2001), sprinkler irrigation (Talekar et al. 1986), trap cropping (Badenes-Perez et al. 2004, Shelton and Nault 2004), crop rotation, and clean cultivation have shown some potential to manage the pest (Talekar and Shelton 1993).

Other serious pests that were reported by respondents were aphids. Webb

(2013) reported that even though aphids occur sporadically in Florida, they follow diamondback moth and cabbage looper in importance. Crucifer plants can be infested by 3 species of aphids: the turnip aphid, Lipaphis erysimi (Kaltenbach); the green peach aphid, Myzus persicae (Sulzer); and the cabbage aphid, Brevicoryne brassicae

(Linnaeus) (Webb 2013, Razaq et al. 2011). Infestations by aphids can cause significant yield loss if management tactics are not implemented to reduce their population (Razaq et al. 2011). Several respondents consider aphids the second most important pest in their crops. Another pest listed as a secondary pest was the cabbage looper, which feeds on other vegetable crops besides crucifers (Shorey et al. 1962, Webb 2013) and is considered a minor pest in North Florida (Webb 2013). However, in the survey, respondents located in central Florida also reported it as the second most important

pest affecting their crucifers. For secondary pests, most respondents applied synthetic, botanical, or microbial insecticides to control their populations.

Microtheca ochroloma was not considered an important pest by most respondents. Those respondents who reported M. ochroloma in their crops have used botanical or microbial insecticides to control them, which are the methods suggested by

Balusu and Fadamiro (2012, 2013).

The results obtained through this on-line survey should be considered cautiously because of the small number of respondents. More participants should have responded to the survey to obtain more comprehensive data sets about the current status of crucifer pests in Florida and the management strategies implemented by farmers. I consider that a higher emphasis on outreach to farmers is needed. Even though research on alternative tactics to pesticides for the management of this pest has been conducted (Talekar et al. 1986, Talekar and Shelton 1993, Åsman et al. 2001 Badenes-

Perez et al. 2004), farmers seem to rely on insecticide use only. Similarly, extension activities to teach not only farmers but also their workers should be considered. Workers in charge of crop maintenance and harvest could play a crucial role in the monitoring and early detection of pests. In Florida, where there is a high proportion of Latin-

Americans working in farms, it is important to make accessible the information needed for insect identification and monitoring through translation of published information into

Spanish. Extension activities in Spanish would also be beneficial.

Figure 6-1. Map of Florida counties where survey respondents’ farms are located.

Figure 6-2. Number of respondents reporting the crucifer crops commonly planted on their farms.

Figure 6-3. Frequency of insect pests on crucifer crops reported by survey respondents.

Figure 6-4. Frequency of primary and secondary pests of crucifer crops reported by survey respondents.

Figure 6-5. Extension training at an organic farm: (a, b) oral presentation; (c) participants responding to the end-of-training survey. Photographs by Diego Ramirez, University of Florida.

Figure 6-6. Change in knowledge before and after extension training.

CHAPTER 7 CONCLUSIONS

The main objective of my dissertation was to study behavioral, biological, and outreach approaches that might contribute to long term management of Microtheca ochroloma Stål (Coleoptera: Chrysomelidae). I concentrated on the following approaches: (1) evaluate the impact of 3 entomopathogenic fungi on the fertility and fecundity of M. ochroloma; (2) assess the effect of 10 non-crucifer plant essential oils on herbivory by adults of M. ochroloma; (3) determine the effect of temperature and photoperiod on M. ochroloma adults’ dormancy response; and (4) determine the insect pest problems of most concern to crucifer farmers in Florida and increase accessibility to information on crucifer pests through a survey and extension activities.

The use of the entomopathogenic fungi Beauveria bassiana (Balsamo) Vuillemin,

Metarhizium anisopliae (Metschnikoff) Sorokin (both Hypocreales: Clavicipitaceae), and

Isaria (=Paecilomyces) fumosorosea Wize (Hypocreales: Cordycipitaceae) did not affect the fecundity and fertility of M. ochroloma. However, M. anisopliae reduced the longevity of females compared to the control treatments. The low mortality caused by these entomopathogenic fungi and the inability to affect the fecundity or fertility of M. ochroloma adults do not portend good management of this pest by using commercially available entomopathogenic fungi. The potential effect of M. anisopliae on the immature stages and the effect on fecundity and fertility of M. ochroloma adults after being treated in the larval stage with sublethal doses of these entomopathogenic fungi should be further tested.

From the 10 non-crucifer essential oils tested, neem, hyssop, and thyme might cause a reduction in leaf damage by M. ochroloma adults. A commercial neem product

containing 1.2% azadirachtin caused a reduction in the percentage of leaf area consumed compared to the control in a no-choice and paired choice tests. The neem commercial product tested might be potentially integrated with trap cropping using turnip to increase the effectiveness of trap cropping by reducing damage on the main crop

(antifeedant effect of neem) and concentrating M. ochroloma on turnips which will allow an easier and more effective control of the pest. Further testing on whole plants in laboratory conditions and field conditions is necessary to determine the effectiveness of the use of neem for the management of M. ochroloma.

The survey conducted showed that M. ochroloma is not considered the main insect problem in crucifer crops but this beetle does cause some problems in production systems with a reduced use of synthetic insecticides. The main insect problems affecting the responder’s crops were the diamondback moth and aphids, therefore research on pest management methods should focus on managing those 2 pests. The use of synthetic, botanical and microbial insecticides is the preferred method of control used by the respondents. A more extensive survey that includes more farmers in other areas of Florida should be conducted.

The isolated use of management tactics is not sufficient to manage M. ochroloma. Therefore, integration with other methods for management is necessary.

The efficacy of various methods can be increased by integrating them into a management program so each tool might contribute to the overall objective of reducing the impact of M. ochroloma in crucifer crops.

APPENDIX A ON-LINE SURVEY QUESTIONNAIRE

Pests in cole crops: a survey of Florida farmers

Thank you for choosing to participate in this survey. We value your opinion and honest feedback. The purpose of the survey is to measure the effect of insect pests on crucifer crops in Florida. Results will provide information on the main insect problems that growers in this area encounter, which will serve as a guide for future research and extension programs. Participation in the survey is voluntary. The survey will take approximately 5 minutes. Personal identity information will not be collected. Data will be summarized by county, farm size, or agricultural production system, but individual survey responses are confidential and will never be shared. All study data will be collected through an online survey-collection program called Qualtrics. Qualtrics is a secure site with SAS 70 certification for rigorous privacy standards. Any data that you provide through this program will be encrypted for security purposes using Secure Socket Layers (SSL). Only the study investigators will have access to the data on Qualtrics. To protect your privacy, all participants’ IP addresses will be masked by Qualtrics and will be unavailable to, and unidentifiable by, investigators or others. Qualtrics’ privacy policy can be obtained at http://www.qualtrics.com/privacy-statement If you have questions about the survey, please contact Angie Nino at [email protected]

Q1 What is your age?  18 to 24 years (1)  25 to 34 years (2)  35 to 44 years (3)  45 to 54 years (4)  55 to 64 years (5)  Age 65 or older (6)

Q2 Do you have any agricultural land or do farming?  Yes (1)  No (2)

Q3 What is your tenure status?  Owner-operator (1)  Leasee (2)  Tenant (3)  Hired laborer (4)  Other (5) ______

Q4 What is the highest degree or level of education you have completed?  Less than high school (1)  High school graduate (includes equivalency) (2)  Some college, no degree (3)  Associate's degree (4)  Bachelor's degree (5)  Master's degree (6)  Doctoral degree (7)  Professional degree (8)  Other (9) ______

Q5 How many years of farming experience do you have?

Q6 What is the size of your farm (in acres)?

Q7 How many acres were actually under cultivation? In 2014 (1) In 2015 (2) In 2016 (3)

Q8 Where is the farm located? County (1) City (2) Zip code (3)

Q9 How do you use the products you cultivate?  For subsistence (1)  Selling at in-state markets (2)  Selling at out-of-state markets (3)  Selling at international markets (4)  U-pick (5)  Other (6) ______

Q10 Have you cultivated crucifer crops such as broccoli, cabbage, turnips, mustard, collards, kale, napa cabbage, arugula, radish, bok choy and any other Asian leafy green (mizuna, mibuna, tatsoi) in the last five years?  Yes (1)  No (2)

Q11 Which of the following crucifers do you consistently grow or produce?  Broccoli (1)  Cabbage (2)  Collards (3)  Turnips (4)  Mustard (5)  Bok choy (6)  Napa cabbage (7)  Arugula (8)  Kale (9)  Other Asian leafy greens (mizuna, tatsoi, yu choy) (10)  Other (11) ______

Q12 What agricultural production system do you use to grow crucifers?  Conventional farming (1)  Transition to organic farming (2)  Organic farming (3)  Conservation farming (4)  Bio-dynamic farming (5)  Other (6) ______

Q13 Where do you grow your crucifers?  Open field (1)  Protected culture (e.g. greenhouse, covers, tends, etc.) (2)  Both (3)

Q14 Which insect pest(s), if any, did you have last cropping season?

 Diamondback Moth (1)  Aphids (2)  Beet Armyworm (3)  Cabbage Looper (4)  Cabbage Webworm (5)  Stinkbugs (8)  Cutworms (6)  Yellowmargined Leaf Beetle (7)  Great Southern White (11)  Other (9) ______ None of the above (10)

Q15 Which pest caused the greatest amount of yield loss to your crucifer crops last season?  Diamondback Moth (1)  Aphids (2)  Beet Armyworm (3)  Cabbage Looper (4)  Cabbage Webworm (5)  Stinkbugs (6)  Cutworms (7)  Yellowmargined Leaf Beetle (8)  Great Southern White (9)  Other (10) ______

Q16 Which one of the following was your second most important pest problem?  Diamondback Moth (1)  Aphids (2)  Beet Armyworm (3)  Cabbage Looper (4)  Cabbage Webworm (5)  Stinkbugs (6)  Cutworms (7)  Yellowmargined Leaf Beetle (8)  Great Southern White (9)  Other (10) ______

Q17 How did you control the primary pest problem?  Chemical control applying synthetic pesticides (e.g. Malathion, Sevin, Furadan, etc.) (1)  Chemical control applying botanical or microbial biopesticides, oils, and insecticidal soaps (e.g. Bt, Entrust, Pyganic, Azadirachtin, etc.) (2)  Biological control (3)  Environmental or cultural control (planting time, resistant varieties, etc.) (4)  Physical control (insect traps, barriers, etc.) (5)  Other (6) ______

Q18 How did you control the secondary pest problem?  Chemical control applying synthetic pesticides (e.g. Malathion, Sevin, Furadan, etc.) (1)  Chemical control applying botanical or microbial biopesticides, oils, and insecticidal soaps (e.g. Bt, Entrust, Pyganic, Azadirachtin, etc.) (2)  Biological control (3)  Environmental or cultural control (planting time, resistant varieties, etc.) (4)  Physical control (insect traps, barriers, etc.) (5)  Other (6) ______

Q19 The yellowmargined leaf beetle is a small beetle (5 mm/2 in long). Adults are black or brown with a white or yellow coloration on the margin of their hard wings. Larvae are brownish and congregate on the underside of the leaves of the plants. Both stages chew holes in the leaves. Have your crucifer crops been attacked by the yellowmargined leaf beetle in the past seasons?

 Yes (1)  No (2)  I do not know (3)

Q20 In which season(s) is the yellowmargined leaf beetle more abundant?  Fall (1)  Winter (2)  Spring (3)  Summer (4)

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Q21 How do you control the yellowmargined leaf beetle?  Applied synthetic pesticides (e.g. Malathion, Sevin, Furadan, etc.) (1)  Applied botanical or microbial biopesticides (e.g. Bt, Entrust, Pyganic, Azadirachtin,etc.) (2)  Trap cropping (3)  Removal of infested plants (4)  Infested plants incorporated into the soil (5)  Do nothing (6)  Other (7) ______

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APPENDIX B EXTENSION TRAINING SURVEY FORMAT SPANISH VERSION

Survey format provided to participants at the end of the extension training.

Evaluación Final de la Actividad de Extensión

En búsqueda de mejores maneras de servirle, por favor tome un momento para completar esta pequeña encuesta, la cual nos servirá para saber qué estamos haciendo y cómo podemos mejorar para suplir sus necesidades futuras.

