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Initial Steps for Developing a Resistance Management Program for the Southern Chinch Bug, Blissus insularis Barber

Permanent Link: http://ufdc.ufl.edu/UFE0023774/00001

Material Information

Title: Initial Steps for Developing a Resistance Management Program for the Southern Chinch Bug, Blissus insularis Barber
Physical Description: 1 online resource (171 p.)
Language: english
Creator: Vazquez, Julie
Publisher: University of Florida
Place of Publication: Gainesville, Fla.
Publication Date: 2009

Subjects

Subjects / Keywords: augustinegrass, blissus, insecticide, insularis, resistance, st
Entomology and Nematology -- Dissertations, Academic -- UF
Genre: Entomology and Nematology thesis, Ph.D.
bibliography   ( marcgt )
theses   ( marcgt )
government publication (state, provincial, terriorial, dependent)   ( marcgt )
born-digital   ( sobekcm )
Electronic Thesis or Dissertation

Notes

Abstract: Blissus insularis Barber, is a serious pest of St. Augustinegrass and has a history of resistance to insecticides in Florida. A resistance management program is needed for this pest but initial steps are required. The goals of this study were to 1) sample select B. insularis populations in Florida to describe their susceptibility to bifenthrin, document new locations of bifenthrin resistance, and evaluate another pyrethroid, permethrin, 2) develop a synchronous rearing method for B. insularis, and 3) develop an improved bioassay that could be used for detecting insecticide susceptibility differences between male and female B. insularis, evaluate and validate both the sprig-dip and the new bioassay under standardized conditions, and determine optimal exposure times and sample sizes to be used for each bioassay for selected insecticides. The results of objective 1 suggest bifenthrin resistance continues to be problematic, is becoming more widespread, and there is a positive relationship between insecticide application and the development of bifenthrin resistance. This study documents the first case of insecticide resistance in the Florida Panhandle and first report of B. insularis resistance to permethrin. Five different rearing methods were attempted for B. insularis. The use of glass jars and a combined diet of fresh corn cob and St. Augustinegrass proved to be the best synchronous rearing method for producing B. insularis of known age and generation. No reduction in body size was observed after nine generations of rearing. In addition, the high number of brachypterus B. insularis produced indicates that populations were not stressed. An airbrush bioassay for testing contact and systemic insecticides was developed, and evaluations were made of both the airbrush and sprig-dip bioassays under standardized conditions to determine sample size and duration of tests. The sprig-dip bioassay was more sensitive in detecting lower LC values than the airbrush bioassay when testing B. insularis against bifenthrin. The airbrush and sprig-dip bioassays will be useful tools for detecting and monitoring of insecticide resistance in B. insularis. The airbrush bioassay would be beneficial for use in studies concerning cross resistance, mechanisms, mode-of-inheritance, and stability of pyrethroid resistance because of the ability to easily detect differences between male and female B. insularis and reduced variability.
General Note: In the series University of Florida Digital Collections.
General Note: Includes vita.
Bibliography: Includes bibliographical references.
Source of Description: Description based on online resource; title from PDF title page.
Source of Description: This bibliographic record is available under the Creative Commons CC0 public domain dedication. The University of Florida Libraries, as creator of this bibliographic record, has waived all rights to it worldwide under copyright law, including all related and neighboring rights, to the extent allowed by law.
Statement of Responsibility: by Julie Vazquez.
Thesis: Thesis (Ph.D.)--University of Florida, 2009.
Local: Adviser: Buss, Eileen A.
Electronic Access: RESTRICTED TO UF STUDENTS, STAFF, FACULTY, AND ON-CAMPUS USE UNTIL 2011-05-31

Record Information

Source Institution: UFRGP
Rights Management: Applicable rights reserved.
Classification: lcc - LD1780 2009
System ID: UFE0023774:00001

Permanent Link: http://ufdc.ufl.edu/UFE0023774/00001

Material Information

Title: Initial Steps for Developing a Resistance Management Program for the Southern Chinch Bug, Blissus insularis Barber
Physical Description: 1 online resource (171 p.)
Language: english
Creator: Vazquez, Julie
Publisher: University of Florida
Place of Publication: Gainesville, Fla.
Publication Date: 2009

Subjects

Subjects / Keywords: augustinegrass, blissus, insecticide, insularis, resistance, st
Entomology and Nematology -- Dissertations, Academic -- UF
Genre: Entomology and Nematology thesis, Ph.D.
bibliography   ( marcgt )
theses   ( marcgt )
government publication (state, provincial, terriorial, dependent)   ( marcgt )
born-digital   ( sobekcm )
Electronic Thesis or Dissertation

Notes

Abstract: Blissus insularis Barber, is a serious pest of St. Augustinegrass and has a history of resistance to insecticides in Florida. A resistance management program is needed for this pest but initial steps are required. The goals of this study were to 1) sample select B. insularis populations in Florida to describe their susceptibility to bifenthrin, document new locations of bifenthrin resistance, and evaluate another pyrethroid, permethrin, 2) develop a synchronous rearing method for B. insularis, and 3) develop an improved bioassay that could be used for detecting insecticide susceptibility differences between male and female B. insularis, evaluate and validate both the sprig-dip and the new bioassay under standardized conditions, and determine optimal exposure times and sample sizes to be used for each bioassay for selected insecticides. The results of objective 1 suggest bifenthrin resistance continues to be problematic, is becoming more widespread, and there is a positive relationship between insecticide application and the development of bifenthrin resistance. This study documents the first case of insecticide resistance in the Florida Panhandle and first report of B. insularis resistance to permethrin. Five different rearing methods were attempted for B. insularis. The use of glass jars and a combined diet of fresh corn cob and St. Augustinegrass proved to be the best synchronous rearing method for producing B. insularis of known age and generation. No reduction in body size was observed after nine generations of rearing. In addition, the high number of brachypterus B. insularis produced indicates that populations were not stressed. An airbrush bioassay for testing contact and systemic insecticides was developed, and evaluations were made of both the airbrush and sprig-dip bioassays under standardized conditions to determine sample size and duration of tests. The sprig-dip bioassay was more sensitive in detecting lower LC values than the airbrush bioassay when testing B. insularis against bifenthrin. The airbrush and sprig-dip bioassays will be useful tools for detecting and monitoring of insecticide resistance in B. insularis. The airbrush bioassay would be beneficial for use in studies concerning cross resistance, mechanisms, mode-of-inheritance, and stability of pyrethroid resistance because of the ability to easily detect differences between male and female B. insularis and reduced variability.
General Note: In the series University of Florida Digital Collections.
General Note: Includes vita.
Bibliography: Includes bibliographical references.
Source of Description: Description based on online resource; title from PDF title page.
Source of Description: This bibliographic record is available under the Creative Commons CC0 public domain dedication. The University of Florida Libraries, as creator of this bibliographic record, has waived all rights to it worldwide under copyright law, including all related and neighboring rights, to the extent allowed by law.
Statement of Responsibility: by Julie Vazquez.
Thesis: Thesis (Ph.D.)--University of Florida, 2009.
Local: Adviser: Buss, Eileen A.
Electronic Access: RESTRICTED TO UF STUDENTS, STAFF, FACULTY, AND ON-CAMPUS USE UNTIL 2011-05-31

Record Information

Source Institution: UFRGP
Rights Management: Applicable rights reserved.
Classification: lcc - LD1780 2009
System ID: UFE0023774:00001


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INITIAL STEPS FOR DEVELOPING A RESISTANCE MANAGEME NT PROGRAM FOR THE SOUTHERN CHINCH BUG, BARBER By JULIE CARA VZQUEZ A DISSERTATION PRESENTED TO THE GRADUATE SCHOOL OF THE UNIVERSITY OF FLOR IDA IN PARTIAL FULFILLMENT OF THE REQUIREMENTS FOR THE DEGREE OF DOCTOR OF PHILOSOPHY UNIVERSITY OF FLORIDA 2009 1

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2009 Julie Cara Vzquez 2

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I dedicate this dissertation to my loving husband, Ricardo Jos Vzquez, for his unending love and support. 3

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ACKNOWLEDGMENTS I am grateful to my major advi sor, Dr. Eileen Buss, for her guidance, financial support, and allowing me to switch projects early on in my progr am. Also, this research would not have been possible were it not for my committee members Dr s. Marjorie Hoy, Nate Royalty, Mike Scharf, and Laurie Trenholm. Words cannot express my gratitude for their expertise, guidance, and utmost professionalism throughout this research. I am truly fortunate to have been mentored by these outstanding scientists and what I have learned from them during this time will follow me throughout my professional career I would also like to thank Dr. Grady Miller. Although you were only a part of my committee for the first tw o years, your advice and expertise was greatly appreciated. I am also deeply indebted to Bayer Environmental Science and C. P. and Lynn Steinmetz for providing funding for this research. This research could not have b een completed without the assist ance of several individuals. First and foremost I thank Paul Ruppert for hi s assistance in collecti ng insects and obtaining supplies. Over the years there have been other individuals that have intermittently worked in our lab, but Jade Cash, Amin Cheikhi, Megan Gilbert, and Rachel Sheahan by far assisted the most with insect colony and plant maintenance. Th e time and care these individuals took to assist with my research is greatly appreciated. Speci al thanks go to Brian Owens and Jason Haugh at the G.C. Horn Memorial Turfgrass Field Laboratory for the use of supplies and bench space. I acknowledge Dr. Phillip Kaufman for use of benc h space, Dr. Phil Koehler for assistance with bioassay development, and Dr. Jacqueline L. Robertson for assistance with the PoloPlus program. I acknowledge and thank Lyle Buss for his time and expertise in photography. I am deeply grateful to Dr. B. Unruh, Advance Tech Pest, Bayer Environmental Science, Citrus Pest Management, Corey Services, Environmental Pest & Lawn Services, Inc. Florida Pest Control and Chemical Co., FMC Corporation, Keith White Lawn-N-Pest Inc., Middelton Pest Control 4

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Inc., TruGreen Companies LLC, and Valley Cr est Com panies for assistance with obtaining populations for my research. I thank the faculty and staff of the Entomology and Nematology Department for their advice, guidance, and friendship. They all have my deepest respect fo r the amount of time and endless support that you provide to students. I am especially gr ateful to Debbie Hall for keeping me informed of deadlines and helping me stay on track. Also, during my time at the University of Florida I have made many wonder ful friends that have made my lif e all that more enjoyable. I would especially like to acknowledge Jay Cee Turner, Olga Kostro mytska, and Jessica Platt for their constant encouragement and support. Special thanks are given to my family for all of their love, support, and understanding of all the times when I could not participate in social gatherings. Most of all I would like to thank my husband Ricky for his sacrifices and service to our great country, and for his love and support while obtaining this degree. Your constant enco uragement helped me to stay focused and make it through the hardest of times. 5

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TABLE OF CONTENTS page ACKNOWLEDGMENTS...............................................................................................................4 LIST OF TABLES................................................................................................................. ..........9 LIST OF FIGURES.......................................................................................................................11 ABSTRACT...................................................................................................................................13 CHAPTER 1 LITERATURE REVIEW.......................................................................................................15 Turfgrasses..............................................................................................................................15 Functional Benefits..........................................................................................................15 Recreational and Aesthetic Benefits................................................................................16 Turfgrass Industry in Florida...........................................................................................16 St. Augustinegrass...........................................................................................................16 ......................................................................................................................17 Host Plants and Distribution............................................................................................17 Biology and Life History.................................................................................................18 Feeding Habits and Damage............................................................................................19 Rearing of spp.....................................................................................................21 Management Practices............................................................................................................24 Biological Control...........................................................................................................24 Host Plant Resistance......................................................................................................25 Cultural Control............................................................................................................... 26 Chemical Control.............................................................................................................27 Organophosphates....................................................................................................28 Carbamates...............................................................................................................29 Pyrethroids...............................................................................................................29 Neonicotinoids.........................................................................................................30 Insecticide resistance in ........................................................................31 Insecticide Resistance......................................................................................................... ....32 Detection and Documentation.........................................................................................32 Choice of Bioassay..........................................................................................................33 Source of variability in insecticide bioassays..........................................................35 Intrinsic factors.........................................................................................................35 Extrinsic factors........................................................................................................36 Resistance Mechanisms...................................................................................................37 Biotic, Genetic, and Operational Factors........................................................................39 Resistance Management..................................................................................................40 Resistance Management Models.....................................................................................40 Research Objectives............................................................................................................ ....43 6

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2 SUSCEPTIBILITY OF POPULATIONS IN FLORIDA TO BIFENTHRIN AND PERMETHRIN.............................................................................................................52 Introduction................................................................................................................... ..........52 Materials and Methods...........................................................................................................54 St. Augustinegrass Maintenance.....................................................................................54 2006 Collection Sites.......................................................................................................54 2008 Collection Sites.......................................................................................................55 Insects..............................................................................................................................55 2006 Tests........................................................................................................................55 Bifenthrin.................................................................................................................55 Permethrin................................................................................................................56 2008 Bifenthrin Test........................................................................................................56 Statistical Analysis..........................................................................................................5 6 Results and Discussion......................................................................................................... ..57 2006 Tests........................................................................................................................57 Bifenthrin.................................................................................................................57 Permethrin................................................................................................................61 2008 Tests........................................................................................................................61 3 SYNCHRONOUS METHOD FOR REARING ON CORN AND ST. AUGUSTINEGRASS.............................................................................................................78 Introduction................................................................................................................... ..........78 Materials and Methods...........................................................................................................81 Test 1. Small-Scale Rearing of A dults on Corn and Nymphs on Grass.........................81 St. Augustinegrass maintenance...............................................................................81 Corn preparation.......................................................................................................82 Insect collection........................................................................................................82 Oviposition and nymph container construction.......................................................82 Egg harvest method..................................................................................................83 Test 2. Assessment of Time of Day for Oviposition......................................................83 Test 3. Rearing Nymphs on Planted Gr ass in Builders Sand and Glass Jars................84 Corn preparation.......................................................................................................84 Insect collection and colony maintenance................................................................84 Test 4. Corn Only Rearing Method................................................................................85 Test 5. Improved Method Using Corn and Grass...........................................................86 Colony jar construction............................................................................................86 Egg harvest method and nymph maintenance..........................................................86 Determining quality and success of rearing method 5.............................................87 Results and Discussion......................................................................................................... ..88 Test 1. Small-Scale Rearing of A dults on Corn and Nymphs on Grass.........................88 Test 2. Assessment of Time of Day for Oviposition......................................................88 Test 3. Rearing Nymphs on Planted Gr ass in Builders Sand and Glass Jars................88 Test 4. Corn Only Rearing Method................................................................................89 Test 5. Improved Method Using Corn and Grass ...........................................................90 7