Satisfacción Por favor encierre en un círculo el número apropiado de su nivel de respuesta. No está Un poco Muy Satisfecho Cuán satisfecho está usted con: satisfecho satisfecho satisfecho La relevancia de la información para sus 1 2 3 4 necesidades La calidad de la presentación realizada por el 1 2 3 4 instructor Conocimiento de la materia del instructor 1 2 3 4 Recursos utilizados en la actividad 1 2 3 4 En general la calidad de la actividad 1 2 3 4

¿Fue la información fácil de entender? 1. Si 2. No

Cambio en prácticas de monitoreo: Por favor encierre en un círculo el número correspondiente a su nivel de conocimiento acerca de los siguientes temas antes y después de completar la actividad de extensión. Por favor use la siguiente escala: 1. Muy bajo = No sabe nada acerca de este tema. 2. Bajo = Sabe un poco acerca de este tema. 3. Moderado = Sabe sobre este tema pero hay más cosas que aprender. 4. Alto = Tiene un buen conocimiento de este tema pero hay más cosas que aprender. 5. Muy alto = Sabe casi todo acerca de este tema

ANTES DE LA ACTIVIDAD DE DESPUES DE LA ACTIVIDAD DE % de Como EXTENSION EXTENSION incremento categoriza su Muy Muy Muy Muy conocimiento Bajo Moderado Alto Bajo Moderado Alto bajo alto bajo alto acerca de: Importancia del monitoreo de 1 2 3 4 5 1 2 3 4 5 plagas. Identificación de insectos plaga y 1 2 3 4 5 1 2 3 4 5 descripción del daño causado.

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Pasos para el monitoreo de 1 2 3 4 5 1 2 3 4 5 estos insectos plaga. Reporte de la presencia de 1 2 3 4 5 1 2 3 4 5 estos insectos plaga.

¿Considera que la actividad de extensión llenó sus expectativas? 1. Si 2. No

¿Recomendaría esta actividad de extensión a otros? 1. Si 2. No Si marcó no, indique por qué______

¿Qué le gustó más de esta actividad de extensión?______

¿Qué fue lo que menos le gustó de esta actividad de extensión?______

¿Cómo podríamos mejorar esta actividad de extensión?______

Demografía

¿Cuál es su género? 1. Masculino 2. Femenino

¿Cómo se identifica? 1. Afroamericano 5. Blanco 2. Nativo Americano/Alaska 6. Nativo Hawaiano/Islas del Pacífico 3. Asiático 7. Otro 4. Hispano/Latino

¿Cuál es su país de origen? ______

Gracias por completar esta evaluación. Apreciamos su aporte para mejorar los programas de extensión.

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APPENDIX C EXTENSION TRAINING SURVEY FORMAT ENGLISH VERSION

Survey format provided to participants at the end of the extension training.

End of Workshop Evaluation

UF/IFAS St Lucie County Extension is always looking for ways to serve you better. Please take a moment to complete this short survey. It will help us know how we’re doing, and how we can better meet your future needs.

Satisfaction Please circle the appropriate number for your level of response. How satisfied are you with: Not Somewhat Satisfied Very Satisfied Satisfied Satisfied The relevance of information to your needs? 1 2 3 4 Presentation quality of instructor(s)? 1 2 3 4 Subject matter knowledge of instructor(s)? 1 2 3 4 Training facilities? 1 2 3 4 The overall quality of the training workshop? 1 2 3 4

Was the information easy to understand? 1. Yes 2. No

Change in monitoring practices: Please circle the appropriate number to indicate your level of knowledge about the following topics before and after completing the program. Please use the following key for rating: 1. Very Low = Don’t know anything about this topic. 2. Low = Know very little about this topic 3. Moderate = Know about this topic but there are more things to learn 4. High = Have good knowledge but there are things to learn 5. Very High = Know almost everything about this topic N=44 BEFORE THIS WORKSHOP AFTER THIS WORKSHOP % How do you rate increase your knowledge Very Low Moderate High Very Very Low Moderate High Very about: low high low high The importance of insect pest 1 2 3 4 5 1 2 3 4 5 monitoring Identification of insect pests and 1 2 3 4 5 1 2 3 4 5 description of damage Steps to monitoring insect 1 2 3 4 5 1 2 3 4 5 pests Report of the presence of 1 2 3 4 5 1 2 3 4 5 insect pests

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Did the training workshop meet your expectation? 1. Yes 2. No

Would you recommend this training workshop to others? 1. Yes 2. No If not, why? ______

What did you like the most about this training workshop?

What did you like the least about this training workshop?

How could this training be further improved?

Demographics

What is your gender? 1. Male 2. Female

How do you identify yourself? 1. African American 5. White 2. American Indian/Alaskan 6. Native Hawaiian/Pacific Islander 3. Asian 7. Other 4. Hispanic/Latino

Which is your country of origin? ______

Thank you for completing this evaluation. We appreciate your input as we make every effort to improve Extension programs.

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APPENDIX D POSTER USED IN THE EXTENSION TRAINING

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APPENDIX E EDIS PUBLICATION ENGLISH VERSION Insect Management for Crucifers (Cole Crops) (Broccoli, Cabbage, Cauliflower, 1 Collards, Kale, Mustard, Radishes, Turnips)

S. E. Webb2, A. Nino2, H. A. Smith2

Cruciferous vegetables are a large and increasingly important crop group in Florida. A number of insects feed exclusively on crucifers and affect all of the crops listed in the title above. Because most of the newer insecticides are being labeled for the entire crop group or for a subset, either head and stem Brassicas (like cabbage and broccoli) or leafy Brassicas (like kale), there are no longer specific tables for individual Brassica crops at the end of this document. Instead, exceptions are given in the Notes column. Other crucifers not listed in the title but which also have the same pest complex include head and stem Brassicas such as Brussels sprouts, Chinese broccoli, and Chinese mustard, and leafy Brassicas such as bok choy, mizuna, and rape greens. Check pesticide labels carefully to see if these crops are included. Radishes and turnip roots are included in the root vegetables group, even though they are also crucifers and have similar pest problems. A separate table of pesticides for radishes is included.

Diamondback moth is the most serious pest of crucifers in Florida. Cabbage looper is also considered a major pest, although it has been less of a problem over the past decade. Insect pests that have been considered major in the past and are only occasionally a problem now include aphids (turnip, green peach, cabbage), harlequin bug, beet armyworm, cabbage webworm, and cutworms (black and granulate). Yellowmargined leaf beetle is a particular problem on mustard and Chinese cabbage, especially for organic growers. Cross-striped cabbageworm is more of a problem on broccoli and cauliflower than it is on other crucifers. Aphids, cutworms, and wireworms are the major insect pests affecting radishes. Diamondback Moth

Plutella xylostella (Linnaeus) (Lepidoptera: Plutellidae)

Description

The adult moth (Figure 1) is small and slender with very long antennae. It is grayish-brown with a broad cream or light brown band along its back. The band can have constrictions, which give it a diamond-like pattern. When viewed from the side, the wing tips appear to turn up slightly. Eggs are oval and flattened, yellow to pale green, and approximately 0.02 inches long and 0.01 inches wide. There are four larval instars. Even the oldest is quite small and very active. Larvae will wriggle violently if disturbed and will drop from the leaf suspended by a strand of silk. The body tapers at both ends and the fifth pair of prolegs (abdominal legs) protrudes from the

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posterior (Figure 2). After the first instar, which is colorless, the larvae are green. Larvae pupate in a loose cocoon on lower or outer leaves or in the florets of cauliflower and broccoli.

Figure 1.

Top and side of diamondback moth. Credit: Lyle J. Buss, University of Florida.

[Click thumbnail to enlarge.]

Figure 2.

Diamondback moth larva. Credit: Lyle J. Buss, University of Florida.

[Click thumbnail to enlarge.]

Biology

The female moth attaches her eggs to the lower leaf surface, either singly or in groups of two or three. Within a few days, the eggs hatch, and the larvae begin to feed on the underside of the leaf. The larval stage can last from ten days to a month, depending on temperature. Diamondback moth larvae slow their feeding at temperatures below 50°F, and population growth is most rapid at temperatures greater than 80°F. The pupal stage is passed within a transparent, loose cocoon, which is usually attached to the underside of leaves. In warm weather, the pupal stage may be completed in 3 to 4 days.

In southern Florida, diamondback moth is most abundant from December to February or March and can attack at any time during the crop cycle. By the end of May, moth counts in pheromone traps fall to near zero. Moth counts may rise in mid-fall through early winter, but activity is limited during that time. Populations build on winter weeds, such as wild mustard, before moving into winter and early spring plantings of cabbage and other crucifers. From mid-winter

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through the spring, when it is a serious pest, diamondback moth may cause losses of up to 70 percent in the absence of control. Populations may decrease after heavy rains.

Damage

Plants at all stages of growth may be attacked. Larvae chew small holes in leaves, with larger larvae making larger holes. Young larvae often feed on one surface of the leaf, leaving a thin layer or “window” of leaf epidermis. Diamondback moth larvae will also attack developing cabbage heads. The resulting damage deforms the heads and leaves entry points for decay pathogens. Cabbage Looper

Trichoplusia ni (Hübner) (Lepidoptera: Noctuidae)

Description

Figure 3. Cabbage looper adult male. Credit: Lyle J. Buss, University of Florida. [Click thumbnail to enlarge.]

Figure 4.

Cabbage looper larva. Credit: Lyle J. Buss, University of Florida.

[Click thumbnail to enlarge.]

The cabbage looper feeds on a variety of crops. The adults (Figure 3) are night-flying moths with brown, mottled fore wings marked in the center with a small, silver figure eight-like spot. Eggs are small, ridged, round, and greenish-white. They hatch into larvae that are green with white stripes running the length of their bodies. The caterpillar (Figure 4) has three pairs of slender legs near its head and then three pairs of thick prolegs near the end of its body. It moves in a

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characteristic looping motion, alternately stretching forward and arching its back as it brings the back prolegs close to its front legs. The caterpillar is about 1.25 inches long when fully grown.

Biology

Eggs are deposited singly or in small clusters on either leaf surface, although more are found on the lower leaf surface. Each female moth can produce 300 to 600 eggs during the approximately 10 to 12 days it is alive. Two to four weeks after hatching, the mature larva spins a thin cocoon on the lower leaf surface, or in plant debris or soil. The pupal stage lasts approximately two weeks. Total time required for development from egg to adult averages 42 days at 69.8°F and 22 days at 89.6°F. Development is abnormal above 94°F.

Populations tend to be highest during the late spring and summer months, and in some years in the late fall. Cabbage looper does not enter diapause and cannot survive prolonged cold weather. The insect remains active and reproduces throughout the winter months only in the southern part of Florida (south of Orlando). In central Florida, cabbage looper populations peak during early fall and again during late spring.

Damage

The cabbage looper is still an important annual pest in north Florida crucifers. It is less of a problem in southern Florida, where it is considered a minor pest. In general, cabbage looper is more of a problem during the fall than during the winter or spring months.

Cabbage looper larvae damage plants by chewing holes in leaves. Smaller larvae remain on the lower leaf surface, while larger larvae produce larger holes throughout the leaf. In addition to feeding on the wrapper leaves of cabbage, larvae may bore into the developing head. Some defoliation can be tolerated before head formation, but feeding damage and excrement left behind on heads make cabbage unmarketable. Cabbage with damage confined to wrapper leaves is marketable but with reduced value. Aphids

Turnip aphid [Lipaphis erysimi (Kaltenbach)], green peach aphid [Myzus persicae (Sulzer)], cabbage aphid [Brevicoryne brassicae (Linnaeus)]

Description

Turnip aphid (Figure 5) and green peach aphid (Figure 6) are the most important aphids on crucifers in Florida. Cabbage aphid (Figure 7) is not as common in Florida. Although aphid problems on crucifers in Florida tend to be sporadic, they follow diamondback moth and cabbage looper in importance. Adults are soft-bodied, pear- or spindle-shaped insects with a posterior pair of tubes (cornicles or siphunculi), which project upward and backward from the dorsal surface of the abdomen and which are used for excreting an alarm pheromone. Aphids have fine piercing- sucking mouthparts with which they penetrate leaf tissue to feed on phloem sap. Nymphs are smaller but otherwise similar in appearance to wingless adults.

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Figure 5

Turnip aphid. Credit: Lyle J. Buss, University of Florida.

Figure 6

Green peach aphid. Credit: Lyle J. Buss, University of Florida.

Figure 7.

Cabbage aphid. Credit: Lyle J. Buss, University of Florida.

[Click thumbnail to enlarge]

Turnip aphid adults are whitish-green or green, about 0.06–0.09 inches long. The antennae are dark and the cornicles are pale with dusky tips. The body is covered with a white secretion. Nymphs are pale greenish yellow. Green peach aphid adults vary from 0.04 to 0.08 inch in length and are light green to yellow to pink and pear-shaped. The tubercles (bumps between antennae) point inward and are a distinguishing characteristic. Winged forms have a black patch on the back of the abdomen. Cabbage aphid adults are very similar in appearance to turnip

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aphids although larger (0.08- 0.1 inches long) with shorter cornicles and are covered with a grayish waxy powder.

Biology

Aphids reproduce very rapidly. In Florida, males are uncommon and females give birth to live nymphs all year without mating or laying eggs. Nymphs mature in 7 to 10 days. When host plant quality deteriorates or if plants become overcrowded, winged forms develop and migrate to new host plants. They are often protected from their many natural enemies by ants, which feed on the sugary waste product of the aphids called honeydew. Aphids are more abundant during the spring and fall and almost disappear in summer.

Damage

Green peach aphid is a major pest of greens (collards, kale, and mustard), as well as many other unrelated crops. They attack cabbage mainly before heading begins. Turnip aphids attack only crucifers, preferring turnips and radishes. Aphids suck plant juices with their piercing-sucking mouthparts, resulting in yellowing and curling of the leaves. The plant, particularly when attacked as a seedling, may become stunted or die as a result of aphid feeding. Foliage may be contaminated with aphid bodies, cast skins, and honeydew. Aphids feeding within the curled leaves or inside the cupped leaves of headed plants are normally protected from contact insecticides. Green peach aphid and turnip aphid transmit turnip mosaic virus in Florida. Beet Armyworm

Spodoptera exigua (Hübner) (Lepidoptera: Noctuidae)

Description and Biology

The beet armyworm has a wide host range, and in addition to crucifers, attacks such vegetables as asparagus, bean, beet, celery, chickpea, corn, cowpea, eggplant, lettuce, onion, pea, pepper, potato, spinach, sweet potato, and tomato. It also feeds on many field crops and weeds.

The highly mobile adult moth (Figure 8) has dark front wings with mottled lighter markings and hind wings thinly covered with whitish scales. Each female can lay over 600 eggs, generally in masses of about 100 on the undersides of leaves in the lower plant canopy. Egg masses are covered with fuzzy, white scales. Very young caterpillars (Figure 9), which are pale with dark heads, feed in groups and then disperse as they grow older (third instar). By the third instar, caterpillars have wavy, light-colored stripes lengthwise down the back and broader stripes on each side. Although often dull green, the color of caterpillars can vary. After feeding from one to three weeks, they construct a cocoon from sand and bits of soil and pupate in the soil, emerging as adults about one week later. Beet armyworm is a tropical insect and survives the winter in southern Florida where it can complete many generations a year. From southern Florida, adults migrate into northern Florida and other parts of the Southeast.

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Figure 8.

Beet armyworm adult. Credit: Lyle J. Buss, University of Florida.

[Click thumbnail to enlarge.]

Figure 9.

Beet armyworm larva. Credit: Lyle J. Buss, University of Florida.

[Click thumbnail to enlarge.]

Damage

Larvae feed on foliage of host plants. Beet armyworm larvae consume greater amounts of leaf tissue than the diamondback moth but not as much as the cabbage looper. An action threshold of 0.3 beet armyworm larvae per plant has been used on cabbage in Texas. Because adults can readily invade a field from nearby crops or weeds, monitoring the crop twice a week for beet armyworm presence and damage is recommended.

Beet armyworm is a sporadic pest on Florida crucifers, and it is usually kept under damaging levels by controls targeted to diamondback moth. It can be a serious pest of napa cabbage in northern Florida in the spring. Beet armyworm populations in southern Florida are highest from late March through mid-June, with a smaller population rise from mid-August through October. The increase in the late summer and fall is thought to be related to beet armyworm activity on late summer weeds, while the population increase in the spring coincides with the vegetable production season in southern Florida. Cabbage Webworm

Hellula rogatalis (Hulst) (Lepidoptera: Crambidae)

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Description

The moth (Figure 10) has yellowish-brown front wings marked with white bands and a dark kidney-shaped spot. The hind wings are grayish-white with a darker margin. The wingspan is about 0.7–0.8 inches. Eggs have a flattened shape and are gray or yellowish-green to begin with but turn pink as they get close to hatching. There are five larval instars. The mature larva (Figure 11) is yellowish-gray with five brownish-purple bands running the length of its body. Its head is black. Moderately long yellow or light brown hairs sparsely cover the body.

Figure 10.

Cabbage webworm adult. Credit: Lyle J. Buss, University of Florida.

[Click thumbnail to enlarge.]

Figure 11.

Cabbage webworm larva. Credit: Lyle J. Buss, University of Florida.

[Click thumbnail to enlarge.]

Biology and Damage

Cabbage webworm eggs are usually laid singly or in small masses on the terminal leaves. Upon hatching, the larvae mine the leaves and also feed on the underside of the leaves producing small holes. At about the third instar, larvae begin to web and fold the foliage. The webs become covered with dirt and excrement. Larger larvae can burrow into buds, stems, and leaves. The insect may feed on the growing point, causing severe damage to young plants. When fully grown, larvae pupate in the buds, on the sides of stems, or on the surface of the soil.

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Like beet armyworm, cabbage webworm is seen sporadically and is controlled by treatments for diamondback moth. A related species, H. phidilealis, or cabbage budworm, can be a problem in southern Florida. Cutworms

Black cutworm [Agrotis ipsilon (Hufnagel)] and granulate cutworm [Feltia subterranea (Fabricius)] (Lepidoptera: Noctuidae)

Description

Black cutworm moths (Figure 12) are large, with a wingspan of 1.5 to slightly over 2 inches. The front wings are dark brown with a lighter band near the end of each wing. The hind wings are whitish to gray. The ribbed eggs are first white, and then turn brown and are usually deposited in clusters. The larvae (Figure 13) are stout, gray caterpillars with a greasy appearance. Black cutworm larvae have numerous dark, coarse granules over most of their bodies.

Figure 12.

Black cutworm adult. Credit: John Capinera, University of Florida.

[Click thumbnail to enlarge.]

Figure 13.

Black cutworm larva. Credit: John Capinera, University of Florida.

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Granulate cutworm moths (Figure 14) are smaller, with a wingspan of 1.2 to 1.7 inches. The front wings are often yellowish-brown and have distinct bean-shaped and round spots in the

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center. The hind wings are mostly white. Eggs are hemispherical and ridged. Like black cutworm eggs, they are initially white and darken with age. Larvae (Figure 15) are grayish to reddish- brown. Each abdominal segment has a dull yellowish oblique mark. A weak gray line occurs along the length of the body with spots of white or yellow.

Figure 14.

Granulate cutworm adult. Credit: Lyle J. Buss, University of Florida.

[Click thumbnail to enlarge.]

Figure 15.

Granulate cutworm larva. Credit: Lyle J. Buss, University of Florida.

[Click thumbnail to enlarge.]

Biology

The black cutworm is one of the most destructive of the cutworms and attacks a wide range of plants. Although cutworm larvae can migrate into a field from adjacent areas, most migration occurs by adults flying into the field. The moth deposits eggs in groups of one to 30 on leaves, stems, stubble, or field debris near ground level. The egg stage lasts from 5 to 15 days, the larval stage lasts from three to four weeks, and the pupal stage takes 12 to 36 days. At high temperatures, when development is more rapid, the life cycle can be completed in six or seven weeks. The life cycle of the granulate cutworm is similar to that of the black cutworm. They are active at night, feeding on the stems and leaves. During the day, they take refuge in the soil at the base of the plants. Larvae tend to curl up into a ring when disturbed or handled. They may also bite and release a greenish-brown fluid.

Damage

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Recently transplanted crucifers are particularly susceptible to attack by cutworms, which can cut thin-stemmed plants off at or slightly below the soil surface. They can also cut out large holes from leaves touching the soil. Several plants in a row are usually affected, and the cutworm often pulls the end of the leaf on which it is feeding into a protected area of the soil. Cutworms can also eat into heading cabbage and may remain within the head during the day. Overall, while some damage to leaves and heads occurs, greatest losses from cutworm damage are the result of reduced stands.

Black cutworms do most of their feeding at ground level. Larvae feed on young plants, cutting off leaves, or in later instars, entire plants. Populations of this pest tend to be higher in weedy and in wet fields. Granulate cutworm larvae can cut off entire seedling plants, as well as climb and feed on leaves of older plants. This cutworm is not associated with weedy fields as is the black cutworm. First instar larvae stay on plants, while older larvae climb and feed on plants only during night. Harlequin Bug

Murgantia histrionica (Hahn) (Hemiptera: Pentatomidae)

Description

Eggs are barrel-shaped, light gray or pale yellow and are encircled by two black bands. They are generally found beneath leaves in clusters of 12 arranged in two rows of six. Young nymphs (Figure 16) are first pale green with black markings but soon turn black or blue with red and yellow or orange markings. Adults (Figure 17) are also brightly colored, mainly black and yellow or black and red.

Figure 16.

Harlequin bug eggs and nymphs. Credit: Lyle J. Buss, University of Florida.

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Figure 17.

Harlequin bug adult. Credit: James L. Castner, University of Florida.

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Biology

Harlequin bug reproduces all year round in Florida. Females produce an average of 115 eggs during their lifetime. Eggs hatch in 4–5 days in warm weather. Newly hatched nymphs stay close to the eggs for one or two days. There are five or six instars. Development times that have been reported range from 30 to 70 days, depending on temperature. Adults can live about two months.

Damage

Harlequin bugs prefer crucifers although they will occasionally feed on other plants. They have piercing-sucking mouthparts and leave white blotches where they feed. Plant may wilt, become deformed, or die, if bugs are abundant. Brown Stink Bugs

Euschistus obscurus (Palisot de Beauvois), Euschistus servus (Say) (Hemiptera: Pentatomidae)

Description

Adult stink bugs are shield-shaped and vary from greyish-yellow to brown in color. They have numerous blackish spots scattered on the head, thorax and hardened part of the forewing. The shoulders are usually rounded with a light transverse band between them. In E. obscurus, the head and anterior part of the thorax have several black dots, making them darker than the rest of the body (Figure 18a). The last two segments at the tip of the antenna in E. servus are darker in color (Figure 18b). The underside is yellow or green. Eggs are somewhat elliptical in shape, semi-translucent, and slightly yellow but become pinkish upon reaching maturity. Eggs are laid in masses of up to 35 eggs on the underside of leaves. Small, rounded, and wingless nymphs, which otherwise resemble adults, emerge from the eggs and go through five instars molting to adults.

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Figure 18. Adult stink bug (a) Euschistus obscurus and (b) Euschistus servus. Credit: Lyle J. Buss, University of Florida.

Biology

Euschistus obscurus is distributed throughout the southern U.S. It has been reported in Florida, Georgia, Mississippi, and Texas. Euschistus servus is distributed in the southern Canada, part of North America, including the southern U.S. Both species are frequently found on cotton and soybean, but they feed on many different crops, including the crucifers mustard and bok choy.

Damage

Stink bugs have piercing-sucking mouthparts for feeding on the plant’s fluids. They feed on leaves, stems, and flowers. Feeding causes injury that results in discolored spots on the leaves due to tissue degrading enzymes injected during feeding. Feeding on growing points causes stunting and development of small heads in broccoli, cabbage, and cauliflower. Large numbers can weaken plants and plant pathogens can enter the punctures created where the bugs insert their mouthparts. Cross-striped Cabbageworm

Evergestis rimosalis (Guenée) (Lepidoptera: Crambidae)

Description

The adult moth (Figure 19) has a wingspan of about one inch. The front wings are straw-colored, marked with olive or purplish-brown, and crossed by narrow transverse lines. Hind wings are transparent and whitish, bordered with a darker band. Eggs, laid in small masses, are oval, yellow, and flattened, and overlap slightly. Larvae (Figure 20), which have 4 instars, are gray with black tubercles to begin with and become bluish-gray with numerous transverse black bands. There is a yellow line along each side of the caterpillar. The mature caterpillar is about 0.6–0.7 inches long. The pupa is yellowish-brown, enclosed in a small cocoon covered with sand.

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Figure 19.

Cross-striped cabbageworm adult. Credit: Lyle J. Buss, University of Florida.

[Click thumbnail to enlarge.]

Figure 20.

Cross-striped cabbageworm larva. Credit: Lyle J. Buss, University of Florida.

[Click thumbnail to enlarge.]

Biology

A crucifer specialist, the cross-striped cabbageworm is more of a problem on broccoli, cauliflower, collards, and Brussels sprouts than it is on kale and cabbage. Development time from egg to adult ranges from 61 days at 68°F to 18 days at 95°F. It can be abundant during the winter and spring cropping period in Florida. Larvae pupate in the soil, near the surface.