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4 CONCENTR ATION-MORTALITY RESPONSES TO FIVE INSECTICIDES BY A SUSCEPTIBLE COLONY OF USING AN AIRBRUSH BIOASSAY...........107 Introduction................................................................................................................... ........107 Materials and Methods.........................................................................................................108 St. Augustinegrass Maintenance...................................................................................108 Insect Collection and Maintenance...............................................................................109 Insecticides....................................................................................................................109 Spray Application Device..............................................................................................110 Determining Uptake for Systemic InsecticidesUsing an Airbrush Bioassay............110 Comparison of Airbrush and Sprig Dip Bioassays........................................................111 Airbrush Bioassay.........................................................................................................111 Statistical Analysis........................................................................................................111 Results and Discussion......................................................................................................... 113 Determining Uptake for Systemic Insecticides.............................................................113 Comparison of Airbrush and Sprig-Dip Bioassays.......................................................113 Bifenthrin...............................................................................................................113 Imidacloprid...........................................................................................................115 Subsampled comparison data--bifenthrin...............................................................116 Subsampled comparison data--imidacloprid..........................................................117 Airbrush Bioassay.........................................................................................................119 5 CONCLUSIONS.................................................................................................................. 137 LIST OF REFERENCES.............................................................................................................147 BIOGRAPHICAL SKETCH.......................................................................................................171 8

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LIST OF TABLES Table page 2-1 Collection sites and the number of insecticide applications made to the populations in Florida in 2006 that were te sted for susceptibility to bifenthrin................64 2-2 Collection sites of the populations in Florida in 2008 that were tested for susceptibility to bifenthrin.................................................................................................66 2-3 Response of Florida populations collected in 2006 to bifenthrin after 72 h using a sprig-dip bioassay...............................................................................................67 2-4 Hypothesis tests comparing the slopes and intercepts of logit regression lines for 15 populations in comparison to the most susceptible population, GE18, after exposure to bifenthrin for 72 h using a sprig-dip bioassay........................................68 2-5 Response to permethrin after 72 h of two populations collected in 2006 using a sprig-dip bioassay..................................................................................................69 2-6 Response of Florida populations collected in 2008 to bifenthrin after 24 h using an airbrush bioassay..............................................................................................70 2-7 Hypothesis tests comparing the slopes and intercepts of logit regression lines for 6 populations in comparison to a sus ceptible laboratory colony, LO, after exposure to bifenthrin for 72 h using an airbrush bioassay...............................................71 3-1 The total number eggs in each replicate at the start of Test 1 and the number of male and female that successfully emerged after 5.5 wk........................................96 3-2 Mean number of eggs collected at each 8-h interval in Test 2.......................97 3-3 The total number eggs in each replicate at the start of Test 3 and the number of adults that su ccessfully emerged after 6 wk.......................................................98 3-4 The total number eggs in each replicate at the start of test 4 and the number and stage of found after 8 wks........................................................................................99 3-5 The number of emerged generation nine adults, percentage survival, wing type, and comparison of mean body lengt h of brachypterus females by replicate for test 5..................................................................................................................... ......100 4-1 Insecticides tested agai nst a susceptible colony of .......................................123 4-2 Concentration-mortality data at different exposure times for a susceptible laboratory colony exposed to St. Augustinegra ss treated with clot hianidin 1, 3, and 7 d before bioassay..............................................................................................................124 9

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4-3 Comp arison of concentration-mortality data for a susceptible laboratory colony to bifenthrin and imidacloprid at 24, 48, and 72 h using the airbrush and sprig-dip bioassays...........................................................................................................125 4-4 Comparison of subsampled concentra tion-mortality data for a susceptible laboratory colony exposed to bifenthr in using the airbrush and sprig-dip bioassays...................................................................................................................... ....126 4-5 Comparison of subsampled comparison test concentration-mortality data for a susceptible laboratory colony exposed to im idacloprid using the airbrush and sprig-dip bioassays....................................................................................................127 4-6 The mean number of male and female that located treated plant material within 1 h of introduction into the airbrush bioassay......................................................128 4-7 Concentration-mortality data compared for males and females from a susceptible laboratory colony treated with five insecticides after 24, 48, and 72 h using the airbrush bioassay........................................................................................................12 9 4-8 Analysis of LC50 values for 24, 48, and 72 h within each sex to determine bioassay time for the contact insecticides bifenthrin, carbaryl, and trichlorfon........................................................................................................................131 4-9 Analysis of LC90 values for 24, 48, and 72 h within each sex to determine bioassay time for the systemic ins ecticides clothianidin and imidacloprid....132 10

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LIST OF FIGURES Figure page 1-1 Severe damage from feeding that stops at the neighboring bahiagrass lawn................................................................................................................................44 1-2 Brachypterus and macropter us male and female ............................................45 1-3 A healthy egg in early and late development.................................................46 1-4 nymphs...................................................................................................47 1-5 Lawns damaged by .........................................................................................48 1-6 St. Augustinegrass lawns with populations encroaching on neighboring lawns..................................................................................................................................49 1-7 St. Augustinegrass with excessive thatch..........................................................................50 1-8 A egg parasitized by and image of an adult ............51 2-1 LC50 Lethal concentration ratios with lo wer 95% confidence limits indicating the fold difference for each population of in comparison to the most susceptible population (GE18) when tested with bifenthrin..............................................72 2-2 LC90 Lethal concentration ratios with lo wer 95% confidence limits indicating the fold difference for each population of in comparison to the most susceptible population (GE18) when tested with bifenthrin..............................................73 2-3 Map showing the distribution of insecticide-resistant populations in Florida between 2003-2008...............................................................................................74 2-4 Linear regression showing the relati onship between the number of insecticide applications made in 2006 to populations and respective lethal concentration ratios (at LC50).............................................................................................75 2-5 LC50 Lethal concentration ratios with lo wer 95% confidence limits indicating the fold difference for each population of in comparison to the most susceptible population (LO) when tested with bifenthrin..................................................76 2-6 LC90 Lethal concentration ratios with lo wer 95% confidence limits indicating the fold difference for each population of in comparison to the most susceptible population (LO) when tested with bifenthrin..................................................77 3-1 Experimental design of Tests 1 and 2 showing the oviposition container used to maintain adults and collect eggs, and the container used for nymph development.................................................................................................................... .101 11

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3-2 7.6-L oviposition jar u sed for maintaining adults and collecting eggs, and an image of the egg roll used in Tests 3, 4, and 5 displaying ..............102 3-3 7.6-L glass jar with grass planted in st erilized builders sand for nymph development used in Test 3................................................................................................................. ..103 3-4 7.6-L glass jar showing wax paper and car dboard assemblage at the bottom and a completely constructed jar with dental castone used in Test 5........................................104 3-5 7.6-L glass jar containing St. A ugustinegrass for development of nymphs used in Test 5................................................................................................................. ..105 3-6 Flow chart of steps and approximate time and labor required to rear one jar of in a synchronous laboratory system (Test 5)....................................................106 4-1 The sprig-dip bioassay conventionally used for testing insecticides against ............................................................................................................................133 4-2 The Paasche airbrush and BioServe bioa ssay tray and lid used in the airbrush bioassay............................................................................................................................134 4-3 The differences in variabili ty between replicates of bife nthrin for the airbrush and sprig-dip bioassays after 24 and 48 h...............................................................................135 4-4 The differences in variabili ty between replicates of imid acloprid for the airbrush and sprig-dip bioassays after 24 and 48 h...............................................................................136 12

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Abstract of Dissertation Pres ented to the Graduate School of the University of Flor ida in Partial Fulfillment of the Requirements for the Degree of Doctor of Philosophy INITIAL STEPS FOR DEVELOPING A RESISTANCE MANAGEMENT PROGRAM FOR THE SOUTHERN CHINCH BUG, BARBER by Julie Cara Vzquez May 2009 Chair: Eileen A. Buss Major: Entomology and Nematology Barber, is a serious pest of St. Augustinegrass and has a history of resistance to insecticides in Florida. A resist ance management program is needed for this pest but initial steps are required. The goals of this study were to 1) sample select populations in Florida to describe their suscepti bility to bifenthrin, document new locations of bifenthrin resistance, and eval uate another pyrethroid, permet hrin, 2) develop a synchronous rearing method for and 3) develop an improved bioassay that could be used for detecting insecticide susceptibility differences between male and female evaluate and validate both the sprig-dip and the new bioassay under standardized conditions, and determine optimal exposure times and sample sizes to be used for each bioassay for selected insecticides. The results of objective 1 suggest bifenthrin resistance continues to be problematic, is becoming more widespread, and there is a positi ve relationship between insecticide application and the development of bifenthrin resistance. This study documents the first case of insecticide resistance in the Florida Pa nhandle and first report of resistance to permethrin. Five different rearing me thods were attempted for The use of glass jars and a combined diet of fresh corn cob and St. A ugustinegrass proved to be the best synchronous 13

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14 rearing method for producing of known age and generation. No reduction in body size was observed after nine genera tions of rearing. In addition, the high number of brachypterus produced indicates that popul ations were not stressed. An airbrush bioassay for testing contact a nd systemic insecticides was developed, and evaluations were made of both the airbrush and sprig-dip bioassa ys under standardized conditions to determine sample size and duration of tests. The sprig-dip bioassay was more sensitive in detecting lower LC values than the airbrush bioassay when testing against bifenthrin. The airbrush and sprig-dip bioassays will be useful tools for detecting and monitoring of insecticide resistance in The airbrush bioassay would be beneficial for use in studies concerning cross resistance, m echanisms, mode-of-inheritance, and stability of pyrethroid resistance because of th e ability to easily detect differences between male and female and reduced variability.

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CHAPTER 1 LITERATURE REVIEW Turfgrass is a vegetative ground cover used in landscapes and is the mo st widely used ornamental crop in the United St ates (Emmons 1995). Humans have used turfgrasses for more than 10 centuries as a means to enhance their environment and quality of life (Beard 1973, Beard and Green 1994). There are several functional, recreational, and aesthet ic contributions of turfgrasses. Turfgrasses are maintained in a long-term stab le state and thus great ly aid in protecting nonrenewable soil resources from water and wi nd erosion (Kageyama 1982, Potter and Braman 1991, Beard and Green 1994). Once a vigorous and dens e turf develops in the landscape, it also plays a significant role in reducing water runoff in urban and suburban areas, especially those near paved surfaces (Kageyama 1982, Potter and Braman 1991, Florida Department of Environmental Protection 2002, Bell and Moss 2008). In addition, the development of a healthy root zone allows greater infilt ration of rain or irrigation by im proving soil structure and reducing soil compaction (Florida Department of Envir onmental Protection 2002). The root zone also aids in facilitating biodegradation of organic pollutants, air contaminants, and pesticides used in lawns, as well as encouraging soil-building processes through the decomposition of organic matter and formation of humus. Healthy turf grass also muffles noise, reduces glare, and modifies temperatures (Kageyama 1982, Potter and Braman 1991, Beard and Green 1994, Florida Department of Environmental Protection 2002). Also, a 15 m 15 m turf area absorbs carbon dioxide, ozone, hydrogen fluoride, and perosyacetyle nitrate and can release enough oxygen to meet the needs of a family of four (Emmons 1995). 15

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Healthy turfgrass prov ides a safe recreational surface with a cushioning effect that reduces injuries to humans compared to walking or running on poorlyor non-tu rfed soils (Beard and Green 1994). Also, the beauty of a well main tained lawn and landscape can have a positive impact on mental health by providi ng green space in urban areas, as well as increase property values by as much as 15% (Kageyama 1982, Potter and Braman 1991, Emmons 1995). Many lawns in Florida are established through so dding. Sod is dense tu rf that is cut in pieces or strips from the soil and sold as ground cover for use in lawns (Emmons 1995, Christians 2004). In a national study, Florida was ranked first in terms of economic impact of sod production (Haydu et al. 2006). In 2003, the tota l sod production in Florida was estimated to be 93,000 ha, with 64% being St. Augustinegrass (Haydu et al. 2005). Only 3% of harvested sod is sold outside of Florida. With so much demand for sod in Florida, there is also a high demand for maintaining it. Florida is second only to California in terms of employment impacts of the turfgrass industry, providing 83,944 j obs in 2002 (Haydu et al. 2006). St. Augustinegrass, (Walt.) Kuntze, is a warm-season, coarsetextured, aggressive, and stolonife rous grass (Turgeon 1996) that is believed to be native to the coastal regions of both the Gulf of Mexico a nd the Mediterranean (Trenholm and Unruh 2005). Carter and Duble (1976) estimated that St. Augustinegrass comprised as much as 96% of lawns in the Gulf Coast area. In Florida, the firs t known record of planting St. Augustinegrass was from a diary by A. M. Reed, where he wr ote on November 11, 1880, George planting St. Augustine grass in avenue in afternoon. It was planted as a turf alongside an avenue at A. M. Reeds Mulberry Grove plantation, at Yukon, near Orange Park, FL (Works Progress 16

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Administration 1939, White and Busey 1987, Busey 1995) Today, it is the prim ary turfgrass in residential lawns and comprises 70% or 1.2 million ha in Fl orida (Hodges et al. 1994, Busey 2003). Most St. Augustinegrass cultivars have good salt (Dudeck et al. 1993) and shade tolerance (White and Busey 1987) and are usually established by plugs or sod (Christians and Engelke 1994, Christians 2004). St. Augustinegrass also grows well in most soils and climatic regions in Florida (Trenholm and Unruh 2005). Its aggressive growth habit gives it good recuperative capability, but it is pron e to thatch buildup (Potter 1998). Blissus insularis The southern chinch bug, Barber (Barber 1918), is considered the most damaging insect pest of St. Augustinegrass (Reinert and Portier 1983, Busey and Coy 1988, Crocker 1993). was at first believed to be a variety of and a member of the complex (Leonard 1966). However, Leonard showed was genetically isolated fr om the other taxa of the complex and gave it species rank. Originally known as the lawn chinch bug, the southern chinch bug was given its current name when it was designated as a distinct sp ecies (Stringfellow 1969, Sw eet 2000). It was first documented as a pest of St. Augustineg rass in 1922 (Newell and Berger 1922). also attacks other lawn gr asses including bahiagrass (Fluegg), bermudagrass [ (L.) Pers.], centipedegrass [ (Munto)], and zoysiagrass ( spp.), but most of the injury to these has occurred near heavily infested St. Augustinegrass (Kerr 1966). has also been found in lawns that contained a mix of St. Augustinegrass and centip edegrass where the St. Augustinegrass was killed and the centipedegrass was left unharmed (Kerr 1966) Buss (E.A.B., unpublished data) observed feeding and damage to a St. Augustin egrass lawn stopped ab ruptly where the 17

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neighboring bahiagrass lawn st arted (Figure 1-1). Other hosts include crabgrass ( Hitchc.), torpedograss ( L.), and Pangolagrass ( Stent) (Slater and Baran owski 1990, Brandenburg and Villani 1995). occurs in the southern U. S. coastal states, Hawaii, a nd Mexico (Henry and Froeschner 1988, Vittum et al. 1999, Sweet 2000). Adult are small insects with the adult body measuring between 2-4 mm long (Cherry and Wilson 2003) and 1 mm wide (Leona rd 1968). Females are usually larger than males (Figure 1-2 A and B). The sclerites at the ventral tip of the abdomen are rounded in males and triangular in females (Figure 1-2 C and D). Wings are white with a distinctive triangularshaped black marking in the middle of the outer edge of each wing and are folded flat over the back causing the tips to overlap. Populations may consist mostly of short-winged forms (brachypterous), long-winged forms (macropterous ), or both [Figure 1-2 A and B] (Wilson 1929, Komblas 1962, Leonard 1966, Reinert a nd Kerr 1973). In Florida, macroptery is greatest during the summer and fall although reasons for this are unknown (Cherry 2001a). However, studies have shown that macroptery in the oriental chinch bug, Okajima is density dependent, and is strongly enhanced by seasonal factors (long day length, high temperature) (Fujisaki 2000). The biology of is well documented. When courting, males and females approach each other, make first contact with th eir antennae, then pair facing opposite directions (Vittum et al. 1999). Copulation may last as long as 2 h and during this time female are more active than males and may walk about and/or feed (Leonard 1966, Vittum et al. 1999). Eggs are laid singly or a few at a time in sheaths, near the grass nodes, in soft soil, or in other protected areas (Beyer 1924a, Kuitert and Nutter 1952, Reinert and Kerr 1973). The eggs are 18