Damage

Larvae feed on leaves, creating small holes. They prefer terminal buds and may also burrow into the center of developing cabbage heads. Great Southern White

Ascia monuste (Linnaeus) (Lepidoptera: Pieridae)

Description

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The adult butterfly (Figure 21) has a wingspan that ranges 2.5 to 3.4 inches. The wings of males are white with a black margin in a zigzag pattern on the forewings. Female color varies from white to gray, and, like the males, they have a black margin and a small black spot on the forewings. Both sexes have turquoise antennal clubs. Yellow, spindle-shaped eggs are laid singly or in clusters on host leaves. Larvae (Figure 22) have yellow and gray longitudinal stripes and multiple small black spots along the body. This insect goes through five instars. Pupae are white or yellow with black markings.

Figure 21. Great southern white adult. Credit: Lyle J. Buss, University of Florida.

Figure 22.

Great southern white larvae. Credit: Lyle J. Buss, University of Florida.

Biology

The great southern white is a subtropical and tropical species, and it is mainly found in coastal areas. It is present year-round in southern Texas, peninsular Florida, and along the Gulf Coast. The developmental time from egg to adult emergence is approximately 32 days at 77ºF. It is considered an important pest of cole plants, including wild species such as beach cabbage (Cakile maritima Scopoli) and cultivated plants such as kale, cauliflower, broccoli, arugula, cabbage, and mustard.

Damage

Larvae chew the leaves of plants. They usually feed in groups. Newly emerged larvae consume their eggshells and eggs that have not hatched. Adults feed on nectar of many flowers.

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Yellowmargined Leaf Beetle

Microtheca ochroloma (Stal) (Coleoptera: Chrysomelidae)

Description

The dark bronze or black adult beetle (Figure 23) is about 0.2 inches long. The edges of the wing covers are bordered with yellow. Eggs are bright orange and elongate and are deposited singly or in small clusters in protected spots on the plant or in leaf litter. The soft-bodied larva is yellowish-brown and covered with a fine layer of hairs (Figure 24). The head is dark brown or black. The mature larva pupates in a loose, net-like case in folds of foliage, or in debris on the soil surface.

Figure 23.

Yellowmargined leaf beetle adult. Credit: Angie Niño, University of Florida.

[Click thumbnail to enlarge.]

Figure 24.

Yellowmargined leaf beetle larva. Credit: James Castner, University of Florida.

Biology

Yellowmargined leaf beetle is native to South America and was first found in the United States in 1947. It is established in the southeastern part of U.S. The life cycle is not well known in Florida. The beetle is capable of completing development in about one month and may be limited

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by food availability because it is restricted to crucifers. Adults are active all winter in Florida and may live from two to over three months. Turnips, radishes, and mustard are more preferred than cabbage and collards, and females that develop on the first three crops produce the most eggs (up to almost 500 eggs per female).

Damage

Yellowmargined leaf beetle is a particular problem for organic growers. It is especially devastating on specialty cruciferous greens like mizuna, but also feeds on turnips, mustard, and other cole crops. The spring crop may suffer more damage than the fall crop. Adults and larvae feed on leaves.

Tables

Table 1.

Diamondback moth

Management Recommendation Option Fields should be scouted weekly. One method is to walk a zigzag or figure eight path through the field in such a way that all four quarters of the field are checked and both interior and border areas are examined. Carry a hand lens (at Scouting/ least 10X) and a notebook and pencil to record observations. Small plastic bags Thresholds are useful for collecting insects for identification. Thresholds of 0.1 to 0.3 larvae per plant have been used in northeast Florida cabbage. Examine 50 to 100 plants per field (stop 10 to 20 times and examine 5 plants in each location). Diamondback moth develops resistance to insecticides easily, particularly pyrethroids. Rotation of insecticide modes of action (MoA) and avoidance of pyrethroids are important for managing diamondback moth. There are at least three types of parasitic wasp in Florida that attack either the larval or pupal stage of diamondback moth. Early season reliance on Bacillus thuringiensis (Bt) products does not interfere with the activity of these natural enemies and can offset the severity of infestations. Diamondback moth has developed resistance to Bt products in some regions; Notes however, Bts remain useful tools for controlling young larvae. It is advised that application of products with the aizawai strain of Bt (i.e. Agree WG, XenTari DF) be alternated with products formulated with the kurstaki strain of Bt (i.e. Biobit HP, Crymax WDG, Dipel DF, Javelin WG). Insect growth regulators (IGRs) that can be used to manage diamondback moth larvae include Rimon (novaluron, MoA Group15) and Intrepid (methoxyfenozide, MoA Group 18). IGRs are slow acting but are useful management tools. Avaunt (indoxacarb, MoA 22) is another important rotation tool for caterpillar management.

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Coragen is a systemic diamide insecticide (MoA Group 28) that can be applied at-plant or through the drip irrigation as well as to foliage to provide protection against diamondback moth and other caterpillars. The active ingredient in Coragen is chlorantraniliprole, which is one of the ingredients in Durivo. Durivo also contains the neonicotinoid thiamethoxam (MoA Group 4A), which provides protection against sucking pests such as whiteflies and aphids. Verimark is another Group 28 insecticide containing cyantraniliprole that is applied in the plant tray, at-planting or through the drip. It provides protection against caterpillars as well as whiteflies and leafminers. Other Group 28 foliar insecticides that can be used to manage diamondback moth include Belt and Exirel. The active ingredient in Belt is flubendiamide, which has been withdrawn by the EPA. Growers may use material that they have on hand. Flubendiamide is also in Vetica, which contains buprofezin to control whitefly nymphs. Exirel contains cyantraniliprole and is also effective against whiteflies and leafminers. The window treatment approach should be used for applying group 28 insecticides to cabbage and other crucifers. If Group 28 insecticides are applied at-planting and during the first five-week treatment window, they should not be applied during the second five-week treatment window. A parasitoid wasp helps control diamondback moth larvae, especially if Bt is Natural the main pesticide used. An egg parasitoid (Trichogramma sp.) and fungal Enemies pathogens also aid in control. Important cultural controls include avoiding the warmer months, and destroying crop residues. Another method of managing diamondback on Cultural cabbage is to plant several rows of collards around the perimeter of the fields Controls as a trap crop to be treated with insecticides. For high-value, specialty crucifers, floating row covers put in place immediately after transplanting may eliminate damage. Table 2.

Cabbage looper

Management Recommendation Option Scouting/ Fields should be scouted weekly. See diamondback moth, above. Thresholds Relying on Bts as the main insecticide and using some of the more specific Notes pesticides (Entrust (spinosad MoA Group 5), Avaunt, Intrepid, Coragen) when needed, will help preserve natural enemies. Natural Parasitoid wasps and flies and general predators help control cabbage looper. Enemies A nucleopolyhedrosis virus also kills loopers. Avoid the warmer months when pests are most abundant, destroy crop Cultural residues, and control weeds. Planting a nectar source for beneficial insects may Controls be helpful—sweet alyssum has been tested in cabbage. For high-value,

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specialty crucifers, floating row covers put in place immediately after transplanting may eliminate damage. Table 3.

Aphids

Management Recommendation Option Scouting/ Fields should be scouted weekly. See diamondback moth, above. No Thresholds information on thresholds is available. Relying on Bts as the main insecticide for caterpillar pests and using some of the more specific pesticides (Entrust, Avaunt, Intrepid, Coragen, and others) when needed, will help preserve natural enemies of aphids. Selective Notes insecticides for controlling aphids include Beleaf (flonicamid MoA Group 9C), Fulfill (pymetrozine MoA Group 9B), and Movento (spirotetramat MoA Group 23). Group 4A products (neonicotinoids) will also control aphids. Parasitoid wasps and general predators, such as ladybeetles, lacewing larvae, Natural and syrphid fly larvae, may completely control aphids if broad spectrum Enemies pesticides can be avoided. Destroy crop residues and control cruciferous weeds. Planting a nectar source for beneficial insects may be helpful—sweet alyssum has been tested in Cultural cabbage. For high-value, specialty crucifers, floating row covers put in place Controls immediately after transplanting may eliminate damage. However, if any aphids are trapped beneath covers, they will multiply freely in the absence of their natural enemies. Table 4.

Beet armyworm

Management Recommendation Option Fields should be monitored at least weekly for damage by caterpillars feeding on leaves. Pheromone traps can be used to monitor occurrence of moths. Young plants are more susceptible to damage. Look for egg masses on the Scouting/ leaves. Look toward the base of leaves for damage and under outer leaves near Thresholds the soil surface for larvae that may hide during the day away from their feeding site. Treat if you find 0.3 larvae per plant, using the sampling procedure described for diamondback moth. Best time to treat for this pest is in early morning or early evening. Insecticides are most effective against the younger instars, with higher rates Notes and more frequent applications needed to try to control later instars. Insecticides are available for foliar applications. Coverage and penetration are

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important in treating for all of the moth species. Use of surfactants to increase surface coverage increases insecticidal control. Natural Insect predators and parasitoids, as well as pathogens aid in control, but Enemies generally do not exert enough pressure to prevent yield loss. Field disking and destruction of crop residues are important for control of all caterpillar and aphid pests to reduce their migration into nearby crops. Beet Cultural armyworms develop well on several weeds in the Amaranth group, so weed Controls control on ditch banks surrounding fields can help reduce populations before they invade fields. Table 5.

Cabbage webworm

Management Recommendation Option Scouting/ Fields should be scouted weekly. See diamondback moth, above. Thresholds Relying on Bts as the main insecticide and using some of the more specific pesticides (Entrust, Avaunt, Intrepid, and others) when needed, will provide control. Younger larvae, less protected by webbing and folded leaves, should Notes be targeted. Early and frequent scouting is critical. Treatment should begin as soon as larvae are found because of their habit of feeding on the terminal bud, which interferes with proper head formation. Natural No important natural enemies are known. Enemies Avoid the warmer months when pests are most abundant, destroy crop residues, and control weeds. Planting a nectar source for beneficial insects may Cultural be helpful—sweet alyssum has been tested in cabbage. For high-value, Controls specialty crucifers, floating row covers put in place immediately after transplanting may eliminate damage. Table 6.

Cutworms

Management Recommendation Option Seedling crops should be scouted as frequently as twice per week to detect cutworms or their damage, particularly in areas known for this pest. Young Scouting/ larvae may be found grouped together on foliage, but older larvae will usually Thresholds be found in soil or beneath leaf trash during the day. Look for wilted foliage or plants with severed stems. Adults can be monitored with black light and pheromone traps.

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Insecticides are available for at-plant, pre- and post-emergence broadcast and Notes banded applications. Post emergence applications are the most efficient. Natural enemies such as parasitic wasps, flies, and predacious ground beetles can exert tremendous control pressure that may approach 80%. However, Natural seedlings emerging in fields without resident natural enemy populations can Enemies experience significant stand loss from first generation cutworms. Larvae are also targets for attack by pathogenic fungi and viruses. Weedy fields quickly rotated to leafy vegetables have higher potential for stand Cultural loss due to surviving older larvae cutting off the emerging plants. Therefore, Controls prepare fallowed fields for production as soon as possible to allow time for surviving larvae to complete development before planting. Table 7.

Cross-striped cabbageworm

Management Recommendation Option Scouting/ Fields should be scouted weekly. See diamondback moth, above. Thresholds Relying on Bts as the main insecticide and using some of the more specific Notes pesticides (Entrust, Avaunt, Intrepid, and others) when needed, will help preserve natural enemies. Natural Parasitoid wasps and possibly general predators help control cross-striped Enemies cabbageworm. Avoid the warmer months when pests are most abundant, destroy crop residues, and control weeds. Planting a nectar source for beneficial insects may Cultural be helpful—sweet alyssum has been tested in cabbage. For high-value, Controls specialty crucifers, floating row covers put in place immediately after transplanting may eliminate damage. Table 8.

Stink bugs

Management Recommendation Option Scouting/ Visual examination of plants and yellow pyramid traps lured with stink bug Thresholds aggregation pheromone can be used to monitor and capture them. The use of synthetic insecticides, primarily pyrethroids, but also some neonicotinoid insecticides (Group 4A), is the most effective control method. Notes Some pesticides (Pyganic (pyrethrins MoA Group 3A) and azadirachtin) can be used in organic farming. The tolerance of stink bugs to most insecticides makes suppression difficult.

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Natural Eggs and nymphs suffer high mortality caused by pathogens and predators Enemies such as ladybeetles, lacewings, and predatory stink bugs. Triticale, buckwheat, sorghum, millet, and sunflower can be used as trap crops by planting them on the periphery of the main crop to intercept stink bugs Cultural before they enter the cash crop. Trap crops should be treated with insecticides Controls once stinkbugs are detected or when the trap crop becomes unattractive to stop them from building up and moving into the main crop.

Table 9.

Yellowmargined leaf beetle

Management Recommendation Option Visual examination of plants looking for larvae and adults or the damage Scouting/ caused by them. Eggs and pupae are more frequently found under fallen leaves Thresholds or at the base of the plant. The use of Pyganic and Entrust, which are allowed in organic farming, is the Notes most effective control method. Natural General predators such as ladybeetles, lacewings, and predatory stink bugs Enemies help controlling eggs, larvae, and adults of the yellowmargined leaf beetle. Turnips can be used as a trap crop when growing less attractive cruciferous crops such cabbage and collards. Turnips should be planted at the periphery of Cultural main crop two weeks in advance. The trap crop should be treated with Controls insecticides once adults or larvae are detected to stop them from building up and moving into the main crop.

Footnotes

This document is ENY-464, one of a series of the Entomology & Nematology Department, Florida Cooperative Extension Service, Institute of Food and Agricultural Sciences, University of Florida. Published July 2002. Revised March 2010 and June 2013. For more publications related to horticulture/agriculture, please visit the EDIS website at http://edis.ifas.ufl.edu/.

S. E. Webb, associate professor, Entomology and Nematology Department, Cooperative Extension Service, Institute of Food and Agricultural Sciences, A. Nino, doctoral candidate, Entomology and Nematology Department, University of Florida, Gainesville, FL 32611-0640, and H. A. Smith, Assistant Professor, Gulf Coast Research and Education Center, University of Florida, Balm, FL 33598.

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The use of trade names in this publication is solely for the purpose of providing specific information. UF/IFAS does not guarantee or warranty the products named, and references to them in this publication does not signify our approval to the exclusion of other products of suitable composition. All chemicals should be used in accordance with directions on the manufacturer's label. Use pesticides safely. Read and follow directions on the manufacturer's label.

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APPENDIX F EDIS PUBLICATION SPANISH VERSION Manejo de Insectos en Crucíferas (Cultivos de Coles) (Brócoli, Repollo, Coliflor, Col, Col 1 Rizada, Mostaza, Rábano, Nabos)

S. E. Webb2, A. Niño2, H. A. Smith2

Los vegetales conocidos como crucíferas son un grupo de cultivos amplio y cada vez más importante en Florida. Un número de insectos se alimenta exclusivamente de crucíferas y afecta todos los cultivos enlistados en el título. Debido a que la mayoría de nuevos pesticidas han sido etiquetados para todo el grupo de cultivos o para un subconjunto, ya sean brásicas de cabeza o tallo (como repollo o brócoli) o brásicas de hojas (como la col rizada), no se incluyen tablas específicas para cultivos individuales de brásicas al final de este capítulo. En cambio, las excepciones están incluidas en la columna de notas. Otras crucíferas, no enumeradas en el título, que albergan el mismo complejo de plagas incluyen brásicas de cabeza o tallo tales como coles de Bruselas, brócoli chino y mostaza china, y brásicas de hojas como col china, mizuna y hojas de nabos. Revise cuidadosamente las etiquetas de los pesticidas para cerciorarse que el cultivo está incluido. Los rábanos y los tubérculos de nabos están incluidos en el grupo de vegetales de raíz, a pesar de que también son crucíferas y tiene problemas de plagas similares. Una tabla separada de pesticidas para rábano está incluida al final de este documento.

La palomilla de dorso de diamante es la plaga principal de crucíferas en la Florida. El gusano falso medidor también se considera una plaga importante, sin embargo, ha causado menores problemas en la última década. Insectos plaga que han sido considerados importantes en el pasado y ahora causan problemas ocasionales incluyen: áfidos (pulgón del repollo y pulgón verde), el chinche arlequín de la col, el gusano soldado, el gusano perforador de las coles y el gusano trozador (negro y granulado). El escarabajo de margen amarillo es un problema particular en mostaza y repollo chino, especialmente para productores orgánicos. El gusano de franjas cruzadas causa más problemas en brócoli y coliflor que en otras crucíferas. Áfidos, gusanos trozadores y gusanos alambre son los principales insectos plaga que afectan rábano. La Palomilla de Dorso de Diamante

Plutella xylostella (Linnaeus) (Lepidoptera: Plutellidae)

Descripción

El adulto de la palomilla (Figura 1) es pequeño y delgado con antenas muy largas. Es de color café grisáceo y tiene una banda amplia de color crema o café claro en su parte dorsal. La banda tiene contriciones, las cuales forman un patrón en forma de diamante. Cuando se observa lateralmente, la punta de las alas parece alargarse ligeramente hacia arriba. Los huevos son de

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forma ovalada y aplanada, de color amarillo a verde pálido, y miden aproximadamente 0.02 pulgadas de largo y 0.01 pulgadas de ancho. Hay cuatro estadios larvales. Incluso las larvas en el último estadio son bastante pequeñas y muy activas. Las larvas se retuercen violentamente si se les molesta y se dejan caer de la hoja quedando suspendidas por un hilo de seda. El cuerpo se estrecha hacia ambos extremos y el quinto par de pseudopatas (patas abdominales) sobresale de la parte posterior (Figura 2). Después del primer estadio, que es incoloro, las larvas obtienen una coloración verdosa. Las larvas empupan dentro de un capullo suelto en las hojas inferiores o externas o en las inflorescencias de coliflor y brócoli.

Figura 1.

Vista superior y lateral de la palomilla dorso de diamante. Crédito: Lyle J. Buss, Universidad de la Florida.

[Haga click en la figura para maximizar.]

Figura 2.

Larva de la palomilla dorso de diamante. Crédito: Lyle J. Buss, Universidad de la Florida.

[Haga click en la figura para maximizar.]

Biología

La hembra deposita sus huevos sobre el envés de la hoja, de forma individual o en grupos de dos o tres. Al cabo de unos días, los huevos eclosionan, y las larvas comienzan a alimentarse del envés de la hoja. La etapa larval puede durar de 10 días a un mes, dependiendo de la temperatura. Larvas de la palomilla dorso de diamante se alimentan más lento a temperaturas por debajo de 50°F, y el crecimiento de la población es más rápido a temperaturas superiores a 80°F. El estado de pupa ocurre dentro de un capullo holgado y transparente el cual es usualmente

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adherido al envés de las hojas. En climas cálidos, el estado de pupa puede completar su desarrollo en 3 o 4 días.

En el sur de la Florida, la palomilla de dorso de diamante es más abundante durante los meses de Diciembre a Febrero o Marzo y puede atacar en cualquier momento durante el ciclo del cultivo. Para finales de Mayo, el conteo de palomillas en trampas de feromonas se disminuye a casi cero. El conteo de palomillas puede aumentar de mitad de otoño hasta comienzos del invierno, pero su actividad es limitada durante esa época. Las poblaciones se desarrollan en malezas silvestres, como la mostaza silvestre, antes de trasladarse a las plantaciones de repollo y otras crucíferas plantadas en invierno y a comienzos de la primavera. Desde mitades de invierno y durante la primavera, cuando esta plaga causa daños severos, la palomilla de dorso de diamante puede causar pérdidas de hasta 70% en la ausencia de medidas de control. Las poblaciones pueden disminuir después de lluvias fuertes.

Daño

Plantas en cualquier estadio de crecimiento pueden ser atacadas. Las larvas producen pequeños hoyos en las hojas, larvas más grandes hacen hoyos más grandes. Las larvas jóvenes con frecuencia se alimentan de una de las superficies de la hoja dejando una capa delgada similar a una “ventana” en la epidermis de la hoja. Las larvas de la palomilla de dorso de diamante también atacan la cabeza del repollo en desarrollo. El daño resultante deforma la cabeza del repollo y permite la entrada de patógenos descomponedores. Gusano Falso Medidor

Trichoplusia ni (Hübner) (Lepidoptera: Noctuidae)

Descripción

Figura 3.

Macho adulto del gusano falso medidor. Crédito: Lyle J. Buss, Universidad de la Florida. [Haga click en la figura para maximizar.]

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Figura 4.

Gusano falso medidor. Crédito: Lyle J. Buss, Universidad de la Florida.

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El gusano falso medidor se alimenta de una amplia variedad de cultivos. Los adultos (Figura 3) son polillas nocturnas con alas anteriores de color café, moteadas y marcadas en el centro con una figura pequeña, plateada y en forma de ocho. Los huevos son pequeños, acanalados, redondos y de color blanco verdoso. Los huevos eclosionan en larvas de color verde con líneas blancas que recorren la longitud del cuerpo. El gusano (Figura 4) tiene tres pares de patas delgadas cerca de su cabeza y tres pares de pseudopatas gruesas en el extremo final del cuerpo. Éste se mueve con un movimiento ondulatorio característico, alternadamente se mueve hacia adelante y, arqueando su parte dorsal, trae las pseudopatas traseras cerca de sus patas delanteras. El gusano mide alrededor de 1.25 pulgadas cuando está completamente desarrollado.

Biología

Los huevos son depositados individualmente o en pequeños grupos en cualquiera de las superficies de la hoja, sin embargo, la mayoría son encontrados en el envés. Cada hembra puede producir de 300 a 600 huevos durante los 10 a 12 días, aproximadamente, que está viva. Dos a cuatro semanas después de la emergencia, la larva madura teje un capullo delgado en el envés de la hoja, sobre los residuos de plantas o el suelo. El estado de pupa dura aproximadamente dos semanas. El tiempo total requerido para completar el desarrollo de huevo a adulto es, en promedio, 42 días a 69.8°F y 22 días a 89.6°F. Desarrollo anormal ocurre por encima de 94°F.

Las poblaciones tienden a ser más altas a finales de la primavera y en los meses de verano, y en algunos años, a finales del otoño. El gusano falso medidor no entra en diapausa y no puede sobrevivir durante periodos prolongados de clima frío. Los insectos permanecen activos y se reproducen durante los meses de invierno solo en la parte sur de la Florida (más al sur de Orlando). En Florida central, las poblaciones del gusano falso medidor se incrementan durante inicios de otoño y de nuevo a finales de la primavera.

Daño

El gusano falso medidor es todavía una plaga anualmente importante en crucíferas al norte de la Florida. Es menos problemático en el sur de la Florida donde es considerado una plaga menor.

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En general, el gusano falso medidor causa más problemas durante el otoño que durante los meses de invierno o primavera.

Las larvas del gusano falso medidor dañan las plantas al producir hoyos en las hojas. Larvas pequeñas permanecen en el envés de la hoja, mientras larvas grandes producen hoyos más grandes a través de la hoja. Adicional al daño al alimentarse de las hojas que envuelven el repollo, las larvas pueden perforar dentro de la cabeza en desarrollo. Cierta defoliación es tolerada antes de la formación de la cabeza, pero el daño por alimentación y los excrementos dejados en las cabezas hace que el repollo no pueda ser comercializado. El repollo con daño confinado a las hojas externas es comercializable pero con un menor valor. Áfidos

Áfido del nabo [Lipaphis erysimi (Kaltenbach)], el pulgón verde del melocotero [Myzus persicae (Sulzer)], y el áfido del repollo [Brevicoryne brassicae (Linnaeus)]

Descripción

El áfido del nabo (Figura 5) y el pulgón verde del melocotero (Figura 6) son los áfidos más importantes en crucíferas en Florida. El áfido del repollo (Figura 7) no es tan común en la Florida. A pesar de que los problemas con áfidos en la Florida tienden a ser esporádicos, ellos siguen en importancia a la palomilla de dorso de diamante y al gusano falso medidor. Los adultos son de cuerpo blando, con el cuerpo en forma de pera, y tienen un par de túbulos (cornículos o sifúnculos) en la parte posterior del cuerpo, los cuales se proyectan hacia la parte superior trasera desde la parte dorsal del abdomen y son usados para excretar un feromona de alarma. Los áfidos tienen un aparato bucal picador chupador con el cual penetran el tejido de la hoja para alimentarse de la savia del floema. Las ninfas son más pequeñas pero similares en apariencia a los adultos que carecen de alas.

Figura 5.

Áfido del nabo. Una apariencia inflada y de color pardo pálido indica que el áfido ha sido parasitado por avispas parasíticas. Crédito: James Castner, Universidad de la Florida.

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Figura 6.

Áfido verde del melocotero. Crédito: Lyle J. Buss, Universidad de la Florida.

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Figura 7.

Áfido del repollo. Crédito: Lyle J. Buss, Universidad de la Florida.

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Los adultos del áfido del nabo son verdes o verde blancuzcos y miden alrededor de 0.06-0.09 pulgadas de largo. Las antenas son oscuras y los cornículos son pálidos con la punta oscura. El cuerpo está cubierto de una secreción blanca. Las ninfas son de color amarillo verdoso pálido. Los adultos del áfido verde del melocotero varían de 0.04 a 0.08 pulgadas de largo y su color varía de verde claro, a amarillo o rosado y tienen forma de pera. Los tubérculos (protuberancias en medio de las antenas) apuntan hacia adentro y son una característica distinguida. Las formas aladas tienen una mancha negra en la parte trasera del abdomen. Los adultos del áfido del repollo son muy similares en apariencia al áfido del nabo, sin embargo, son más grandes (0.08-0.1 pulgadas de largo) con cornículos cortos y están cubiertos con un polvo ceroso grisáceo.