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white when first laid (Eden and Self 1960), turn ing beige (Figure 1-3 A) then bright orange (Figure 1-3 B) just before hatching. Young nymphs ar e as sm all as 1.0 mm, are reddish-orange with a white band across the dorsal side of the abdomen, and become black in color as they mature (Figure 1-4 A-E). Many nymphs crawl betw een the folds of the sheath located at the lower portion of the grass leaf (Christians 2004 ), and may remain hidden for up to 10 d (Kerr 1966). Development from egg to adult depends on location and temperature. In Florida, Kerr (1966) reported can complete development from egg to adult in 34.7 d at 28.3C and in 93.4 d at 21.1C. All life stages are present throughout th e year in most of the state with three to four generations occurring in northern Florida and seven to ten in southern Florida each year (Kerr 1966, Reinert and Kerr 1973). Although capable of flight, adult move between lawns mainly by walking and many have been observed crawling across pa ved areas bordering heavily infested lawns (Kerr 1966). All life stages are distributed vertic ally through the turf th atch and into the upper organic layer of the soil, with densities of up to 2,000 /0.1 m2 being reported (Reinert and Kerr 1973). Light to moderate infestations are aggregated in small areas in the lawn, but can occur throughout the entir e lawn in heavily infest ed areas (Cherry 2001b). spp. are sap feeders (Slater 1976) and feed on the phloem and xylem in meristematic regions of the grass (Painter 1928) causing wilting, chlorosis, stunting, and eventually death (Painter 1928, Negron and Riley 1990, Spike et al. 1991). As the grass dies, the insect s continue to move outward to feed on more-succulent grass, en larging the damaged area (Figure 1-5), and may easily encroach onto neighboring St. Augustinegrass lawns (Figure 1-6). St. Augustinegrass cultivated on high, dry, sandy, or she ll soil is especia lly vulnerable to damage (Wilson 1929, Woods 2007). 19

PAGE 20

prefer open sunny areas of St. Augustinegrass, especially areas with abundant thatch (Reinert and Kerr 1973). Thatch is the layer of accumu lated decomposing leaf blades, stems, and roots on top of the soil su rface (Figure. 1-7) (Emmons 1995, Trenholm and Unruh 2005). Where temperatures are warmer, partic ularly in South Florid a, the grass may grow continuously and create a thick, spongy thatch (V ittum et al. 1999). Thatch that is 10 15 cm thick is common and can be up to 30 cm deep (Vittum et al. 1999), providing with shelter and possibly protecting them from predat ion and environmental stress (Reinert and Kerr 1973). The abundance of Montandon was also closely linked to thatch thickness in lawns (Davis and Smitley 1990). The effect of moisture on populations and their feeding injury to turf is equivocal. may thrive when the grass is most tender and succulent, and its feeding may prevent normal growth and cause a dwarfed condition to the grass (Beyer 1924a, Vzquez and Buss 2006). Warm and fairly dry w eather is most favorable for hatching of eggs (Beyer 1924a). injury may be more evident during dry weather because dryness reduces turf vigor and favors the rapid increase in populations (Wilson 1929). Kerr (1966) suggested that moisture had a marked but paradoxical effect on populations. Heavy irrigati on or rainfall may make the gr ass more succulent and able to tolerate some feeding damage, while at the same time making the grass more attractive to However, destructive outbreaks of are sometimes prevented by heavy rainfall (Beyer 1924a) by killing the y oung nymphs, and this is true for other spp. as well (Webster 1907). Long-term feeding damage may look lik e drought stress, but not be a result thereof. Also, could already be present a nd feeding in a lawn, but a secondary stress, like drought, may intensify the damage (Vzquez and Buss 2006). 20

PAGE 21

Several authors have attempted to rear spp. under laboratory and greenhouse conditions to better understand its biology, life history, and f eeding habits. The following provides a brief review of previously re ported rearing procedures for Blissidae. Blissus Yamada et al. (1984) reared the oriental chinch bug, Okajima, on maize, Kentucky bluegrass, sorghum, and sugarcan e. Sugarcane leaves were the best diet on which to rear more than two generations of However, Yamada et al. (1984) reported that only 40% of the second generation successfully survived to the adult stage. Dahms (1947) and Todd (1966) reared the common chinch bug, (Say), on plants maintained in a speciall y prepared nutrient solution. However, the insects were only maintained on a limited basis. Later, Parker and Randolph (1972) reared in the laboratory on alternating stacked laye rs of maize and sorghum stalk sections. Each stalk section end was dipped in melted paraffin wax and allowed to dry before placement in heat-sterilized 3.78-L cardboard cartons. Cartons were maintained in growth chambers at 32 2C with a 14L:10D photoperiod. Pathogens were controlled by washing the stalk sections with warm soapy water and rinsing in a 1.0% solution of benzalkonium chloride before placement in cardboard cartons. The carton tops were covered with heat-ste rilized Purelin singlefold no. 515 towels. eggs, nymphs, and adults were easily removed from the top stalks and used to start ne w colonies. Each 3.78-L cardboa rd carton could produce 800-1000 chinch bugs (Parker and Randolph 1972). Wilde et al. (1987) also reared but used small grains, maize, sorghum, and millet. Ten to fifteen maize, sorghum, or m illet plants were germinated in 15-cm pots. Two to 3 wk after planting, 25 unsexed adults were pl aced in each pot and c onfined with 15 45 cm plastic cages with ventilation hol es on the side. Sand was used at the base with Teflon 21

PAGE 22

(DuPont, Wilmington, DE) sprayed on the upper insi de surfaces of cages to prevent insect escape. Ad ults were transferred to new plants every 2 wk. Cages were maintained in the greenhouse with a 16L:8D photoperiod and 25-30 C. Between 300 to 400 chinch bugs developed on each plant. Meehan and Wilde (1989) also successfully reared on pearl millet in the greenhouse (21 32C) and in growth chambers (24 30C) with a 16L:8D photoperiod. Baker et al. (1981) attempted to rear the hairy chinch bug, ,using Parker and Randolphs (1972) technique, but early-instar mortality was high, which appeared to be associated with fungal growth on the corn sections. When sections of young maize plants were treated with 2% sodium hypochlor ite (instead of 1.0% benzalkonium chloride) and placed in 236.6-ml cardboard cartons in growth chambe rs [16L:8D photoperiod, at 26C, and 40-75% RH], was reared year round (Baker et al. 1981). survival from egg to adult increased to 80%. Busey and Zaenker (1992) maintained populat ions of the southern chinch bug on 10-20 stolon cuttings (~100 mm long with three to four nodes) of sus ceptible Florida Common St. Augustinegrass for host-plant resist ance studies. Insects were c onfined in plastic bins (14.5 18.0 9.0 cm deep) covered with a double sheet of cellulose tissue (Kimwipes, Kimberly-Clark, Roswell, GA) glued to the tops of the bins. St olon cuttings were placed in water-filled glass vials that were sealed with parafilm and were replaced at least once a week (Busey and Zaenker 1992). Percentage survival, the number of generations produced, and the existence of overlapping generations were not re ported. It is possible that th e insects were only maintained long enough to complete the study. 22

PAGE 23

Anderson (2004) reared on 15-cm pots of Ralei gh St. Augustinegrass in a potting mixture of sand-soil-peat-per lite in a 2:1:3:3 ratio. Plants were covered with ventilated tubular 15 45 cm plastic cages that were embedded 2-3 cm into the soil. The cages were sealed with organdy fabric and sand was placed around the bottom of the cages to prevent insect escape. Infested plants were kept in a growth cham ber at 28 2C with a 24L:0D photoperiod and 40 75% RH. As the plants began to die, insects we re sifted through a 2-mm mesh screen, aspirated, and placed on new plant material. was reared for five generations but the population peaked at a total of 500 insects and rapidly declin ed (Anderson 2004). Spider predation in the cages, limited air movement and fungal development due to caging negatively affected the population, and constant li ght may not have been suitable for development (Anderson 2004). Anderson (2004) also reared and with the procedures described by Wilde et al. (1987). This method a llowed the use of whole pl ants instead of stalk sections and did not require tr eating plants (Parker and Randol ph 1972, Baker et al. 1981). One pot could support ~400 chinch bugs for about 3 wk. However, greenbug [ (Rondani)] populations would rapi dly build, crowding out preferre d chinch bug feeding sites and excreting copious amounts of honeydew, resulting in sooty mold (Anderson 2004). Several authors successfully produced > 1 generation of spp. under greenhouse and growth chamber conditions (Wilde et al 1987, Meehan and Wilde 1989, Anderson 2004). However mass-rearing of in our greenhouse has not been feasible. Daily ambient summer temperatures in three of the greenhouses we used have exceeded 37.8 C, which is lethal for (personal observation), and St. Augustineg rass pots have become infested with aphids, thrips, scales, mites, other populations, and natural enemies. 23

PAGE 24

Reinert (1978) observed spiders ( sp.) and predatory insects such as Stal (Hemiptera: Nabidae), (Reuter) (Hemiptera: Anthocoridae), Reuter (Hemiptera: Anthocoridae), spp. (Hemiptera: Reduviidae), Pallas (Dermaptera: Labiduridae), and (F.) (Hymenoptera: Formicidae), feeding on but none were able to suppress populations below damaging levels. Another pest in lawns, the red imported fire ant, Buren, is also ineffective at controlling populations (Cherry 2001c). (Balsamo) Vuillemin was pathogenic to all life stages of ; however, was only present when moisture and humidity levels were high (Reinert 1978). Woods (2001) tested the virulence of on healthy via direct contact and was only able to obtain a 6.1% rate of transmission. (Say) and (Say) (Hemiptera: Geocoridae) have been observed preying on (Reinert 1978). (Say) also found in Florida turfgrass (Mead 2004), can prey on up to 19 firstto third-instar nymphs in 24 h under laboratory conditions (Congdon 2004). However, young nymphs can hide within the sheaths of St. Augustinegrass, thus avoiding predation by spp. When were given a choice between pea aphids (Hemiptera: Aphididae) and nutritionally superior eggs of the corn earworm (Lepidoptera: Noctuidae), consistently attacked (Eubanks and Denno 2000). The authors reported that prey mobility, not prey nutritional quality, ap peared to be the most important criterion used by in choosing their prey. Geocoridae are generalist predators and are cannibalistic (Chiravathanapong and Pitre 1980, James 2004, Mead 2004). Also, Geocoridae in turfgrass can 24

PAGE 25

be mistaken for and unnecessary insecticide appl ications can ensue (Caplan 1968, Mead 2004). Perhaps the natural enemy that holds the most promise for controlling can be found in the family Scelionidae. spp. (Hymenoptera: Sce lionidae) are parasitic wasps (Figure 1-8 A-B) that attack the eggs of several spp. (McColloch and Yuasa 1914, 1915; Dicke 1937; Reinert 1972; Wright and Danielson 1992; Sadoyama 1998). Gahan attacks the eggs of year-round in Florida; Reinert (1972) found an average abundance of 35 wasps/0.1 m2 in lawns containing populations of 90.0/0.1 m2 (Reinert 1972). However, research needs to be done to determine host-specificity and suitability for use in th e biological control of Tolerance of is a major consideration in selection of St. Augustinegrass cultivars, as the success of Floratam demonstr ates. Floratam is an improved cultivar of St. Augustinegrass that was released as a chinch bugresistant cultivar in 1973 by the University of Florida and Texas A&M (Anonymous 1973, Buse y 1979, Horn et al. 1973, White and Busey 1987, Trenholm and Unruh 2005). After its releas e, Floratam was confirmed resistant to (Reinert et al. 1980, Crocker et al. 1982, Reinert et al. 1986) and it quickly became the most widely used cultivar in South Flor ida (Busey 1986, Busey and Center 1987). Today, Floratam is the most widely produced and used cultivar of St. Augustinegrass and accounts for 80% of sod production in Florida (H aydu et al. 2005). It has a very coarse leaf texture, and poor cold and shade tolerance (Trenholm and Nagata 2005), but is resistant to the St. Augustine decline virus, and successfully minimized problems for years (Busey 1979, Busey and Center 1987). However, has overcome resistance to Floratam in Florida and Texas (Busey and Center 1987; Crocker et al. 1989; Busey 1990a, 1990b). Plant breeders 25

PAGE 26

developed FX-10 St. Augustinegrass that was registered in 1993 and was resistant against (Busey 1993). However, it was never exte nsively grown because of its very coarse texture, toughness, and poor ground coverage (Busey 1993, Nagata and Cherry 2003). Two additional cultivars, NUF-76 and NUF-216, were de veloped and found to be highly resistant to (Nagata and Cherry 2003, Rangasamy et al. 2006). NUF-76, now known as Captiva, has a lush, dark green color and can be mowed less frequently. It is now in production for use in lawns, but its long term effectiveness is yet to be determined. Proper cultural practices such as fertilization can help to pr omote healthy grass, which may be able to better tolerate chin ch bug damage. Current nitrogen recommendations suggest a range from 100 to 300 kg N ha-1 yr-1 to maintain St. Augustinegrass lawns in Florida, depending on soil type, geographical location in the state, time of year, amount of shade in the lawn, and the preference for a high-or low-i nput lawn (Trenholm and Unruh 2007) Applications must be made carefully as heavy fertil ization may cause excessive turf grass growth (Christians 2004), thatch development, reduced tolerance to en vironmental stresses (E mmons 1995), and nutrient leaching (Kelling and Peterson 1975, Weaver et al. 1988, Petrovic 1990, Coale et al. 1994, Correll 1998, Sims et al. 1998, Wulff et al. 1998, Florida Department of Environmental Protection 2002, Sinaj et al. 2002, Park et al 2008). Phytophagous insects such as meet nutritional requirements by feeding on heal thy host plants (Heinrichs 1988). Nitrogen appears to be a crucial, limiting factor in ins ect growth and survival (Mattson 1980, Scriber and Slansky 1981, Heinrichs 1988). With this said, high fertility with readily available N applied to St. Augustinegrass results in higher populations than gra ss treated with lower N amounts (Horn and Pritchett 1963, Busey and Snyde r 1993). Busey and Snyder (1993) noted that early population regulation effects, such as the number of eggs per fe male per week and rate 26

PAGE 27

of development of early instars, may explain the observed response of to high fertilization with quick -release fertilizers. Irrigation and mowing are also important components in maintaining healthy St. Augustinegrass. Irrigation should occur on an as-needed basis (Emmons 1995, Trenholm and Unruh 2005), with St. Augustinegrass being watered at the first sign of rolling leaf blades, wilting, and/or footprints that remain on the lawn At these signs of water deficit, applying 1.3 1.9 cm of irrigation to the entire lawn should su pply water to a depth of ~15-23 cm for most Florida soils. Watering in this manner will help to encourage deep root growth (Emmons 1995). Excess irrigation may lead to problems such as a shallow root system, increased pest problems, and increased thatch (Emmons 1995, Trenholm a nd Unruh 2005). Mowing too infrequently can also cause a thatch buildup and excessive thatch (exceeding 2.5 cm) may need to be professionally removed by mechanical thatch rem oval (verticutting or ae rification) (Christians and Engelke 1994, Christians 2004). Also, mowe r height should be properly set to avoid possible scalping. Most St. Augus tinegrass cultivars should be mowe d to a height of 8-10 cm. Newer semi-dwarf varieties can be mowed to a height of 3.8.3 cm (Trenholm and Unruh 2005). Control of damaging populations is mainly achiev ed through insecticide use. Currently, 20-25 per 1 ft2 warrant cont rol (Short et al. 1982). Lawn care companies have at times made up to six to twelve insecticid e applications a year to control this pest in Florida (Reinert 1978, Reinert and Niemczyk 1982). Insecticides historically used against include tobacco dust, calciu m cyanide, nicotine sulfat e, DDT, parathion, dieldrin, aldrin, chlordane, chlorpyrif os, propoxur, diazinon, and bifent hrin (Beyer 1924b; Watson and Bratley 1929a, 1929b; Kelsheimer 1952; Wolf enbarger 1953; Kerr 1956; Brogdon and Kerr 27

PAGE 28

1961; Reinert 1982a, 1982b; Reinert and Portier 1983; Cherry and Nagata 2005). Currently, organophosphates, carbamates, pyrethroids, neonic otinoids, and combination products are used for control. The following provides a brief description of each insecticide class and their mode of action. The synthesis of organophosphates (OPs) bega n in the 1800s; howev er their potential toxicity went unrecognized until the 1930s (Chambers et al. 2001). By 1940, work conducted in England and Germany had produced several hi ghly toxic compounds for possible use as chemical warfare agents. The most notable wo rk was done in Germany by Gerhard Schrader with the development of nerve gases during Worl d War II. It was not until the capture of Schraders research records that interest in OP insecticides grew (Chambers et al. 2001). OPs are derivatives of phosphoric acid and are highly toxic (Yu 2008). There are several subclasses of OPs but, in general, they are considered to be biodegradable and nonpersistent. OPs exert their toxic action by i nhibiting acetylcholinesterase, an enzyme that occurs in the central nervous system (Scharf 2003, Yu 2008). Under normal circumstances, acetylcholinesterase removes acety lcholine from its postsynaptic r eceptor, resulting in hydrolysis of acetylcholine into acetate and choline. This initiates at precise inte rvals electrical impulses (action potentials) that travel along neurons and provide the basi s of nervous system function (Scharf 2003). OPs attach to acetylcholinesterase resulting in prolonged binding of acetylcholine to its postsynaptic receptor, leading to deat h in the organism from prolonged neuroexcitation (Scharf 2003). The reaction with OPs and acetylc holinesterase is very slow, typically taking days or even weeks (Yu 2008). This is because OPs must be enzymatically activated by cytochrome P450s before they can effectively in hibit acetylcholinesterase and also because the nature of the OP-acetylcholinesterase interaction (Scharf 2003). As a result, OPs can become 28

PAGE 29

irreversible inhibito rs of acetylcholinestera se. The only OP currently registered for control is trichlorfon (Dylox, Bayer Environmental Science, Research Triangle Park, NC). Carbamates are esters of carbamic acids (P limmer 2001). Their herb icidal and fungicidal activities were demonstrated in the early 1930s, but interest in inse cticidal activity did not begin until the mid-1950s (Ecobichon 2001). Carbamates are slightly to moderately soluble in water, moderately volatile, and readily bi odegradable (Yu 2008). They are used to control a wide range of chewing and sucking insects. Like the OPs, carbamates are acetylcholinesterase inhibitors. However, carbamates are faster inhibitors than OPs, and are generally more hazardous although their effects are readily reve rsible (Scharf 2003). Carbaryl (Sevin, Bayer Environmental Science, Research Triangle Park, NC) was the first carbamate to be commercially developed (Wickham 1995) and is the sole chemical in th is class that is avai lable for control of Pyrethroids are synthetic insecticide deriva tives of pyrethrum. Pyrethrum extracts are obtained from chrysanthemum flowers ( ). Originally grown in the former Yugoslavia, the ground, dried flower heads were later known as Dalmation Insect Powder and the product was used to control body lice during the Napoleonic Wars (van Emden and Service 2004). Over the years, research was done to find synthetic replacements for agricultural use that ha d greater photochemical stability a nd longer field life (Plimmer 2001). The first pyrethroid, allethrin, was developed in 1949 (Ware and Whitacre 2004). Pyrethroids are divided into two groups, type I and type II; the difference between the two being that type II pyrethroids have an -cyano group (Yu 2008). Type I pyrethroi ds permethrin and bifenthrin are used for control of Type II pyrethroids used to control include betacyfluthrin, cypermethrin, deltamethrin, and -cyhalothrin. Pyrethroids are slightly soluble in 29

PAGE 30

water and have mini mal volatility. They have exceptional photostability and generally provide good residual control (Yu 2008). Py rethroids are also lipophilic a nd adhere strongly to organic matter (Elliott et al. 1978, Laskow ski 2002). Because of these char acteristics, pyrethroids are widely used to control agricultural pests. Pyrethroids interfere with voltage-gated sodium channels of both the peripheral and central nervous system. Voltage-gated sodium cha nnels are responsible for the initiation and perpetuation of action and receptor potentials in neurons (Scharf 2003). Pyrethroids affect sodium channels by causing activatio n at lower thresholds or inac tivation later than would occur under normal circumstances, resultin g in prolonged flow of sodium currents into neurons and excessive neuroexcitation (Scharf 2003). Three generations of chemicals are involved in the history of nico tinoids (Yamamoto and Casida 1999), the first of which was nicotine. Nicotine was extracted from tobacco in 1828 by Posselt and Reimann and was named after Jean Nicot, who introduced tobacco to the French court around 1560 (Posselt and Reimann 1828, Ujv ry 1999). Due to its high toxicity to mammals, research continued in search of more sele ctive compounds. The second generation of chemicals emerged in 1970 when the Shell Development Company was investigating heterocyclic nitromethylenes as potential insecticides (Sheet s 2001, Tomizawa and Casida 2005). Nitromethylenes had potency, selectivity, and systemic properties but were not photostable (Yamamoto and Casida 1999). Imidacloprid is the first commercial product of the third generation of chemicals, the neonicotinoids. Im idacloprid was developed by Bayer in 1984 and has greater insecticidal activity and lower mamm alian toxicity than its predecessors (Sheets 2001). Neonicotinoids generally have low toxi city to mammals (Thyssen and Machemer 1999, Yamamoto 1999, Anatra-Cordone and Durkin 2005, Tomizawa and Casida 2005, birds (Anatra30

PAGE 31

Cordone and Durkin 2005, Tomizawa and Casida 2005), and fish (TDC Environm ental 2003, Tomizawa and Casida 2005, Jemec et al. 2007). Since the introduc tion of imidacloprid, neonicotinoids have become the fastest-growing class of insectic ides introduced to the market since the commercialization of pyrethroids (Nauen and Bretsc hneider 2002, Jeschke and Nauen 2005). Neonicotinoids are broadspectrum insecticides that possess contact, stomach, and systemic activity (Jeschke and Nauen 2005). Neonicotinoids labeled for control of include imidacloprid, clothianidin, dinotefuran, and thiamethoxam. Neonicotinoids act on the insect central ner vous system as agonists of the postsynaptic nicotinic acetylcholine receptors (nAChRs) (Jeschke and Nauen 2005). nAChRs and muscarinic acetylcholine receptors are considered the two major acetylcholine receptors based on their sensitivity to agonists (Nauen et al. 2001, Scha rf 2003). nAChRs are agonized by nicotine and muscarinic acetylcholine receptors are agonized by the mushroom toxin muscarine. nAChR is a highly insect-specific target site. Compared to the ~250:1 ratio of mu scarinic to nicotinic receptors in the mammalian central nervous syst em, insects have more than 10 times as many nicotinic as muscarinic receptors (Scharf 2003). Neonicotinoids mimic acetylcholine by acting as agonists to activate the nAChR, causing an influx of sodium i ons and the generation of action potentials (Yu 2008). Under normal conditions, the synaptic action of acetylcholine is terminated by acetylcholinesterase which hydrolyzes the neurotransmitter. However, neonicotinoids are not hydrolyzed by acetylcholines terase and the persistent activation leads to hyperexcitation, convulsion, paralysis, a nd death of the insect (Yu 2008). B. insularis Limited research on the eff ects of natural enemies on lack of host-plant resistance options for the last two decades, and minimal and/or improper use of cultural methods have helped generate near-constant reliance on chemi cal control of populations. As 31

PAGE 32

a result, has developed resistance to organochl orines, organophosphates, carbamates, neonicotinoids, and pyrethroi ds (Wolfenbarger 1953; Ke rr 1958, 1961; Reinert 1982a, 1982b; Reinert and Niem czyk 1982; Re inert and Portier 1983; Cherry and Nagata 2005, 2007). Resistance is defined as a heri table physiological and/or beha vioral (Sparks et al. 1989) adaptation that confers a selectiv e advantage in the presence of a pesticide, and that leads to control failures (Sawicki 1987, ffrench-Constant and Roush 1990). In 2006, there were 550 arthropod species resistant to one or more pesticides worldwid e (Onstad 2008, Whalen et al. 2008). Pesticide resistance has been estimated to have a global annual eco nomic impact of over $4 billion dollars (Pimentel et al. 1991, Mota-Sanchez et al. 2002). Since th e first report of pesticide resistance in 1914 (Melander 1914), sign ificant progress has been made in 1) the development of methods for detection and documenta tion of resistance, 2) research on resistance mechanisms, 3) the identification of biotic, ge netic, and operational f actors influencing the evolution of resistance, and 4) pest management practices that incorporate resistance-delaying measures (Georghiou and Saito 1983). Early detection and documentation of insectic ide resistance is an important step in resistance management because if resistance can be detected before control failures occur, then preventive measures possibly can be impl emented (ffrench-Constant and Roush 1990). However, it is first important to have a standardized method for ev aluating toxicity to the insect in question. The toxicity of an insecticide is typically determin ed using concentrationor doseresponse bioassays with response expressed in terms of lethal concentration, dose, knockdown, or time (Yu 2008). Evaluation of toxicity to is typically expressed as lethal concentration (LC50 and/or LC90) with the response being death (concentration-mortality). For 32

PAGE 33

analysis, the raw data are transformed to a log scale, form ing a sigmoid or S-shaped curve (Yu 2008). The percentage mortalit y is then converted to probits using probit (for normally distributed data) or logit analysis to create a l og concentration-probit (or logit) line from which the LC50 is estimated (Yu 2008). Ideally, five concen trations that cause mortality ranging from 5-95% with at least 100 insect s per concentration should be used for estimation of LC50 (Robertson and Preisler 1992, Robe rtson et al. 2007). When c onducting concentration-mortality tests it is important to choose an appropriate bioassay for measuring response. It is also important to standardize insecticide bioassays to reduce variability. Attributes to standardize bioassays inlcude temperature, da ylength, relative humidity, sex of the insect, age of the insect, previous exposure(s) to other chemicals, and subs trate. This will ensure that the observations made are due to the effect of the ins ecticide and not to some other variable. Most insecticide bioassays can be classified based on the manner in which the pesticide is applied (i.e., dipping, topical, feed ing, or residual). The dipping method involves dipping insects in solutions of known concentration (Yu 2008). This method is often regarded as unrealistic or imprecise because the results cannot be expressed in terms of toxicant per gram of body weight (ffrench-Constant and Roush 1990). While dippi ng tests ensure uniform contact, it would be extremely time consuming when working with small insects such as Topical application involves applying a known dose of insecticide direc tly to individual insects via microsyringe (Yu 2008). However, as with th e dipping method, topical application is time consuming when dealing with small insects and the exact amount of pest icide penetrating the insect is not known. Feeding assays entail prov iding immature insects with insecticide-treated diets (Yu 2008). The residual or contact method cons ists of exposing insects to a dry residue of pesticide on a natural (e.g., leaf) or artificial (e.g., glass, filter paper) substrate (ffrench-Constant 33

PAGE 34

and Roush 1990). Some researchers have reported that tests such as le af-residue assays can im prove the accuracy of resistance detection as we ll as help to establish the relationship between laboratory bioassays and field-control failures (ffrench-Cons tant and Roush 1990). However, others have expressed concern th at insects may be repelled by trea ted leaves and never come in contact with the insecticide, and that the exact dose accumu lated by the insect is unknown (Brown and Brogdon 1987, ffrench-Constant and Roush 1990). Nonetheless, the residual method has been used in many hemipteran studi es (Studebaker and Kring 2003, Snodgrass et al. 2005, Fleury et al. 2007), including (Reinert and Por tier 1983; Congdon and Buss 2004, 2006; Cherry and Nagata 2005, 2007). The most commonly used residual bioassay for evaluating insecticide efficacy in is the sprig dip (Reinert and Portie r 1983; Congdon and Buss 2004, 2006; Cherry and Nagata 2005, 2007). This method involves cutti ng sections of St. Augustinegrass stolons, dipping them into insecticide solutions, allowing th em to dry, and placing them into petri dishes containing ten adult The set up for the sprig-dip bioassay can be conducted quickly and it is inexpensive. However, a large degree of variability in response occurs. Tests are usually conducted in different laboratories under varying environmental conditions, or with fieldcollected of unknown age and/or from different locations (Reinert and Portier 1983; Cherry and Nagata 2005, 2007; Congdon and Buss 2006; Chapter 2). It woul d be beneficial to evaluate the sprig-dip bioassay under more standardized conditions to va lidate the use of the assay. In addition to variability in the sprig-dip bioassay, scoring multiple individuals in the same dish can be cumbersome when they are not a ll moribund. A more standardized bioassay that could detect differences between male and female would greatly aid in 34

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understanding how insecticide resistan ce develops in this pest (i.e., mode of inherita nce, stability of resistance). Replication and reliability of insecticide bi oassays can depend on the stability of the environmental conditions (e.g., light, temperatur e, and humidity) (Sun 1960, Rozman et al. 2001) and the insects used for testing (Sun 1960). Any variability in the biological and/or environmental factors in a bioassay may change th e insects susceptibility to an insecticide. These factors can be divided into two categor ies, intrinsic and extrinsic (Busvine 1980). The insect species, stage, and strain used may generate variability in insecticide bioassays (Sun 1960). Data from one species cannot be used to detect resistance in another as differences in susceptibility may occur. However, related sp ecies tested using the same technique may result in similar susceptibility levels (Busvine 1980). Choosing the appropriate life stage is also important. Sun (1960) noted that inactive stages (eggs and pupae) are usually more tolerant of an insecticide than active stages (immatures a nd adults). Also, using homogeneous susceptible or resistant laboratory colonies will help to reduce variability in bioassays. Other intrinsic factors that may introduce variab ility into insecticide bioassays include age, size, sex, and nutrition (Rozman et al. 2001). Adult insects are ofte n more susceptible just after molting, followed by a period of greater tolerance, and then increasing suscep tibility with advancing age (Busvine 1980). In immature insects, tolerance to insectic ides increases as the insect gains weight (ie., first inst ar vs. fifth instar). In additio n, male insects tend to be more susceptible to insecticides than females (Sun 1960). Insects in a single field population or laboratory colony have also shown changes in re sponse to insecticides depending on the time of 35