Biología

Los áfidos se reproducen muy rápido. En Florida, los machos son poco comunes y las hembras dan a luz a ninfas vivas durante todo el año sin aparearse o producir huevos. Las ninfas maduran en 7 a 10 días. Cuando la calidad de la planta hospedera se ha deteriorado o si las plantas están infestadas por muchos insectos, formas con alas se desarrollan y migran a nuevas plantas hospederas. Los áfidos son muchas veces protegidos de sus enemigos naturales por hormigas, las

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cuales se alimentan de la sustancia azucarada producida por los áfidos llamada mielecilla. Los áfidos son más abundantes durante la primavera y el otoño y casi desaparecen en el verano.

Daño

El áfido verde del melocotero es una plaga importante de verduras (col, col rizada y mostaza), y también de muchos otros cultivos no relacionados. Esta especie ataca el repollo principalmente antes que la formación de la cabeza ocurra. Los áfidos del nabo atacan solo crucíferas, prefiriendo el nabo y el rábano. Los áfidos chupan los jugos de la planta con sus aparatos bucales tipo picador chupador, resultando en amarillamiento y enroscamiento de las hojas. La planta, particularmente cuando es atacada en el estado de plántula, puede quedarse enana o morir como resultado de la alimentación de los áfidos. El follaje puede contaminarse con cuerpos de los áfidos muertos, la piel que queda después de la muda y la mielecilla. Los áfidos que se alimentan dentro de las hojas enroscadas o deformadas están normalmente protegidos del contacto con insecticidas. El áfido verde del melocotero y el áfido del nabo transmiten el virus del mosaico del nabo en la Florida. Gusano Soldado

Spodoptera exigua (Hübner) (Lepidoptera: Noctuidae)

Descripción y Biología

El gusano soldado tiene un amplio rango de hospederos. Además de crucíferas, este insecto ataca vegetales como espárragos, fríjoles, remolacha, apio, garbanzo, maíz, caupí, berenjena, lechuga, cebolla, arveja, pimentón, papa, espinaca, camote y tomate. También se alimenta de muchos otros cultivos y malezas.

La polilla adulta (Figura 8), de gran movilidad, tiene las alas anteriores oscuras moteadas con marcas más claras y las alas posteriores están finamente cubiertas por escamas blancuzcas. Cada hembra puede producir alrededor de 600 huevos, generalmente en masas de alrededor de 100, en el envés de las hojas de la parte baja del dosel. Las masas de huevos están cubiertas por escamas blancas y difusas. Los gusanos jóvenes (Figura 9), los cuales son de color pálido con cabeza oscura, se alimentan en grupos y luego se dispersan a medida que envejecen (tercer estadio). Para el tercer estadio, los gusanos tienen unas líneas ondulantes de color claro que recorren la longitud del cuerpo sobre la parte dorsal y unas rayas amplias en cada lado. Aunque generalmente son verde opaco, el color de los gusanos puede variar. Luego de alimentarse por 1 a 3 semanas, ellos construyen un capullo usando arena y pedazos de suelo y empupan en el suelo, emergiendo como adultos una semana después. El gusano soldado es un insecto tropical y sobrevive durante el verano en el sur de la Florida donde puede completar varias generaciones por año. Del sur de la Florida, adultos migran al norte de la Florida y otras partes del sureste.

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Figura 8.

Adulto del gusano soldado. Crédito: Lyle J. Buss, Universidad de la Florida.

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Figura 9.

Larva del gusano soldado. Crédito: Lyle J. Buss, Universidad de la Florida.

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Daño

Las larvas se alimentan del follaje de la planta hospedera. Las larvas del gusano soldado consumen una mayor cantidad de tejido foliar que las palomillas de dorso de diamante, pero no tanto como el gusano falso medidor. Un umbral de acción de 0.3 larvas del gusano soldado por planta ha sido usado en repollo en Texas. Debido a que los adultos pueden invadir el cultivo desde otros cultivos cercanos o malezas, se recomienda el monitoreo del cultivo dos veces por semana en búsqueda de la presencia o daños del gusano soldado.

El gusano soldado es una plaga esporádica en crucíferas en Florida, y usualmente es mantenida por debajo de los niveles de daño a través de los controles implementados para la palomilla del dorso de diamante. Puede volverse un problema serio en repollo de napa en el norte de Florida en la primavera. Las poblaciones del gusano soldado en el sur de Florida son mayores desde finales de Marzo hasta la mitad de Junio, con pequeñas poblaciones ocurriendo desde mitad de Agosto hasta Octubre. El incremento a final del verano y otoño se considera que está relacionado con actividad del gusano soldado sobre malezas a finales del verano, mientras que el incremento de las poblaciones en primavera coincide con la etapa de producción de vegetales en el sur de la Florida.

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Gusano Perforador de las Coles

Hellula rogatalis (Hulst) (Lepidoptera: Crambidae)

Descripción

La polilla (Figura 10) tiene las alas anteriores de color amarillo-café con bandas blancas y un punto negro en forma de riñón. Las alas posteriores son blancas grisáceas con un margen oscuro. La envergadura de las alas es de alrededor de 0.7-0.8 pulgadas. Los huevos tienen una forma aplanada y son de color gris o amarillo-verdoso en un inicio, pero luego se tornan rosados a medida que se acercan a la emergencia. Hay cinco instares larvales. La larva madura (Figura 11) es amarillo-grisáceo con cinco bandas café-púrpura que recorren la longitud del cuerpo. La cabeza es negra. El cuerpo está cubierto de pelos moderadamente largos de color amarillo o café claro.

Figura 10.

Adulto del gusano perforador de las coles. Crédito: Lyle J. Buss, Universidad de la Florida.

[Haga click en la figura para maximizar.]

Figura 11.

Larva del gusano perforador de las coles. Crédito: Lyle J. Buss, Universidad de la Florida.

[Haga click en la figura para maximizar.]

Biología y Daño

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Los huevos del gusano perforador de las coles son usualmente ovipositados individualmente o en pequeñas masas en las hojas terminales. Apenas emergen, las larvas penetran las hojas y también se alimentan del envés, produciendo unos hoyos pequeños. Cuando alcanzan el tercer estadio, las larvas construyen una telaraña y doblan el follaje. Las telarañas son cubiertas de mugre y excremento. Larvas grandes son capaces de penetrar los brotes, tallos y hojas. El insecto puede alimentarse de los puntos de crecimiento de la planta, causando un daño severo a plantas jóvenes. Cuando las larvas se han desarrollado por completo, empupan en los brotes, sobre los tallos o en la superficie del suelo.

Similar al gusano soldado, el gusano perforador de las coles ocurre esporádicamente y es controlado por las medidas implementadas para la palomilla de dorso de diamante. La especie relacionada, H. phidilealis (Walker), o el gusano del brote de la col, puede ser problemático en el sur de la Florida. Gusanos Trozadores

Gusano trozador negro [Agrotis ipsilon (Hufnagel)] y el gusano trozador granulado [Feltia subterranea (Fabricius)] (Lepidoptera: Noctuidae)

Descripción

Las polillas del gusano trozador negro (Figura 12) son grandes, con una envergadura de 1.5 a 2.0 pulgadas. Las alas anteriores son café oscuro con una línea clara cerca del final de cada ala. Las alas posteriores varían entre blanco y gris. Los huevos acanalados son en un inicio blancos, y luego se tornan café y son usualmente depositados en grupos. Las larvas (Figura 13) son gusanos robustos, de color gris y con un aspecto grasoso. El gusano trozador negro tienen numerosos gránulos oscuros y gruesos en la mayor parte de su cuerpo.

Figura 12.

Adulto de gusano negro trozador. Crédito: John Capinera, Universidad de la Florida.

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Figura 13.

Larva del gusano trozador negro. Crédito: John Capinera, Universidad de la Florida.

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Las polillas del gusano trozador granulado (Figura 14) son más pequeñas, con una envergadura de 1.2 a 1.7 pulgadas. Las alas anteriores son frecuentemente amarillo-café y tienen manchas distintivas en forma de fríjol en el centro. Las alas posteriores son blancas en su mayoría. Los huevos son semi-esféricos y acanalados. Al igual que los huevos del gusano trozador negro, éstos son inicialmente blancos y se oscurecen con el tiempo. Las larvas (Figura 15) varían de color gris a rojizo. Cada segmento abdominal tiene una mancha oblicua de color amarillo opaco. Una línea débil de color gris se presenta a lo largo de la longitud del cuerpo con manchas blancas o amarillas.

Figura 14.

Adulto del gusano trozador granulado. Crédito: Lyle J. Buss, Universidad de la Florida.

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Figura 15.

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Larva del gusano trozador granulado. Crédito: Lyle J. Buss, Universidad de la Florida.

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Biología

El gusano trozador negro es uno de los trozadores más destructivos y ataca una amplia variedad de plantas. Aunque las larvas del trozador pueden migrar al cultivo desde áreas adyacentes, la mayoría de la dispersión ocurre por adultos volando al cultivo. La polilla deposita los huevos en grupos de uno a 30 sobre hojas, tallos, o residuos del cultivo cerca al nivel del suelo. El estado de huevo dura de 5 a 15 días, el estado de larva dura de 3 a 4 semanas, y el estado de pupa toma 12 a 36 días. A temperaturas altas, cuando el desarrollo es más rápido, el ciclo de vida puede ser completado en 6 o 7 semanas. El ciclo de vida del gusano trozador granulado es similar al del gusano trozador negro. Ellos son activos en la noche, se alimentan de hojas y de tallos. Durante el día, se refugian en el suelo a la base de las plantas. Las larvas tienden a enroscarse cuando son perturbadas o manipuladas. Ellas pueden morder y secretar un fluido verde café.

Daño

Crucíferas recién trasplantadas son particularmente susceptible al ataque por gusanos trozadores, los cuales pueden cortar plantas con tallos delgados a nivel o por debajo el nivel de la superficie del suelo. Ellos también pueden cortar hoyos grandes en las hojas que están en contacto con el suelo. Muchas plantas en la misma línea son usualmente afectadas, y los gusanos trozadores regularmente halan la punta de la hoja de la que se están alimentando a un área protegida del suelo. Los gusanos trozadores también pueden alimentarse de la cabeza del repollo y pueden permanecer dentro de la cabeza durante el día. En general, mientras algún daño a las hojas y tallos puede ocurrir, las pérdidas mayores ocasionadas por los gusanos trozadores son el resultado de una reducción en la composición uniforme de las plantas.

Los gusanos trozadores negros se alimentan principalmente al nivel del suelo. Las larvas se alimentan de plantas jóvenes, cortando hojas o, en los estadios más avanzados, plantas completas. Las poblaciones de esta plaga tienden a ser mayores en áreas con malezas y en cultivos con alta humedad. Las larvas del trozador granulado pueden cortar plántulas por completo, y también pueden trepar y alimentarse de hojas en plantas más grandes. Este trozador no está asociado a cultivos con malezas como el trozador negro. Larvas de primer estadio permanecen sobre las plantas, mientras que larvas más viejas trepan y se alimentan sobre las plantas solo durante la noche. Chinche Arlequín

Murgantia histrionica (Hahn) (Hemiptera: Pentatomidae)

Descripción

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Los huevos son en forma de barril, de color gris claro o amarillo pálido y están rodeados por 2 bandas negras. Ellos son generalmente encontrados debajo de las hojas en grupos de 12 organizados en 2 líneas de 6. Las ninfas jóvenes (Figura 16) son en un principio verde pálidas con marcas negras, pero pronto se vuelven negras o azules con marcas rojas y amarillas o naranjas. Los adultos (Figura 17) son también de colores vivos, principalmente negro con amarillo o negro con rojo.

Figura 16.

Huevos y ninfas del chinche arlequín. Crédito: Lyle J. Buss, Universidad de la Florida.

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Figura 17.

Adulto del chinche arlequín. Crédito: James L. Castner, Universidad de la Florida.

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Biología

El chinche arlequín se reproduce todo el año en Florida. Las hembras producen en promedio 115 huevos durante su vida. Los huevos emergen en 4 a 5 días en climas cálidos. Las ninfas recién emergidas permanecen cerca de los huevos por 1 o 2 días. Las ninfas pasan por 5 o 6 estadios. Los tiempos de desarrollo que han sido reportados varían entre 30 y 70 días, dependiendo de la temperatura. Los adultos pueden vivir alrededor de dos meses.

Daño

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El chinche arlequín prefiere crucíferas, sin embargo, se alimenta ocasionalmente de otras plantas. Tiene un aparato bucal picador chupador y deja manchas blancas donde se alimenta. Las plantas pueden marchitarse, deformarse o morir, si los chinches son abundantes. Chinches Hediondas

Euschistus obscurus (Palisot de Beauvois) y Euschistus servus (Say) (Hemiptera: Pentatomidae)

Descripción

Las chinches adultos tienen forma de escudo y varían en color de gris-amarillo a café. Ellas tienen numerosos puntos negros dispersados en la cabeza, tórax y la parte endurecida de las alas anteriores. Los hombros son usualmente redondeados con una banda clara transversal entre ellos. La cabeza y la parte anterior del tórax de E. obscurus tienen varios puntos negros, haciéndolos más oscuros que el resto del cuerpo (Figura 18a). Los últimos dos segmentos en la punta de la antena en E. servus son oscuros (Figura 18b). La parte ventral es amarillo o verde. Los huevos son elípticos, semi-translúcidos, y un poco amarillentos, pero toman una coloración rosada a medida que maduran. Los huevos son puestos en masas de hasta 35 huevos en el envés de las hojas. Las ninfas son similares en forma a los adultos, pero más pequeñas, redondeadas y sin alas. Emergen de los huevos y pasan por cinco estadios hasta que mudan a adultos.

Figura 18. Chinches hediondas adultos (a) Euschistus obscurus y (b) Euschistus servus. Crédito: Lyle J. Buss, Universidad de la Florida.

Biología

Euschistus obscurus está distribuido a lo largo del sur de los Estados Unidos. Las chinches han sido reportadas en Florida, Georgia, Mississippi y Texas. Euschistus servus está distribuido en el sur de Canadá hasta el sur de los Estados Unidos. Ambas especies son frecuentemente encontradas en algodón y soya, pero ellos se alimentan de varios cultivos, incluyendo crucíferas como la mostaza y el bok choy.

Daño

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Las chinches tienen un aparato bucal picador chupador para alimentarse de los fluidos de la planta. Ellas se alimentan de hojas, tallos y flores. Este tipo de alimentación causa daños que resultan en puntos descoloridos en las hojas debido a enzimas que son inyectadas durante el proceso de alimentación y degradan los tejidos de la planta. Alimentación en los puntos de crecimiento causa retrasos en el desarrollo y la formación de cabezas pequeñas de brócoli, repollo y coliflor. Grandes números de chinches pueden debilitar las plantas y patógenos de plantas pueden entrar a través de las perforaciones creadas por las chinches cuando insertan sus aparatos bucales. Gusano de Franjas Cruzadas

Evergestis rimosalis (Guenée) (Lepidoptera: Crambidae)

Descripción

La polilla adulta (Figura 19) tiene una envergadura alrededor de una pulgada. Las alas anteriores son de color paja con marcas de color verde oliva o purpura-café, y con líneas delgadas transversas. Las alas posteriores son transparentes y blancas, con una banda oscura en el borde. Los huevos, ovipositados en pequeñas masas, son ovalados, amarillos y aplanados y se superponen ligeramente. Las larvas (Figura 20), las cuales pasan por cuatro estadios, son inicialmente grises con tubérculos negros y luego toman una coloración azul grisácea con numerosas líneas transversales. Hay una línea amarilla a cada uno de los lados del cuerpo del gusano. La larva madura mide alrededor de 0.6-0.7 pulgadas de largo. La pupa es color amarillo- café, encerrada en un capullo pequeño cubierto con arena.

Figura 19.

Adulto del gusano de franjas cruzadas. Crédito: Lyle J. Buss, Universidad de la Florida.

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Figura 20.

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Gusano de franjas cruzadas. Crédito: Lyle J. Buss, Universidad de la Florida.

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Biología

Un especialista en crucíferas, el gusano de franjas cruzadas es más un problema en brócoli, coliflor, col y col de Bruselas que lo que es en col rizada y repollo. El tiempo de desarrollo de huevos a adultos varía de 61 días a 68°F a 18 días a 95°F. Pueden ser abundantes durante el invierno y primavera en la Florida. Las larvas empupan en el suelo, cerca de la superficie.

Daño

Las larvas se alimentan de hojas, creando pequeños hoyos. Ellas prefieren los brotes terminales y también pueden enterrarse en el centro de las cabezas del repollo en desarrollo. Mariposa Blanca Mayor del Sur

Ascia monuste (Linnaeus) (Lepidoptera: Pieridae)

Descripción

La mariposa adulta (Figura 21) tiene una envergadura que varía entre 2.5 a 3.4 pulgadas. Las alas de los machos son blancas con un margen negro en forma de zigzag en las alas anteriores. Las hembras varían de color blanco a gris, y, como los machos, ellas tienen un margen negro y una pequeña mancha negra en las alas anteriores. Ambos sexos tienen la punta de las antenas de color turquesa. Huevos amarillos, en forma de huso, son ovipositados singularmente o en grupos sobre las hojas de su hospedero. Las larvas (Figura 22) tienen líneas longitudinales amarillas y grises y múltiples pequeños puntos negros a lo largo del cuerpo. Este insecto pasa a través de cinco estadios. Las pupas son blancas o amarillas con marcas negras y se encuentran tejidas a las plantas.

Figura 21. Adulto de la mariposa blanca mayor del sur. Crédito: Lyle J. Buss, Universidad de la Florida.

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Figura 22.

Larvas de la mariposa blanca mayor del sur. Crédito: Lyle J. Buss, Universidad de la Florida.

Biología

La mariposa blanca mayor del sur es una especie subtropical y tropical, y es principalmente encontrada en áreas costeras. Está presente durante todo el año en el sur de Texas, la península de la Florida, y a lo largo de la costa del golfo. El tiempo de desarrollo desde huevo a la emergencia del adulto es aproximadamente 32 días a 77ºF. Es considerado una plaga importante de plantas de col, incluyendo especies silvestres como el repollo de la playa (Cakile maritima Scopoli) y plantas cultivadas como la col rizada, coliflor, brócoli, arúgula, repollo y mostaza.

Daño

Las larvas mordisquean las hojas de la planta. Ellas usualmente se alimentan en grupos. Las larvas recién emergidas consumen las cáscaras de los huevos y los huevos que aún no han emergido. Los adultos se alimentan del néctar de muchas flores.

Escarabajo del Margen Amarillo

Microtheca ochroloma (Stål) (Coleoptera: Chrysomelidae)

Descripción

Los escarabajos adultos son de color bronce oscuro o negro (Figura 23) y miden alrededor de 0.2 pulgadas de largo. Los bordes de las alas son de color amarillo. Los huevos son de color anaranjado brillante y elongados y son depositados singularmente o en pequeños grupos en áreas protegidas sobre la planta o en la hojarasca sobre el suelo. Las larvas de cuerpo blando son de color amarillo-café y están cubiertas con una fina capa de pelos (Figura 24). La cabeza es café oscura o negra. La larva madura empupa en una envoltura en forma de red en pliegues del follaje, o en la hojarasca sobre la superficie del suelo.

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Figura 23.

Adultos del escarabajo de margen amarillo. Crédito: Angie Niño, Universidad de la Florida.

[Haga click en la figura para maximizar.]

Figura 24.

Larva del escarabajo del margen amarillo. Crédito: James Castner, Universidad de la Florida.

Biología

El escarabajo del margen amarillo es originario de Sur América y fue primero encontrada en los Estados Unidos en 1947. Se ha establecido en la parte sureste de los Estados Unidos. El ciclo de vida no es muy bien conocido en Florida. El escarabajo es capaz de completar su desarrollo en alrededor de 1 mes y puede estar limitado por la disponibilidad de alimento porque está restringido a las crucíferas. Los adultos están activos durante todo el invierno en la Florida y pueden vivir 2 a 3 meses. Nabo, rábano y mostaza son más preferidos que el repollo y las coles, y las hembras que se desarrollan en los tres primeros cultivos producen una mayor cantidad de huevos (hasta casi 500 huevos por hembra).

Daño

El escarabajo del margen amarillo es un problema particularmente para productores orgánicos. Este insecto es especialmente devastador en crucíferas de especialidad como mizuna, pero también se alimenta de nabo, mostaza u otros cultivos de col. Los cultivos en primavera pueden sufrir más daño que los de otoño. Los adultos y las larvas se alimentan de hojas.

Tablas

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Tabla 1.

La palomilla de dorso de diamante

Opción de Recomendación Manejo Los cultivos deben ser monitoreados semanalmente. Un método consiste en caminar en zigzag o haciendo una figura en forma de ocho a través del cultivo de manera que los cuatro cuartos del cultivo sean revisados y las áreas de interior y borde sean examinadas. Lleve una lupa (de al menos 10X), un cuaderno de notas Monitoreo/ y un lápiz para anotar las observaciones. Bolsas de plástico pequeñas son útiles Umbral para colectar insectos para su identificación. Umbrales de 0.1 y 0.3 larvas por planta son utilizados en cultivos de repollo en el noreste de Florida. Examine de 50 a 100 por cultivo (deténgase 10 o 20 veces y examine 5 plantas en cada ubicación). La palomilla de dorso de diamante desarrolla resistencia a insecticidas fácilmente, particularmente a piretroides. La rotación de insecticidas con diferente modo de acción (MoA por sus siglas en inglés) y evitar el uso de piretroides son importantes para el manejo de la palomilla. Hay al menos tres tipos de avispas parasíticas en Florida que atacan el estado de larva o de pupa de la palomilla. La aplicación al inicio de la temporada del cultivo de productos de Bacillus thuringiensis (Bt) no interfiere con la actividad de estos enemigos naturales y puede ayudar a reducir la severidad de las infestaciones. La palomilla de dorso de diamante ha desarrollado resistencia a productos Bt en algunas regiones; sin embargo, Bt siguen siendo una herramienta útil para controlar larvas jóvenes. Se recomienda que la aplicación de productos de Bt de la cepa aizawai (i.e. Agree WG, XenTari DF) sea alternado con productos de Bt formulados con la cepa kurstaki (i.e. Biobit HP, Crymax WDG, Dipel DF, Javelin WG). Los insecticidas reguladores de crecimiento (IGRs por sus siglas en inglés) que pueden ser usados para manejar la palomilla de dorso de diamante incluyen Notas Rimon (novaluron, MoA Grupo15) e Intrepid (metoxifenocide, MoA Grupo 18). IGRs son de acción lenta, pero son herramientas de manejo útiles. Avaunt (indoxacarb, MoA 22) es otra herramienta de rotación importante para el manejo de gusanos. Coragen es un insecticida sistémico del grupo de las diamidas (MoA Grupo 28) que puede ser aplicado a la planta a través de la irrigación por goteo o al follaje para proveer protección contra la palomilla de dorso de diamante y otros gusanos. El ingrediente activo en Coragen es clorantraniliprol, el cual es uno de los ingredientes en Durivo. Durivo también contiene el neonicotinoide tiametoxam (MoA Grupo 4A), el cual provee protección contra insectos chupadores como mosca blanca y áfidos. Verimark es otro insecticida perteneciente al Grupo 28 que contiene ciantraniliprol que es aplicado en la bandeja durante la siembra o a través de la irrigación. Éste provee protección tanto contra gusanos como para las moscas blancas y minadores. Otros insecticidas foliares del Grupo 28 que pueden ser usados para manejar la palomilla de dorso de diamante incluyen Belt y Exirel.

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El ingrediente activo en Belt es flubendiamida, el cual ha sido retirado por el EPA (Agencia de Protección Ambiental por sus siglas en inglés). Los productores pueden usar el material que tienen a la mano. Flubendiamida también está presente en Vetica, el cual contiene buprofezin para controlar ninfas de mosca blanca. Exirel contiene ciantraniliprol y es también efectivo contra mosca blanca y minadores. El enfoque de tipo “ventanas de aplicación” debe ser usado para aplicar insecticidas del Grupo 28 al repollo y otras crucíferas. Si insecticidas del Grupo 28 son aplicados durante la siembra y durante las primeras cinco semanas de la ventana de tratamiento, éstos no deben ser aplicados durante las segundas cinco semanas de la ventana de tratamiento. Avispas parasitoides ayudan al control de las larvas de la palomilla de dorso de Enemigos diamante, especialmente si Bt es el principal insecticida utilizado. Un parasitoide Naturales de huevos (Trichogramma sp.) y hongos entomopatógenos también ayudan al control. Controles culturales importantes incluyen: evitar lo meses cálidos, y destruir los residuos de los cultivos. Otro método para el manejo de la palomilla de dorso de Control diamante en el repollo, es plantar varias líneas de col en el perímetro del cultivo Cultural para que sirva de cultivo trampa y que sea tratado con insecticidas. Para crucíferas de alto valor, cubiertas flotantes colocadas inmediatamente después del trasplante puede eliminar el daño. Tabla 2.