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day and/or season (Sun 1960). Also, well-nouris hed insects will generally be more tolerant to insecticides than malnourished ones. Temperature, humidity, and light can possibly alter tests by changing the effectiveness of the insecticide (Busvine 1980, Rozman et al. 2001) and/or insect activity (Busvine 1980). The relationship between temperature and insecticide toxicity in insects is well documented (Scott 1995, Valles et al. 1998), particularly for DDT an d pyrethroids. DDT (Vinson and Kearns 1952) and type I pyrethroids may show a negative temperature coefficient, meaning the insecticide becomes more toxic to insects with decreasing temper ature (Wadleigh et al. 1991, Valles et al. 1998, Musser and Shelton 2005). Type II pyrethroids may show a positive, negative (Scott and Matsumura 1983, Sparks et al. 1983), or no temp erature coefficient (Toth and Sparks 1990). Most carbamates and organophosphates have a pos itive temperature coefficient (Norment and Chambers 1970, Chalfant 1973, Grafius 1986, Li et al. 2006b), as well as the neonicotinoid imidacloprid (Elbert et al. 1991, Richman et al. 1999) Low humidity is a nother extrinsic factor that can alter the outcome of insecticide bioassa ys because it can cause desiccation of sensitive insects (but should also be evident in the contro ls). Increased or constant light conditions may lead to greater insect activity and as a result, the insects would come into contact with the insecticide faster in residu al tests (Busvine 1980). While insecticide resistance has been documented in populations in Florida, these conclusions were made based on tests either conducted in th e field under varying environmental conditions (Kerr 1958, 1961) or unde r laboratory conditions using field-collected insects of unknown age and sex (Reinert 1982a; Re inert and Niemczyk 1982; Reinert and Portier 1983; Cherry and Nagata 2005). A more sta ndardized method of testing conducted under 36

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laboratory conditions is needed to better understand the development of insect icide resistance in There are several ways that insects can devel op resistance to insectic ides and these can be classified into two main categ ories: behavioral and physiologi cal. Behavioral resistance is defined as the ability to avoid a dose or concentration th at would prove lethal (Yu 2008). It is thought that insects with behavioral resistance contain receptors which can better detect low concentrations of insecticides than normal insects (Yu 2008). Physiological resistance has been shown to involve three factors: reduced penetration, target site insensitivity, and metabolism (increased detoxification) (Yu 2008) Penetration resistance occurs when resistant insects absorb toxins through the cuticle more slowly than su sceptible insects. Reduced penetration by itself results in only slight resistance (Soderlund a nd Bloomquist 1990, Yu 2008). However, in the presence of other mechanisms, reduced penetra tion confers considerable resistance to some insecticides (Yu 2008). Target site insensitivity usually involves point mutations (the replacement of one nucleotide by another [Hoy 2003]). There are three types of target site insensitivity involved in insecticide resistance in insects: nerve insensitiv ity, altered acetylcholines terase, and reduction in midgut target site binding (Yu 2008). Nerve insensitivity is involved in organochlorine, pyrethroid, neonicotinoid, and phenylpyrazole insecticide resistance in many insects. For example, resistance to cyclodienes in (ffrench-Constant et al. 1993) and fipronil in diamondback moths (Li et al. 2006a) was due to a point mutation (substitution of alanine to serine) of the GABA receptor prot ein, causing receptor insensitivity. Also, knockdown resistance (kdr) to pyrethroids in was due to several point mutations in the sodium channel gene (Yu 2008). Altered acetyl cholinesterase is associ ated with resistance 37

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to organophosphate and carbamate insecticides. This type of resistance o ccurs in several insect and acarine species, including cattle ticks, fall armyworms, houseflies, green rice leafhoppers, mosquitoes, tobacco budworms, and two-spotted spider mites (Smissaert 1964; Fournier and Mutero 1994; Gunning and Moores 2001; Yu 2006, 2008). Examples of reduction in midgut target site binding include insects resistant to (Bt). Ferre and Van Rie (2002) reported that re duced binding of toxin is a primary mechanism of insect resistance to the Cry proteins of Bt, but some in sects are able to alter th e sugar structure of the glycolipid (receptors for Bt toxin) molecule so th e Bt toxin cannot attach itself, and as a result, become resistant (Griffitts et al. 2005). Metabolic resistance results when an insect det oxifies and excretes the toxin faster than a susceptible insect, enabling the re sistant insect to quickly rid it s body of the insecticide. Three detoxification enzymes associated with re sistance in insects are cytochrome P450 monooxygenases, hydrolases, and glutathione S-tran sferases (GSTs) (Yu 2008). Resistance to insecticides can be due to enhanced oxida tive metabolism caused by cytochrome P450 monooxygenases. This important enzyme is non-sp ecific to organic com pounds and can result in cross-resistance to other insec ticides (Yu 2008). Carboxylesterase s (hydrolases) are involved in resistance to ester-containing insecticides su ch as organophosphate, carb amate, and pyrethroid insecticides (Yu 2008). GST is a phase II enzyme associated w ith resistance to nearly all pesticide classes. Cross resistance refers to a situation in whic h an insect population b ecomes resistant to two or more insecticides (with different active ingr edients) as a result of selection by a single insecticide (Winteringham and Hewlett 1964). Multiple resistance occurs after simultaneous or 38

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successive exposure to two or mo re insecticides. Resistan ce mechanisms are not known for The development of resistance is determined by a variety of genetic, biological or ecological, and operational factor s (Georghiou and Taylor 1986). Genetic factors would include the number, frequency, and/or dominance of resistant alleles; past selection with other chemicals; and the extent of integration of the resistance genes with fitness factors. Important biological factors include time per generation, offspring per generation, monogamy or polygamy, mobility, diet, and refugia (Georg hiou and Taylor 1977a). can be difficult to control in Florida because it produces multiple generations per year, has a high number of offspring per generation, is highly mobile and encroaches onto neighboring lawns, is able to survive on other grass sources until new St. Augustinegrass is lo cated, and is able to avoid insecticides. Operational factors that lead to resistance are those related to the application of pesticides and include the dosage used, treatment history, tr eatment schedule (rotation or no application), treatment thresholds, life stage selected, and method of application (Georghiou and Taylor 1977b). Operational factors are considered und er human control and their manipulation may help to delay the onset of insecticide resistance. Multiple insecticide applications are made each year to control damaging populations; however, it ha s been unclear whether treatment history plays a role in development of insecticide resist ance in this pest. Also, with respect to treatment history, St reu (1973) suggested that excessi ve pesticide usage may cause stress in turfgrass that contributes to accumulation of thatch, possibly providing insect pests shelter from insecticides. 39

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Roush (1989) has suggested that if created at the earliest opportunity, a properly-structured resistance management (RM) program can be de veloped without having made a serious error in recommendations. Even if based on limited information of the insecticides used and the population dynamics of the pest involved, it may be possible to improve the design of the RM program as new information (ie., mechanisms, cross-resistance, mode of inheritance, and stability of resistance) is obt ained (Roush 1989). However, the research involved in acquiring this information and the time needed to implemen t it into a RM program can take several years, and relies on employing the correct genetic model (Hoy 1995). Several models have been developed that evaluate options for RM and try to predict ho w quickly a pest will develop resistance if certain conditions are met. There are four RM management modeling appr oaches: analytical, simulation, optimization, and empirical (Tabashnik 1990). Analytical models (Tabashnik 1990, Hoy 1999) use simple mathematical descriptions and attempt to an alyze general trends to define fundamental principles. These models do not provide realisti c details and are relatively simple. Analytical models assume that insect population dynamics are simple with discrete generations and no age structure. Also, population growth is usually de termined by some form of the logistic equation (Tabashnik 1990). However, few arthropods have discrete generations and may be prone to developing resistance (Hoy 1999). Also, many insects, such as are multivoltine and have overlapping generations. Simulation models are more complex and realistic as they attempt to assess the influence of a large number of factors (e.g., biology, behavior, and ecology of the population) (Tabashnik 1990, Hoy 1999). These models may contain complex population dynamics, including age 40

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structure, overlapping generations, and temporal and spatial variation in pe sticide dose. Simulation models can be used to evaluate differe nt options for delaying re sistance by including empirical data in the parameters included in the m odel. Parameters can be varied in a systematic way to determine how important each is. Ho wever, the details of the population biology, ecology, and structure may influence the rate of resistance development. These models may become extremely complex or diffi cult to simulate field conditions. Optimization models focus on economic analysis and evaluate which management strategy will maximize profit when pest susceptibility to a pesticide is considered a non-renewable natural resource. This approach ai ms to balance the future cost of reduction in pest susceptibility with the present losses in crop yield due to the e ffects of the target pest. However, information on the target pest is simplified (biology, ecology, behavi or) and is often viewed as a constraint (Tabashnik 1990). As a result, optimization models may not properly predict the longevity of a product and lead to inaccurate predictions of th e costs of losing a specific product (Hoy 1999). Empirical models are based on actual observa tions among variables and no assumptions are made about causal mechanisms (Tabashnik 1990) These models are derived from data and may only be appropriate for the specific conditions of the observed populations (Tabashnik 1990, Hoy 1999). Empirical models may not be usef ul for developing a st rategy for delaying resistance in an unknown situati on if it is assumed that the important variables (mode of inheritance, cross resistance, fitness costs, a llele frequency, and selec tion intensity) can vary between populations (Hoy 1999). Mitigation models involve the use of mixtures, mosaics, rotations, natural enemies, and/or high-dose strategies (Tabashnik 1990, Hoy 1999). For mixtures to be appropriately applied, resistance to each product should be monogenic. No cross resistance can occur between 41

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products in the mixture and they m ust have equal persistence. Also, some of the population must remain untreated (refuge). Mitiga tion models also assume that resistant individuals are rare in the population and that resistance is functionally recessive. While mixtures exist for control of it would be difficult to provi de untreated refuges for this pest due to the amount of damage their feeding can cause. Also, as with the previous models, the genetic basis of insecticide resistance in is not known. With a mosaic strategy, susceptible individuals are maintained and able to move into surrounding areas; this model may require negative cross-resistance or f itness costs associated with re sistance (Tabashnik 1990, Hoy 1999). Rotation strategies assume the frequency of individuals resistant to one product will decline after the application of an alternative pro duct, which is true if there is negative crossresistance, a fitness cost associated with the resistance, and/or immigration of susceptible individuals occurs (Hoy 1999). Natural-enemy strategies may be used if food limitations are sufficient to constrain the ability of natural enem ies to develop resistance in the field. The highdose strategy assumes complete coverage, effectiv e kill of all individuals, and ignores negative effects on secondary pests, natural enem ies, or the environment (Hoy 1999). Hoy (1995) suggested that the de velopment of resistan ce is likely inevitabl e and at best we can only delay the onset of resistance in orde r to preserve existing products. Long-term resistance management must be a broad-base d multitactic endeavor, in which resistance management is combined with integrated pest management (IPM) and involv es altering pesticide use patterns (Hoy 1995). IPM was first developed by Stern et al. (1959) for control of spotted alfalfa aphid, (Buckton) (Homoptera: Aphididae), in alfalfa in California. The authors noted that IPM included a variet y of tactics, involving monitoring, assessing economic injury levels, use of selective pestic ides, and integrating chemical and biological 42

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control (Stern et al. 1959). While some aspects of IPM exist for (biological, cultural, and chemical control), research on some of these aspects has been limited. Historically, once develops resistance to an ins ecticide (bifenthrin being the mo st current), that insecticide is replaced by another without an understanding of mechanisms, cross-resistance patterns, mode of inheritance, or stabil ity of resistance. The distribution of bifenthrin resistance in Florida is not known. Several other conventi onal and newer insecticides are currently available for control; however, baseline susceptibilities to them are also not known. In addition, it is unclear how effectivene the sprig-dip assay is for systemic insecticides and variability in this bioassay needs to be reduced. A resistance management pr ogram needs to be developed for this pest. However, it is important to obtain initial information upon which to build a foundation. With the above mentioned rationale in mind, the objectives of this research were to: (1) sample select populations in 2006 and 2008 in nor thern and central Florida to describe their susceptibility to bifenthrin, docum ent new locations of bifenthrin resistance to bifenthrin, and evaluate another pyre throid, permethrin (Chapter 2), (2) develop a synchronous rearing method for that produces insects of known age and generation (Chapter 3) and (3) develop an improved bioassay that could be used for detecting insecticide susceptibility differences between male and female evaluate and validate both the sprig-dip and the new bioassay under standardized cond itions, and determine optimal exposure times and sample sizes to be used for each bioassa y for selected insecticides (Chapter 4). 43

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Figure 1-1. Severe damage from feeding (right) that stops at the neighboring bahiagrass lawn (left) (Photo credit: E. A. Buss). 44

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45 Figure 1-2. Images showing A) brachypterus and B) macropterus male (left) and female (right) respectively. The ventra l tip of the abdomen of C) male and D) female (Photo credit: L. Buss).

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Figure 1-3. Photograph of A) healthy egg in early development, and B) healthy egg in late developmen t (Photo credit: L. Buss). 46

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Figure 1-4. A) first, B) second, C) third, D) fourth, and E) fifth instars (Photo credit: L. Buss). 47

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Figure 1-5. Lawns damaged by [Photo credit: A) Rick Lewis, B) and D) J. C. Vzquez, and C) R. Levin]. 48

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Figure 1-6. St. Augustinegrass lawns with populations encroaching on neighboring lawns (Photo credit: R. Clemenzi). 49

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Figure 1-7. St. Augustinegrass with excessi ve thatch (Photo cr edit: R. Clemenzi). 50

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51 Figure 1-8. Photographs of A) a egg parasitized by and an B) adult (Photo credit: L. Buss).

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CHAPTER 2 SUSCEPTIBILITY OF B. POPUL ATIONS IN FLORIDA TO BIFENTHRIN AND PERMETHRIN St. Augustinegrass ( [Walt.] Kuntze) is the most widely used lawn grass in tropical and subtr opical climatic regions (Sauer 1972) It is the primary turfgrass in residential lawns and comprises ~70% or 1.2 million ha in Florida (Hodges et al. 1994, Busey 2003). The southern chinch bug, Barber, is considered the most damaging insect pest of this grass (Kerr 1966, Reiner t and Kerr 1973, Reinert a nd Portier 1983, Crocker 1993). Kerr (1966) speculated that was one of the most economically important plant feeding arthropods in Florida, being second only to the citrus rust mite in amount of money spent for control. By 1983, the combined annual lo sses and cost in Florida to manage this pest was estimated at $5 million (Hamer 1985). Given that the number of hou sing units in Florida increased from ~3.9 million in 1980 to 8.5 million in 2006 (an increase of 118%), the potential for damage and increased cost for management is likely higher now. With over 18 million people and an annual growth rate of 1.8% (U .S. Census Bureau 2006), the demand for quality turf and maintenance in Florida continues to increase (Haydu et al. 2005). Florida is second only to California in terms of employment impacts of the turfgrass industry, providing 83,944 jobs in 2002 (Haydu et al. 2006). Similar to other feeding habits, nymph and adult damage St. Augustinegrass by feeding in the phloem sieve el ements of the grass (Rangasamy et al. 2009) causing wilting, chlorosis, stunt ing, and eventually death (Painter 1928, Negron and Riley 1990, Spike et al. 1991). As the grass dies, the insect s continue to move outwa rd to feed on moresucculent grass, thus enlarging the damaged area. 52

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Although capable of flight, adult move between lawns ma inly by walking and many have been observed crawling across paved areas bordering heavily infested lawns (Kerr 1966). All life stages are distribu ted vertically through the turf thatch and in to the upper organic layer of the soil, with densities of up to 2,000 /0.1 m2 being reported (Reinert and Kerr 1973). Light to moderate in festations are aggregated in small areas in the lawn, but can occur throughout the entir e lawn in heavily infest ed areas (Cherry 2001b). can be difficult to control because it has overcome host-plant resistance (Busey and Center 1987, Cherry and Nagata 199 7), it produces multiple generations per year, has a high number of offspring per generation, is highly mobile and disperses to neighboring lawns (i.e., encroachment), is able to surviv e on other grass sources until new St. Augustinegrass is located (Kerr 1966, Reinert and Kerr 1973), and is able to avoid insecticides. Currently, 20-25 per 0.09 m2 warrant control (Short et al. 1982). Insecticides are currently the only economical management option for lawn-care comp anies in Florida, with some making as many as twelve insecticide ap plications per year to control th is pest (Reinert 1978, Reinert and Niemczyk 1982). With near-constant reliance on chemical control, this insect has developed resistance to organochlorines, organophosphate s, and carbamates (Wolfenbarger 1953; Kerr and Robinson 1958; Kerr 1958, 1961; Reinert 1982a, 1982b; Reinert and Niemczyk 1982; Reinert and Portier 1983). In a 2003 University of Florida survey, the pyrethroid bifenthr in was the insecticide used most by lawn and ornamental professionals in Florida (Bu ss and Hodges 2006). Cherry and Nagata (2005) reported resi stance to bifenthrin in 14 populations in central and south Florida. In 2006, our lab received multiple complaints of field fail ures with bifenthrin and other pyrethroids as far north as Pe nsacola, FL. Additionally, pyreth roids are widely available to 53

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homeowners and professionals and their overuse m ay make pyrethroid resistance more widespread. In an effort to develop a resistance manageme nt program, it is important to determine where bifenthrin-resis tant populations occur in the st ate and the severity of the problem. Thus, I tested 16 populations in 2006 and 6 populations in 2008 in northern and central Florida to de scribe their susceptibility to bifenthrin, document new locations of resistance to bifenthrin, and ev aluate another pyrethroid, permethrin. Commercially-obtained plugs of Palmetto St. Augustinegrass were planted in 15.2-cm plastic pots filled with Farfard #2 potting soil (Conrad Farfard Inc., Agawam, MA). Plants were maintained in a University of Florida greenhous e in Gainesville, FL and held under a 14L:10D photoperiod with day and night temperatures of 27 and 24 C, respectively. Plants were fertilized weekly with a 20-20-20 water-solubl e complete nitrogen source (NH4NO3) at 0.11 kg N/0.09 m2, watered as needed, and cut to a height of ~7.6 cm. populations were collected between May and August 2006. Two populations were collected from areas where in secticides had not been used, three were randomly collected (treatment history unknown), a nd 11 were from lawns where control failures with bifenthrin had been reported (Table 2-1). The number of times lawns were treated prior to collection and the active ingred ients used during 2006 were documented for each site, where possible, and GPS coordinates were recorded. Se veral populations were collected from the same neighborhood or street, but were considered dist inct because their treatment history varied. Populations were named based on location within a neighborhood. 54

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populations were collected in Ju ly 2008. Six populations were from lawns where control failures with bifenthrin had been reported (Table 2-2). The active ingred ients used during 2008 were documented for each site, however, I was unable to obtain the number of times lawns were treated. GPS coordinates were recorded. Populations were named based on location within a neighborhood. Insects were collected using a modified Weed Eater Barracuda blower/vacuum (Electrolux Home Products, Augusta, GA) (Crocker 1993, Nagata and Cherry 1999, Congdon 2004), transported to the laboratory, si fted from debris, and fifth instar s and adults were placed into colony as outlined in Chapter3. Tests were conducted using a sprig-dip bioassay similar to that of Reinert and Portier (1983) and Cherry and Nagata ( 2005). Bioassays were run for 72 h because mortality results after 24 and 48 h for some of the populations did not fit a probit or logit model. This could have been due to a delay in response or because some insects initially avoided the plant material. Serial dilutions were made with formulated bifenthrin (Talstar One, FMC Corporation, Philadelphia, PA) and prepared fresh on each test date. Eight concentrations were tested and mortality ranged from 5 to 95% with the excep tion of three outliers, populations DAR, HF and GE18 (Table 2-1). Fresh Palmetto St. Augustinegrass stolon sections (5.0 6.4 cm long, with three leaflets and one node) were dipped in one solution and air dried on wax paper (~2 h). Ten unsexed adult of unknown age were placed into plastic petri dishes (100 15 mm) containing one treated stolon and one 70-mm Whatman filter paper moistened with 0.5-ml of 55

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distilled water to prevent desiccation. All tests were conducted between 1330 1500 h at room temp erature (25 2C) and a 14L:1 0D photoperiod. The number of dead was assessed at 24, 48, and 72 h using a dissecting microscope. Insects were scor ed as dead if they were on their backs or unable to walk. One population (JC) had control failu res with both TalstarOne and Permethrin-G Pro (permethrin, Gr o-Pro LLC, Inverness, FL), so both products were tested. Permethrin-G Pro solutions and testing were conducted as described with TalstarOne. Population HF was used as the susceptible standard. Tests were conducted using an ai rbrush bioassay as described in Chapter 4. A bifenthrinsusceptible laboratory population, LO (Chapter 4), was used as a st andard in this test. Serial dilutions were made with formulated bifenthr in (TalstarOne, FMC Co rporation, Philadelphia, PA) and prepared fresh on each test date. Eight or nine concentrations were tested for each population and mortality ra nged from 5-95%. Tests were held for 24 h and insects were scored as previously described. The LC50 and LC90 values, 95% confidence limits (CL), slopes of the regression lines, and the likelihood ratio test to test the hypothesis of parallelism and equality of the regression lines were estimated by logit analysis using Polo Plus (LeOra Software 2002). Differences in susceptibility between populations were tested by the 95% confidence limits (CL) of lethal concentration ratios (LCRs) at the LC50 and LC90 (Robertson and Priesler 1992, Robertson et al. 2007). Populations were individually compared to the most susceptible population (GE18) and LCR confidence intervals (95%) that did not include 1.0 were considered significant ( < 0.05) 56

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(Robertson and Priesler 1992, Robe rtson et al. 2007). Conventi onally, if the 95% confidence limits of the letha l concentrations overlapped, then the lethal concentrations were not considered significantly different. However, the ratio test has greater sta tistical power and lower Type I error rates, so this statistical test was used in this study (Wheel er et al. 2006, Robertson et al. 2007). The relationship between the number of insecticide applications made in 2006 and respective LCRs (at LC50) was analyzed using regression an alysis (Systat Software 2006). LC50 values for bifenthrin from the 16 populations (Table 2-3) were highest in populations that received two or more insecticide a pplications. Populations P, BH, and JC received the most insecticide applicat ions (8 11) and had the highest LC50 values for bifenthrin (3,835, 3,748 and 2,737 g/ml, respectively). Populati ons that received two to five insecticide applications (V, GE12, LF4, FS, BP, and CT) had LC50 values for bifenthrin ranging from 93 1,127 g/ml. Populations with one or no appl ications (DAL, DAR, HF, and GE18) had the lowest LC50 values, ranging from 0.9 42 g/ml. LCR50 values for all populations (with the exception of DAR and HF) were significantly di fferent from the most susceptible population, GE18, and increased with increasing insec ticide applications (Figure 2-1). LCR90 values for all populations treated with bifenthrin (with the exception of DAL and DAR) were significantly different from the most susceptible population, GE18 (Figure 2-2). The highest LCR90 values for bifenthrin we re recorded from populati ons BH, JC, GE12, LF4, L, FS, and BP. LCR90 values for these populations indi cated they were 1,077 13,000 g/ml more resistant to bifenthrin than the most susceptible population, GE18 (Figure 2-2). LC90 values for these same populations ranged from 53,120 to 642,527 g/ml. The lowest LC90 values were 57

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from populations GE18, DAR, and DAL (Tab le 2-3). Of the 11 populations collected, nine were actual cont rol failures (highest label rate of TalstarOne = 209 g/ml). Populations DAL and DAR demonstrated LC90 values that were below the recommended label rate, but control failure in these two sites may have been due to application error. Alternately, it is possible that different resistance mechanis ms are present in the DAL and DAR populations and the bioassay was unable to detect them. These data describe new locat ions of bifenthrin-resistant populations, as well as in counties similarly reported by Cherry and Nagata (2005) (Fi gure 2-3). In 2003, Cherry and Nagata (2005) reported 8 cases of resistance to bifenthrin in Flagler, Hernando, Lake, Manatee, Monroe, Sarasota, and Volusia counties, showing a 4.6 736 fold reduced susceptibility to bifenthrin. The data I coll ected in 2006 show a 45 to 4,099 fold reduced susceptibility to bifenthrin in Citrus, Escam bia, Flagler, Hillsborough, Orange, Osceola, and Volusia counties. These data are the first to report bifenthrin resist ance in Citrus, Hillsborough, Orange, and Osceola counties. In addition, pop ulation P from Escambia County is the first known in the Florida Panhandle to be resistant to insecticid es of any kind in (Figure 2-3). The results of the hypothesis tests of parallelism and equality show that the regression lines of 13 of the populations collected in central and northern Florida in 2006 were parallel but not equal to the mo st susceptible population, GE18 (T able 2-4). Even though their intercepts differ significantly, their slopes are not significantly different. This could mean that the field-collected populations are heterogeneous and represen t a range of susceptible and resistant individuals (as can be seen in population SCL with an LC50 of 47 and an LC90 of 4,039 g/ml). Alternately, the hypothesis test re sults may indicate that the different 58

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populations have qualitatively iden tical but quantitatively different levels of detoxification enzymes (Robertson et al. 2007). Population DAL, with a steep slope of 4.3, had significantly different intercepts and slopes from the GE18 popul ation. This may indicate that DAL was more uniform in its response to bifenthrin, their detoxification enzymes differ quali tatively, or that this population has entirely different detoxification enzymes (Robertson et al. 2007). Intercepts and slopes for populations DAR and GE18 were sim ilar, demonstrating a similar response to bifenthrin. It is interesting to note that the data obtained from the 2006 bifenthr in test indicate that individual lawns may represent a single population. In Palm Coast, sites GE12 and GE18 were located a few houses from each other, on the same side of the street, and were maintained by the same company at the time of this study. GE12 had received four insecticide applications between January and July 2006 and the collected from this lawn demonstrated an LC50 of 1,048 g/ml and an LC90 of 186,000 g/ml for bifenthrin. Meanwhile, lawn GE18 showed the presence of damage for the first time in 2006 and thus had not been treated at the time of collection. The collected from this lawn demonstrated an LC50 of 0.9 g/ml and an LC90 of 49 g/ml for bifenthrin. Also, population V was located in the same neighborhood, just one street away from GE12 and GE18. Although, the V population was under the same insecticide schedule as GE12, the collected from this lawn demonstrated an LC50 of 1,127 g/ml and an LC90 of 28,641 g/ml for bifenthrin. Populations FS and L, also located in Palm Coast but in a different neighborhood, were located directly across the street from each other, and were not maintained by the same lawn care company. The FS population received three inse cticide applications between January and July 2006 and the collected from this lawn demonstrated an LC50 of 652 g/ml and an LC90 of 53,120 59

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g/ml for bifenthrin, while the L population, with unknown treatm ent history had an LC50 of 521 g/ml and an LC90 of 62,612 g/ml for bifenthrin. Although, it is possible the sampled from these lawns did not fully represent the population as a whole, treatment effects on individual lawns, effects of encro achment, and population dynamics of within neighborhoods warrants further study. Based on the known treatment history for the populations where co ntrol failures with bifenthrin were reported in 2006, the number of applications ma de with bifenthrin, carbaryl, clothianidin, cypermethrin, imidacloprid, permethr in, and/or trichlorfon was positively correlated to their respective bifenthrin lethal concentration ratio (at LC50) values (Figure 2-4). While there are several documented cases showing a positiv e relationship between insecticide application frequency and selection for resistance (Geo rghiou 1986, Rosenheim and Hoy 1986, Croft et al. 1989, He et al. 2007, Magana et al. 2007), these st udies were based on knowledge of treatment history over a period of several years. Because I was only able to obtain the treatment history for 2006, it is uncertain whether application fre quency caused, or merely resulted from the development of resistance to bifenthrin in in this study. However, it is well documented in other organisms that resistance to pyrethroids often e volves quickly on the foundation of DDT resistance (Chadwick et al 1977, Prasittisuk and Busvine 1977, McDonald and Wood 1979, Omer et al. 1980, Priester a nd Georghiou 1980, Malcolm 1983, Miller et al. 1983, Georghiou 1986, Cochran 1995). Cases of DDT resistance in were documented in Sarasota (Kerr and Robinson 1958) and Miami (Kerr 1958), but, it is unclear how widespread the problem was and if cross resistan ce to pyrethroids is cu rrently occurring as a result. Due to the number of different insect icides used in 2006 to treat the populations I 60

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collected, cr oss resistance and/or multiple resistance may have occurred, but I did not have enough insects to test this. Population JC was 212.4-fold more tolerant of permethrin than the susceptible population, HF (Table 2-5). The hypothesis te st for equality was rejected ( 2 = 141; df = 2; < 0.05) and the hypothesis test for parallelism was not rejected ( 2 = 0.53; df = 1; = 0.47) showing that intercepts differed significantly, while slopes did not. Population JC from Orange County represents the first report of pe rmethrin resistance for the state. By 2007, Cherry and Nagata (2007) documented resistance to the pyrethroids deltamethrin a nd lambda-cyhalothrin, clearly indicating the occurrence of cross resistance in Florida. In addition, Cherry and Nagata (2007) documented the first case of resistance to a neon icotinoid, imidacloprid as well as finding six additional locations of bi fenthrin resistance. LC50 values for bifenthrin from the 6 populations collected in central Florida in 2008 ranged from 99 366 g/ml compared to the LC50 of 3.0 g/ml from the susceptible laboratory population, LO (Table 2-6). All 6 field-collected populations were actual control failures (highest label rate of TalstarOne = 209 g/ml), with LC90 values ranging from 293 1,439 g/ml (Table 2-6). Slopes of the regression lines from the populations tested were steep, indicating a uniform response to bifenthrin, w ith the exception of populat ion PA (Georghiou and Metcalf 1961; ffrench-Constant and Rous h 1990; Prabhaker et al. 1996, 2006). The regression lines of the 6 populations had significantly different intercepts from that of the most susceptible population LO (Table 2-7) The hypothesis test for parallelism was not rejected for populations JP, JH, and TG ( 2 = 0.5; df = 1; = 0.46, 2 = 1.5; df = 1; = 0.21, and 2 = 0.2; df = 1; = 0.66, respectively). For these populations, the slopes were similar to that of 61