Gusano falso medidor

Opción de Recomendación Manejo Monitoreo/ El cultivo debe ser monitoreado semanalmente. Vea la información sobre la Umbral palomilla de dorso de diamante, arriba. Utilizando Bt como el insecticida principal y usando algunos pesticidas Notas específicos (Entrust (spinosad MoA Grupo 5), Avaunt, Intrepid, Coragen) cuando sea necesario, ayudará a preservar enemigos naturales. Enemigos Avispas y moscas parasitoides, y depredadores generalistas ayudan al control del Naturales gusano falso medidor. Virus de la poliedrosis nuclear también mata este gusano. Evitar cultivar en los meses más cálidos cuando la plaga es más abundante, destruir los residuos del cultivo y controlar malezas. Plantar una fuente de néctar Controles para insectos benéficos puede ser beneficioso—aliso marítimo ha sido evaluado Culturales en repollo. Para crucíferas de alto valor y crucíferas de especialidad, cubiertas flotantes colocadas inmediatamente después del trasplante pueden eliminar el daño. Tabla 3.

Áfidos

149

Opción de Recomendación Manejo El cultivo debe ser monitoreado semanalmente. Vea la información sobre la Monitoreo/ palomilla de dorso de diamante, arriba. No hay información disponible acerca de Umbral umbrales de daño. Utilizando Bt como el insecticida principal para controlar gusanos y usando algunos pesticidas más específicos (Entrust, Avaunt, Intrepid, Coragen, y otros) cuando sea necesario, ayudará a preservar enemigos naturales de áfidos. Notas Insecticidas selectivos para el control de áfidos incluyen Beleaf (flonicamid MoA Grupo 9C), Fulfill (pimetrozina MoA Grupo 9B) y Movento (spirotetramat MoA Grupo 23). Productos del Grupo 4A (neonicotenoides) sirven para el control de áfidos. Avispas parasitoides y depredadores generalistas, como las mariquitas, larvas de Enemigos crisopas y larvas de sírfidos, puede controlar completamente los áfidos si se evita Naturales el uso de pesticidas de amplio espectro. Destruir residuos del cultivo y controlar malezas crucíferas. Plantar una fuente de néctar para insectos benéficos puede ser beneficioso—aliso marítimo ha sido evaluado en repollo. Para crucíferas de alto valor y crucíferas de especialidad, Controles cubiertas flotantes colocadas inmediatamente después del trasplante pueden Culturales eliminar el daño. Sin embargo, si áfidos quedan atrapados por debajo de la cubierta, ellos podrán reproducirse con libertad en la ausencia de enemigos naturales. Tabla 4.

Gusano soldado

Opción de Recomendación Manejo El cultivo debe ser monitoreado al menos semanalmente en búsqueda de daños causados por los gusanos al alimentarse de hojas. Trampas de feromonas pueden ser usadas para monitorear la presencia de polillas. Las plantas jóvenes son más susceptibles al daño. Buscar las masas de huevos sobre las hojas. Buscar en la Monitoreo/ base de las hojas por daño y bajo las hojas más externas y cercanas a la superficie Umbral del suelo, por larvas que pudieran esconderse durante el día lejos del sitio de alimentación. Tratar si encuentra 0.3 larvas por planta, usando la metodología de monitoreo descrito para la palomilla de dorso de diamante. El mejor momento para tratar esta plaga es temprano en la mañana o temprano en la noche. Los insecticidas son más efectivos contra las larvas jóvenes, altas concentraciones y aplicaciones más frecuentes son necesarias para controlar los últimos estadios. Hay insecticidas disponibles para aplicaciones foliares. Notas Cobertura y penetración son importantes en el tratamiento de todas las especies de polillas. Use surfactantes para incrementar la cobertura de la superficie e incrementar el control insecticida.

150

Parasitoides y depredadores de insectos, al igual que patógenos ayudan en el Enemigos control, pero generalmente no ejercen suficiente presión para prevenir la pérdida Naturales de rendimiento. Labrar el campo con discos y la destrucción de residuos del cultivo son importantes para controlar las plagas de gusanos y áfidos y reducir su migración Controles a cultivos cercanos. El gusano soldado se desarrolla bien sobre varias malezas Culturales del grupo amaranto, por lo tanto el control de malezas en los canales y alrededor del cultivo pueden ayudar a reducir las poblaciones antes que invadan el cultivo. Tabla 5.

Gusano perforador de las coles

Opción de Recomendación Manejo Monitoreo/ El cultivo debe ser monitoreado semanalmente. Vea recomendaciones para la Umbral palomilla de dorso de diamante. Utilizando Bt como insecticida principal y usando algunos pesticidas más específicos (Entrust, Avaunt, Intrepid y otros) cuando sea necesario, proveerán control. Las larvas jóvenes que están menos protegidas por las telarañas y hojas Notas dobladas, deben ser el blanco de control. Monitoreo temprano y frecuente es crítico. El tratamiento debe iniciarse tan pronto las larvas son detectadas por su hábito de alimentarse de los brotes terminales, lo cual interfiere con la formación adecuada de la cabeza. Enemigos No se conocen enemigos naturales importantes de esta plaga. Naturales Evitar los meses cálidos en donde la plaga es más abundante, destruir los residuos del cultivo y controlar malezas. Plantar una fuente de néctar para insectos Controles benéficos puede ser beneficioso—aliso marítimo ha sido evaluado en repollo. Culturales Para crucíferas de alto valor y crucíferas de especialidad, cubiertas flotantes colocadas inmediatamente después del trasplante pueden eliminar el daño. Tabla 6.

Gusano trozador

Opción de Recomendación Manejo Plántulas deben ser muestreadas con una frecuencia de dos veces por semana para detectar el gusano trozador o su daño, particularmente en áreas propensas Monitoreo/ para esta plaga. Larvas jóvenes pueden encontrarse agrupadas en el follaje, pero Umbral larvas viejas son usualmente encontradas en el suelo o, en el día, debajo de hojas caídas. Busque follaje marchito o plantas con tallos rotos. Los adultos pueden ser monitoreados usando trampas de feromonas o de luz negra.

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Hay insecticidas disponibles para aplicación a la planta, pre- y post- emergencia Notas y aplicaciones en bandas. Aplicaciones post emergencia son las más eficientes. Enemigos naturales como avispas, moscas parasíticas y escarabajos depredadores en el suelo pueden ejercer una presión de control bastante alta, cercana al 80%. Enemigos Sin embrago, plántulas que emergen en cultivos sin una población de enemigos Naturales naturales residentes pueden experimentar significantes pérdidas debido a la primera generación de gusanos trozadores. Las larvas también pueden ser atacadas por hongos entomopatógenos y virus. Áreas con malezas que luego son usadas en rotación con vegetales de hojas tienen un mayor potencial de pérdidas debido a que larvas viejas sobrevivientes Controles puede cortar las plantas que están emergiendo. Por lo tanto, la preparación el área Culturales para producción se debe hacer lo más rápido posible para dar suficiente tiempo a las larvas sobrevivientes para completar su desarrollo antes de plantar. Tabla 7.

Gusano de franjas cruzadas

Opción de Recomendación Manejo Monitoreo/ El cultivo debe ser monitoreado semanalmente. Vea las recomendaciones para la Umbral palomilla de dorso de diamante. Utilizando Bt como el insecticida principal y usando algunos pesticidas más Notas específicos (Entrust, Avaunt, Intrepid y otros) cuando sea necesario, ayudará a preservar enemigos naturales. Enemigos Avispas parasitoides y depredadores generalistas ayudan al control del gusano de Naturales franjas cruzadas. Evitar cultivar en los meses más cálidos cuando la plaga es más abundante, destruir los residuos del cultivo y controlar malezas. Plantar una fuente de néctar Controles para insectos benéficos puede ser beneficioso—aliso marítimo ha sido evaluado Culturales en repollo. Para crucíferas de alto valor y crucíferas de especialidad, cubiertas flotantes colocadas inmediatamente después del trasplante pueden eliminar el daño. Tabla 8.

Chinches hediondas.

Opción de Recomendación Manejo Examinación visual de las plantas y el uso de trampas amarillas en forma de Monitoreo/ pirámide aplicadas con feromona de agregación para chinches hediondas pueden Umbral ser utilizados para monitorear y capturar chinches.

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El uso de insecticidas sintéticos, primariamente piretroides, pero también algunos nicotenoides (Grupo 4A), es el método de control más efectivo. Algunos Notas pesticidas (Pyganic (piretrinas MoA Grupo 3A) y azadiractina) pueden ser usados en producción orgánica. La tolerancia de las chinches a la mayoría de insecticidas dificulta su supresión. Enemigos Huevos y ninfas sufren una alta mortalidad causada por patógenos, avispas Naturales parasíticas y depredadores como mariquitas, crisopas y chinches depredadores. Triticosecale, trigo sarracero, sorgo, mijo, y girasol pueden ser usados como cultivos trampa plantándolos en la periferia del cultivo principal para interceptar Controles chinches antes de que entren al cultivo comercial. Cultivos trampa deben ser Culturales tratados con insecticidas una vez los chinches son detectados, o cuando los cultivos trampa dejan de ser atractivos para detenerlos y evitar que aumente la población y se muevan al cultivo principal.

Tabla 9.

Escarabajo del margen amarillo.

Opción de Recomendación Manejo Examinación visual de plantas en búsqueda de larvas o adultos o el daño causado Monitoreo/ por éstos. Huevos y pupas son frecuentemente encontrados bajo hojas caídas o en Umbral la base de la planta. El uso de Pyganic y Entrust, los cuales están permitidos en agricultura orgánica, Notas es el método de control más efectivo. Enemigos Depredadores generalistas como mariquitas, crisopas y chinches depredadores Naturales ayudan a controlar huevos, larvas y adultos del escarabajo del margen amarillo. Nabo puede ser usado como cultivo trampa cuando se plantan cultivos de crucíferas menos atractivas para el escarabajo como repollo y col. Los nabos Controles deben ser plantados en la periferia del cultivo dos semanas antes de sembrar el Culturales cultivo principal. El cultivo trampa debe ser tratado con insecticidas una vez adultos o larvas son detectadas para detener que aumente la población y se muevan al cultivo principal.

Notas

Este documento es ENY-464, uno de una serie del Departamento de Entomología y Nematología, Servicio de Extensión Cooperativa de la Florida, Instituto de Alimentos y Ciencias Agrícolas, Universidad de la Florida. Publicado Julio 2002. Revisado Marzo 2010, Junio 2013 y Noviembre 2016. Para más publicaciones relacionadas con horticultura/agricultura, por favor visite la página web de EDIS en http://edis.ifas.ufl.edu.

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S. E. Webb, profesora asociada, Departamento de Entomología y Nematología, Servicio de Extensión Cooperativa de la Florida, Instituto de Alimentos y Ciencias Agrícolas, Universidad de la Florida, Gainesville, FL 32611-0640, A. Niño, candidata a doctorado, Centro de Investigación y Educación del Indian River, Fort Pierce, FL 34945, y H. A. Smith, profesor asistente, Centro de Investigación y Educación en la Costa del Golfo, Universidad de la Florida, Balm, FL 33598.

El uso de nombres comerciales en esta publicación es únicamente con el propósito de proveer información específica. UF/IFAS no garantiza los productos nombrados, y las referencias de ellos en esta publicación no indican nuestra aprobación para la exclusión de otros productos de composición adecuada. Todos los químicos deben ser usados en conformidad con las direcciones de la etiqueta del fabricante. Use pesticidas de manera segura. Lea y siga las direcciones de la etiqueta del fabricante.

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BIOGRAPHICAL SKETCH

Angie Niño started her research career focusing on biological control of mites.

She graduated with a bachelor’s degree in applied biology from the Universidad Militar

Nueva Granada in Colombia. Angie gained professional experience working for 2 years as a research assistant with the Biological Control Research group and the Biodiversity and Ecology of Wild Bees Research group at the Universidad Militar Nueva Granada.

As a Fulbright-Colciencias scholarship recipient, Angie obtained a Master of

Science degree in Entomology at the University of Florida in 2013. She continued her graduate studies at the same university, beginning her Doctor of Philosophy degree program in Entomology in fall 2013. Angie received her Ph.D. from the University of

Florida in the fall of 2016.

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

Delta Honor Society of Agriculture, and the Delta Epsilon Iota Academic Honor Society.

She received the Francisco Luis Gallego award given by the Sociedad Colombiana de

Entomología as the best undergraduate study in 2008. Additionally, she received 2 awards given by the Ft. Pierce Garden Club and the UF-Saint Lucie County Extension

Office.

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