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population LO. Populations LU, PA, and OR had significantly different intercepts and slopes from the LO population (Table 2-7). LCR50 values for all populations, ranging from 33 121 g/ml, were significantly different from the most susceptible population, LO (Figure 2-5). LCR90 values for these populations indicated they were 19 98 g/ml more resistant to bifenthrin than population, LO (Figure 2-6). The results of this chapter show that bifent hrin resistance continue s to spread and is particularly problematic in central Florida (Fi gure 2-3). Although, it is possible that pyrethroid resistance may be more widespread. In addi tion, these data along with reports by Cherry and Nagata (2007) show that cross resistance to other pyrethroids is occurring. Currently, pyrethroids, carbamates, neonicotinoi ds, and organophosphates are used for control in Florida. Carbamate (propoxu r) and organophosphate (chlorpyrif os) resistance was reported in the 1970s and 80s (Reinert and Niemczyk 1982, Rein ert and Portier 1983). Cross-resistance patterns and the stability of propoxur and chlorpyrifos resistance in are not known, making it unclear as to their effects on the cu rrent use of the carbamate, carbaryl, and the organophosphate, trichlorfon. In addition, the impact of insecticide us e on St. Augustinegrass grown in sod farms remains unknown. In a national study, Florida was ranked first in terms of economic impact of sod production (Haydu et al. 2006). In 2003, the tota l sod production in Florida was estimated to be 93,000 ha, with 64% being St. Augustinegrass (Haydu et al. 2005). Only 3% of harvested sod is sold outside of Florida. A summary of agri cultural pesticide use in Florida in 1995-1998 and 1999-2002 noted that chlorpyrifos was the sole insecticide used in sod farms (Shahane 1999, 2003). It is likely that St. Augustinegrass s od has already received several insecticide applications before it is even planted in residential neighborho ods. I have observed 62

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63 already present in St. Augustinegrass sod before it had been planted in a residential lawn. A better understanding of insecticide use on sod farms would be greatly benefi cial in understanding their role (if any) in selection for resistance to pesticides in populations in Florida. It is clear that further information is needed in order to solve the resistance problem in Florida. Once this is done, a resistance mana gement strategy can be made. An effective resistance management strategy should be multi-tactic (Hoy 1999) and include not only traditional integrated pest management (IPM) st rategies (monitoring pests, use of cultural controls, preservation of natural enemies, and hos t plant resistance) but al so include the possible use of synergists, effective educational programs, and monitoring of progr ess to ensure tactics are properly put into place. This would not only require cooperati on by professionals in academia and pest management, but should includ e sod growers, homeowner associations, and homeowners as well. In addition, solving the resistance problem in Florida will require much more research and cooperation and/or coordination with insect icide manufacturers. It is important to delay the development of resi stance to chemical classes available for control. Although doing so will be challenging be cause all registered products (carbamates, neonicotinoids, organophosphates, and pyrethroids) are currently used multiple times per year in attempts to control and prevent damage from in Florida lawns.

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Table 2-1. Collection sites a nd the number of insecticide applications made to the populations in Florida in 2006 that were tested for susceptibility to bifenthrin. Population County City GPS coordinates Month No. insecticide Active ingredients collected applications in 2006 usedc P Escambia Pensacola N30.70676, W87.7228 August 11 Bifenthrin Trichlorfon BH Citrus Beverly Hills N28.9644, W82.9684 August 11 Bifenthrin Carbaryl Imidacloprid Trichlorfon JC Orange Windermere N28.33244, W81.15464 June 8 Bifenthrin Permethrin Carbaryl Trichlorfon Acephate 64Va Flagler Palm Coast N29.81518, W81.87286 July 4 Bifenthrin Cypermethrin GE12a Flagler Palm Coast N29.78872, W81.93536 July 4 Bifenthrin Cypermethrin LF4 Flagler Palm Coast N29.69246, W81.93052 July 3 Bifenthrin Cypermethrin FSb Flagler Palm Coast N29.994833, W8110.11883 July 3 Bifenthrin Cypermethrin Lb Flagler Palm Coast N29.98482, W81.161 July unknown BP Hillsborough Sun City N27.516, W82.618 May 2 Bifenthrin CT Hillsborough Sun City N27.416167, W82.86733 June 5 Bifenthrin Carbaryl Imidacloprid PC Flagler Palm Coast N29.2641, W81.55944 May unknown

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Table 2-1. Continued SCL Osceola St. Cloud N28.20868, W81.0191 July unknown DAL Volusia Port Orange N29.101333, W81.952833 July 1 Bifenthrind Imidaclopridd DAR Volusia Port Orange N29.3879, W81.33 222 July 1 Clothianidine HF Alachua Gainesville N29.83908, W82.0241 JuneAugust 0 ------GE18a Flagler Palm Coast N29.78644, W81.93704 July 0 ------a Denotes populations in the same neighborhood. b Denotes populations located acro ss the street from each other. c Products are listed in descending order of application frequency.d A single application of Allectus SC was used at this site, which contai ns both bifenthrin and imidacloprid. e Control failure with bifenthrin was reporte d in 2005 but, at the time of collection, only clothianid in had been used in 2006. 65

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Table 2-2. Collection sites a nd the number of insecticide applications made to the populations in Florida in 2008 that were tested for susceptibility to bifenthrin. Population County City GPS coordinates Month No. insecticide Active ing redients collected applications in 2008* usedc LU Lake Clermont N28.8664, W814.9164 July N/A Bifenthrin Trichlorfon JP Orange Winter Garden N28.6611, W8138.9364 July N/A Bifenthrin Carbaryl Imidacloprid JH Orange Winter Garden N28.65, W81.5522 July N/A Bifenthrin Carbaryl Imidacloprid Fipronil PA Orange Windermere N28.0283, W8133.7480 July N/A Bifenthrin Carbaryl 66 Imidacloprid TG Orange Windermere N28.2447, W8134.6830 July N/A Bifenthrin Carbaryl Imidacloprid OR Orange Orlando N28.07361, W81.31778 July N/A Bifenthrin Carbaryl Imidacloprid I was unable to obtain the number of insecticid e applications that were made to these sites.

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Table 2-3. Response of Florida populations collected in 2006 to bifenthrin after 72 h using a sprig-dip bioassay at 25.5C, 14L:10D photoperiod. 67Population Slope SEa LC50 (95% CL)b LC90 (95% CL)b 2 (df)c P 240 2.0 0.3 3,835 (1,619,923) 44,798 (17,078,547) 5.1(5)d BH 80 1.0 0.2 3,748 (678,707) 642,527 (89,477,530,686) 4.0(5)d JC 240 1.1 0.1 2,737 (1,058,557) 260,786 (81,799,538,682) 3.2(5)d V 240 1.6 0.2 1,127 (490,358) 28,641 (11,496,575) 2.0(5)d GE12 240 1.0 0.1 1,048 (48,027) 186,000 (16,864,753,603) 14.1(5) LF4 240 1.1 0.2 817 (18,187) 71,506 (6,573,990,382) 18.2(5) FS 240 1.1 0.2 652 (32,649) 53,120 (8,599,135,424) 8.0(5)d L 240 1.1 0.2 521 (41,347) 62,612 (12,217,833,448) 5.9(5)d BP 240 0.9 0.1 459 (31,122) 143,891 (14,638,945,632) 11.0(5) CT 240 1.8 0.3 93 (10) 1,447 (310,491) 14.0(5) PC 240 2.0 0.4 62 (24) 785 (345,435) 0.6(5)d SCL 240 1.1 0.2 47 (1) 4,039 (661,205) 8.2(5)d DALL 240 4.3 1.1 42 (18) 137 (84) 0.1(5)d DAR 240 2.7 0.7 10 (3) 62 (32) 0.2(5)d HF 1,200 1.2 0.1 8 (2) 652 (349,436) 4.0(4)d GE18 240 1.3 0.4 0.9 (0) 49 (11) 0.5(5)d a Slope of the logit mortality line. b LC50, LC90, and 95% confidence limits (CL) are expressed in g/ml. c Pearson chi-square statis tic (degrees of freedom). d Good fit of the data to the logit model ( > 0.05).

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Table 2-4. Hypothesis tests compar ing the slopes and intercepts of logit regression lines for 15 populations in comparison to the most susceptible population, GE18, after exposure to bifenthrin for 72 h usi ng a sprig-dip bioassay at 25.5C, 14L:10D photoperiod. Population Hypothesis test Hypothesis test for equality for parallelism P reject; 2 = 168; df = 2; < 0.05 accept; 2 = 1.7; df = 1; = 0.19 BH reject; 2 = 74; df = 2; < 0.05 accept; 2 = 0.4; df = 1; = 0.56 JC reject; 2 = 126; df = 2; < 0.05 accept; 2 = 0.1; df = 1; = 0.71 V reject; 2 = 110; df = 2; < 0.05 accept; 2 = 0.3; df = 1; = 0.58 GE12 reject; 2 = 97; df = 2; < 0.05 accept; 2 = 0.47; df = 1; = 0.49 LF4 reject; 2 = 92; df = 2; < 0.05 accept; 2 = 0.10; df = 1; = 0.75 FS reject; 2 = 80; df = 2; < 0.05 accept; 2 = 0.07; df = 1; = 0.79 L reject; 2 = 78; df = 2; < 0.05 accept; 2 = 0.23; df = 1; = 0.63 BP reject; 2 = 78; df = 2; < 0.05 accept; 2 = 0.88; df = 1; = 0.35 CT reject; 2 = 44; df = 2; < 0.05 accept; 2 = 0.97; df = 1; = 0.32 PC reject; 2 = 33; df = 2; < 0.05 accept; 2 = 1.4; df = 1; = 0.24 SCL reject; 2 = 29; df = 2; < 0.05 accept; 2 = 0.08; df = 1; = 0.78 DALL reject; 2 = 35; df = 2; < 0.05 reject; 2 = 10; df = 1; = 0.001 DAR accept; 2 = 6; df = 2; = 0.06 accept; 2 = 3; df = 1; = 0.08 HF reject; 2 = 14; df = 2; = 0.001 accept; 2 = 0.04; df = 1; = 0.85 68

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Table 2-5. Response to perm ethrin after 72 h of two populations collected in 2006 using a sprig-dip bioa ssay at 25.5C, 14L:10D photoperiod. Population Slope SEa LC50 (95% CL)b LCR50 (95% CL)c LC90 (95% CL)b LCR90 (95% CL)c 2 (df)d JC 240 3.5 0.7 341 (130 750) 212 (104 434)* 1,431 (668 9,885) 157 (53.7 457)* 6.0 (5)f HFe 240 2.9 0.5 1.6 (1.0 2.7) 1 9.1 (4.9 28) 1 4.4 (5)f a Slope of the logit mortality line. b LC50, LC90, and 95% confidence limits (CL) are expressed in mg/ml. c Lethal concentration ratios w ith 95% confidence limits indicating the fold-difference for each population in comparison to the most susceptible population at LC50 and LC90. Confidence limits that include 1.0 indicate no significant difference from the susceptible (HF) population. Shows ratio s that are significant ( 0.05, Robertson and Preisler 1992; Robertson et al. 2007). d Pearson chi-square statis tic (degrees of freedom). e Susceptible population. f Good fit of the data to the logit model ( > 0.05). 69

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70 Table 2-6. Response of Florida populations collected in 200 8 to bifenthrin after 24 h using an airbrush bioassay at 25.5C, 14L:10D photoperiod. Population Slope SEa LC50 (95% CL)b LC90 (95% CL)b 2 (df)c LU 270 5.0 0.8 366 (291) 1014 (736) 3.2(6)d JP 54 4.1 1.3 333 (172) 1140 488,203) 3.6(5)d JH 288 4.0 0.5 129 (87) 457 (270) 10.4(6)d PA 288 2.1 0.3 124 (54) 1439 (382,909) 21.2(6)d TG 288 3.5 0.6 116 (68) 499 (277,370) 9.6(6) OR 288 4.7 0.6 99 (49) 293 (156,084) 25.1(6) LO 256 3.2 0.4 3.0 (1) 15 (8) 12.3(5)d a Slope of the logit mortality line. b LC50, LC90, and 95% confidence limits (CL) are expressed in g/ml. c Pearson chi-square statis tic (degrees of freedom). > 0.05).d Good fit of the data to the logit model (

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Table 2-7. Hypothesis tests compar ing the slopes and intercepts of logit regression lines for 6 populations in comparison to a sus ceptible laboratory colony, LO, after exposure to bifenthrin for 72 h using an airbrush bioassay at 25.5C, 14L:10D photoperiod. Population Hypothesis test Hypothesis test for equality for parallelism LU reject; 2 = 290; df = 2; < 0.05 reject; 2 = 4.5; df = 1; = 0.03 JP reject; 2 = 142; df = 2; < 0.05 accept; 2 = 0.5; df = 1; = 0.46 JH reject; 2 = 260; df = 2; < 0.05 accept; 2 = 1.54; df = 1; = 0.21 PA reject; 2 = 190; df = 2; < 0.05 reject; 2 = 5.25; df = 1; = 0.02 TG reject; 2 = 228; df = 2; < 0.05 accept; 2 = 0.20; df = 1; = 0.66 OR reject; 2 = 257; df = 2; = 0.001 reject; 2 = 5.0; df = 1; = 0.03 71

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72 L GE18 HF DAR DAL SCL PC P H CT B FS LF4 GE12 V JC B P 4000 3500 3000 2500 2000 1500 1000 500 0 * * * Figure 2-1. LC50 Lethal concentration ratios with lo wer 95% confidence limits indicating the fold difference for each population of in comparison to the most susceptible population (GE18) when tested with bifenthrin. Lower confidence limits that do not include 1.0 are significantly different from the GE18 population. Shows ratios that are significant ( 0.05, Robertson and Preisler 1992, Robertson et al. 2007).

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13,500 12,000 10,500 9,000 7,500 73 6,000 4,500 3,000 1,500 0 LF4 LF4 GE12 GE12 JC JC BH GE18 HF DAR DAL SCL CL PC PC CT CT BP BP L L FS FS V V P P * * * Figure 2-2. LC90 Lethal concentration ratios with lo wer 95% confidence limits indicating the fold difference for each population of in comparison to the most susceptible population (GE18) when tested with bifenthrin. Lower confidence limits that do not include 1.0 are significantly different from the GE18 population. S hows ratios that ar e significant (P 0.05, Robertson and Preisler 1992, Robertson et al. 2007). Figure 2-2. LC90 Lethal concentration ratios with lo wer 95% confidence limits indicating the fold difference for each population of in comparison to the most susceptible population (GE18) when tested with bifenthrin. Lower confidence limits that do not include 1.0 are significantly different from the GE18 population. S hows ratios that ar e significant (P 0.05, Robertson and Preisler 1992, Robertson et al. 2007).

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= Bifenthrin-resistant populations found by Cherry and Nagata in 2003 = Bifenthrin-resistant populations found by Vzquez in 2006 = Permethrin-resistant populations found by Vzquez in 2006 = Bifenthrinand deltame thrin-resistant populations found by Cherry and Nagata in 2007 = Imidacloprid-resistant populations found by Cherry and Nagata in 2007 = Lambda-cyhaloth rin-resistant population found by Cherry and Nagata in 2007 = Bifenthrin-resistant populations found by Vzquez in 2008 POL K HILLSBO ROUGH OR ANGE HERNA N DO OSCEOL A PASCO SANTA ROS A OKALOOSA ESCAMBIA WALTON LEVY GILCHRIST CIT R US ALACHU A COLUMBIA UNION B R ADFORD BA KE R N EE SUW ANHA MILTON DIXIE ETTE TAYLOR MADISON LAFAYJEFFERSON FRA N KLIN WAKULLA LEONLIBERTY GADSENGULF CALHOUN BAY WA SHINGTON JACKSON HOLMES N ASSAU DUVAL CLAY ST. JOHNS FLAGLER PUT N AM MARION VOLU S I A SUMTER LAKE SEMINOLE BREVARD PINELLAS INDIAN RIVEROKEECHO B EE HIGHLANDS MAN A TEE HARDEE ST. LUCIE SARASOTA DESOTO MART IN C HARLOTTE GLAD E S PALM BEACH HENDRY LEE BR OW ARD COL L IER MIAMI-DADE MONROE Figure 2-3. Map showing the dist ribution of insecticide-resistant populations in Florida between 2003-2008. The legend iden tifies counties where populations have been found and identifying authors. 74

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024681 01 2 0 1000 2000 3000 4000 5000 y = 346 slope = 379 r = 0.91 Figure 2-4. Bifenthrin resistance in populations from central and northern Florida in 2006: relationship between the number of inse cticide applications made (regardless of active ingredient used) a nd respective lethal conc entration ratios (at LC50) See Table 2-1 for locations sampled. 75

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140 130 120 110 100 90 80 70 60 50 LO OR TG PA JH JP LU * 40 76 30 20 10 0 Figure 2-5. LC50 Lethal concentration ratios with lo wer 95% confidence limits indicating the fold difference for each population of in comparison to the most susceptible population (LO) when tested with bifenthrin. Lower confidence limits that do not include 1.0 are significantly di fferent from the LO population. S hows ratios that are significant ( 0.05, Robertson and Preisler 1992, Robertson et al. 2007).

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77LO OR TG PA JH JP LU * 110 100 90 80 70 60 50 40 30 20 10 0 Figure 2-6. LC90 Lethal concentration ratios with lo wer 95% confidence limits indicating the fold difference for each population of in comparison to the most susceptible population (LO) when tested with bifenthrin. Lower confidence limits that do not include 1.0 are significantly di fferent from the LO population. S hows ratios that are significant ( 0.05, Robertson and Preisler 1992, Robertson et al. 2007).

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CHAPTER 3 SYNCHR ONOUS METHOD FOR REARING ON CORN AND ST. AUGUSTINEGRASS The southern chinch bug, Barber, is the most destru ctive insect pest of St. Augustinegrass (Reinert and Kerr 1973, Brut on et al. 1983). Similar to other feeding habits, nymphal and adult damage St. Augustinegrass by f eeding in the phloem sieve elements of the grass (Rangasamy et al. 2009) causing wilting, chlo rosis, stunting, and eventually death (Painter 1928, Negron and Rile y 1990, Spike et al. 1991). Populations may consist mostly of long-winged forms (macropter ous), short-winged forms (brachypterous), or both (Wilson 1929, Komblas 1962, Leonard 1966, Reinert and Kerr 1973). In Florida, macroptery is greatest during the summer and fa ll when populations are high (Cherry 2001a). Eggs are laid singly or a few at a time in leaf sheat hs, soft soil, or in other protected areas. Eden and Self (1960) reported that eggs hatch in 14 d in Mobile, AL, while Kelsheimer and Kerr (1957) state eggs can hatch in 7-10 d during the summer in Florida. Young nymphs are as small as 0.87 mm (Leonard 1968), are reddish-ora nge with a white band across the dorsal side of the abdomen, and become black in color as th ey mature. Development from egg to adult can vary depending on location and temperature (Swe et 2000): 35 d in Florida (Kelsheimer and Kerr 1957), 49-56 d in Alabama (Eden and Self 1960) and 30-45 d in Mississippi (Burton and Hutchins 1958). Kerr (1966) reported that development from egg to adult is completed in 93 d at 21C and in 35 d at 28C. In addition, female and male longevity was 70.4 and 42.1 d, respectively, with females laying an average of 4.5 eggs per day under laboratory conditions (Kerr 1966). Control for is mainly achieved through insecticide use. Because is multivoltine and has overlapping generati ons (Kerr 1966, Reinert and Kerr 1973), damaging 78

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populations have received as many a s 6 to 12 insect icide applications a year in Florida (Reinert 1978, Reinert and Niemczyk 1982). With n ear-constant insecticide exposure, has developed resistance to organoc hlorines, organophosphates, ca rbamates, neonicotinoids, and pyrethroids (Kerr 1958, 1961; Re inert 1982a, 1982b; Reinert and Niemczyk 1982; Reinert and Portier 1983; Cherry and Nagata 2005, 2007; Chap ter 2). To conduct in secticide-resistance studies, it is important to c onduct tests with quality insects of known age and generation (ffrench-Constant and Roush 1990) Therefore synchronized rear ing methods are needed for Several attempts have been made to rear spp. under laboratory or greenhouse conditions (Parker and Randolph 1972, Baker et al. 1981, Yamada et al. 1984, Wilde et al. 1987, Meehan and Wilde 1989, Anderson 2004). However, laboratory-reared often incurred high mortality, overlapping generations were pr oduced so that insect age could not be determined prior to bioassays, or percentage success in developm ent from egg to adult was not reported. Several authors successf ully produced > 1 generation of spp. under greenhouse and growth-chamber conditions (Wilde et al 1987, Meehan and Wilde 1989, Anderson 2004). However, mass rearing in the greenhouse at the Entomology and Nematology Department at the University of Florida has not been feasible becau se daily ambient summer temperatures in the greenhouses can exceed 37.8 C, which is lethal for Also, potted St. Augustinegrass can become infested with aphids, thrips, scales, mites, other populations, and natural enemies. In 2004 and 2005, preliminary tests were conduc ted to evaluate appropriate rearing containers, food sources, and oviposition substr ates to minimize handling while maximizing growth and speed of development. Differ ent food sources tried included Palmetto St. 79

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Augustinegrass (both stolon sections and ground plant material m ixe d with plain gelatin), green beans ( .); ryegrass ( .); frozen peas ( ); and baby, canned, fresh, frozen, and organic seedling corn ( .). attempted to feed on the green beans but were unable to insert their stylets into intact beans. Of all the food choices, performed best on fresh St. Augustinegrass and corn on the cob. For the remaining food choices, either did not feed or mold developed on food in 24 h. Ovipositional substrates tried included 70-mm Whatman filter paper, colored felt cloth (black, white, and green) (Michaels Stores, Inc ., Irving, TX), colored cardstock (black, white, and green) (Michaels Stores, Inc., Irving, TX), colored foam sheets (black, white, green) (Michaels Stores, Inc., Irving, TX), paper towe ls (brown and white), cheesecloth, moistened cottonballs, cotton diaper towel (C lean-Rite Products, Lincolnshire IL), and moistened Publix brand cosmetic squares (Publix Super Markets, Inc., Lakeland, FL). either didnt lay eggs on the substrates (felt, cardstock, foam, paper towels, cheesecloth) or nymphs became trapped and were unable to free themselves (moistened cottonballs and cosmetic squares). However, nymphs were able to emerge without becoming trapped in the diaper towel. To control moisture in the rearing chambe rs, Feline-Pine cat litter (Natures Earth Products, Inc., West Palm Beach, FL), compressed sponges (The Color Wheel Company, Philomath, OR), Plaster of Paris (Lowes Companie s, Inc., Mooresville, NC), and dental castone (Henry Schein Inc., Indianapolis IN) were tried. Various sized plastic containers, petri dishes, cardboard containers, glass jars and 15.2-cm potted Palmetto St. Augustinegrass with 15.2-cm stolon sections enclosed in fibe r floral sleeves (Temkin International, Inc., Miami, FL) were 80

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evaluated. The Feline-Pine cat litter and compressed sponge s worked well at controlling excess moisture, but were suitable for shelter of and as an oviposition substrate. This made it difficult to remove insects and to co unt eggs to determine percentage survival. The plaster of Paris was difficult to work with becau se it dried before being placed into the rearing containers. However, the dental castone stayed in a semi-liquid form long enough to work with. Plastic containers and glass jars proved to be the best for housing because appropriate relative humidity could be maintained. Other materials allowed too much air flow so that eggs desiccated, resulted in excess moistu re, or were not big enough to contain large numbers of insects. Thus, fresh corn cobs and St. Augustinegrass, cotton diaper towel, dental castone, plastic containers, and glass jars were chosen as can didate materials for rearing Several tests were then conducted be tween 2005 and 2008 to develop a synchronized rearing method to produce pure populations of known age and generation for use in insecticide-resistance studies. Commercially-obtained plugs of Palmetto St. Augustinegrass were planted in 15.2-cm plastic pots filled with Farfard #2 potting soil (Conrad Farfard Inc., Agawam, MA). Plants were maintained in a greenhouse near the Univer sity of Florida Entomology and Nematology Department in Gainesville, FL, and held under a 14L:10D photoperiod with day and night temperatures of 27 and 24C, respectively. Heating mats were used to keep plants from going dormant during the winter months. Because of the slightly acidic (pH=5.5) greenhouse water, all plants were provided with 32 ml of a hydrated lime solution on a weekly basis (Oldcastle Stone Products, Thomasville, PA) (10.6 g/3785 ml) to enhance nutrient uptake. Plants were fertilized 81

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weekly with 20-20-20 (N-P-K) wate r-soluble fertilizer (United Indus tries, St. Louis, MO) at a rate of 0.11 kg N/0.09 m2, watered as needed, and cut to a height of ~ 7.6 cm. Commercially obtained bushels of yellow corn cob were shucked, soaked in a 3% bleach solution (600 mL of 6% sodium hypochlorite in 1.94 L tap water) for 10 min, and rinsed. Corn cobs were then dried and stored in 7.6-L Ziplock clear plas tic bags (S. C. Johnson and Son, Inc., Racine, WI) containing four sheets of B ounty paper towels (Proctor and Gamble Co., Cincinnati, OH) to absorb excess moisture, and refrigerated. were collected from St. Augustineg rass lawns using a modified Weed Eater Barracuda blower/vacuum (Electrol ux Home Products, Augusta, GA) (Crocker 1993, Nagata and Cherry 1999, Congdon 2004) and tran sported in a mesh-covered bucket to the laboratory. Adults were aspirate d from debris and placed into oviposition containers, as outlined below. Plastic containers (15.2-cm diam eter, 6.4-cm deep) were soaked in a solution containing 1 L of 6% bottled bleach and 19 L tap water for 20 min, rinsed, and allowed to dry. Fluon (Ag Fluoropolymers, Chadds Ford, PA) was applied to th e top 3 cm of the contai ner to prevent insect escape. Castone dental stone Type III (DentSpl y Inc., York, PA) was then poured to a 0.5-cm depth in each container and allo wed to dry for 24 h. Four holes (2-cm diameter) were cut into the plastic lids and chiffon mesh was glued over the holes to allow airflow. For nymph containers, one hole (1.6-cm diameter) was cut 1.3 cm from the bottom of the container for placement of a 7.6-cm long water tube. A small hole (0.28-cm diameter) was drilled into the water tube so water could be adde d without disturbi ng the insects. 82

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Cotton diaper towels were used to create egg rolls for collecting eggs. Egg rolls were created by cutting diap er towels into 7.6-cm 7.6-cm sections and rolled to a 1.7-cm diameter by wrapping around a plasti c 10-mL graduated cylinder. Thirty unsexed were placed into each of three oviposition containers and provided with one corn section (5 2.5 cm) and two egg rolls as an ovipo sition substrate (Figure 3-1 A). The egg rolls were collected daily for 4 d, eggs were counted, and placed into nymph containers containing one 15.2-cm long stolon of Palmetto St. Augustinegrass with the excised end inserted into a floral tube filled with wate r (Figure 3-1 B). There were 12 replicates. All oviposition and nymph containers were maintained in the laboratory under a 14L:10D photoperiod, 26-31 C and 60-70% RH. Stolons were cha nged weekly and tubes filled with water as needed. Dental castone was moistened with 2.5 ml of deionized water 5 days a week to maintain relative humidity. The number and percentage of adult that successfully developed, sex ratio, and the ratio of wing t ype were determined for each replicate. A subsequent test was designed to determine if the previous method would be appropriate for larger-scale rearing and to determine the time of day oviposition was highest. Oviposition and nymph containers were set up and maintained as previously described. The egg rolls were collected three ti mes a day (0700, 1500, and 2300 h) from two oviposition containers (Figure 3-1A ), eggs were counted, and tran sferred into nymph containers. There were 20 replicates for each 8-h interval. Nymph containers (Figure 3-1B) were checked daily until the first sign of adult emergence, then were checked twice a day (0800 and 1600 h), and newly emerged adults were counted and stored in vials of 95% EtOH. The number of eggs collected at each time interval, number of days for chinch bug development from egg to adult, 83

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ratio of males to fem ales, ratio of brachypterus and macropterus adults, an d percentage survival were determined. An analysis of variance (ANO VA) was used to determine if the mean number of eggs collected differed among time intervals. Treatment means were analyzed using the Tukeys Studentized Range (HSD) test (SAS Institute, Inc. 2001). The purpose of this test was to reduce competition for food, which was observed in Test 2. To accomplish this, 7.6-L glass jars (Heritage Hi ll Collection, Anchor Hocking, Lancaster, OH) were used in place of plastic containers to house more and food. The jars were washed, dried, and a 5-cm band of Fluon was applie d with a wash bottle to the inside top of the jars, and dried using a hairdryer. This created a barrier to preven t insects from escaping. At the bottom of each oviposition jar, dental castone was poured to a 1.3-cm depth and allowed to dry for 24 h. The corn was prepared as previously desc ribed. For placement into colony jars, the bottoms of 59-ml plastic souffl solo cups (G ainesville Paper Supply, Gainesville, FL) were removed, the cups cut in half, and then inserted into the bottom of each cob to keep it from rolling. Excess moisture from around the cup sta nds was blotted with paper towels to reduce the development of mold. were collected as previously desc ribed. Adults were aspirated from debris and placed into jars containing two full-sized, surface-sterilized, fresh yellow corn cobs and 12 egg rolls provided as an oviposition substrate (Figur e 3-2A and B). Corn cobs and stands were replaced twice a week in jars containing adult An air stream (~ 1.4 m/s) was gently applied to the corn cobs to cause the insects to stop feeding and 84

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fall to the bottom of the jars unharm ed. Dead insects or eggs lying at the bottom of the jars were vacuum aspirated before adding new corn cobs an d stands. Excrement that was deposited on the inner sides of the jars was wiped clean using Clorox disi nfecting wipes. For nymph development, one Palmetto St. A ugustinegrass (15.2-cm diameter) plant was planted in jars containing 1500 g of sterilized build ers sand (Figure 3-3) the week of egg introduction. Plants were watered as needed. Oviposition and nymph jars were enclosed with chiffon mesh after insects were introduced. All jars were main tained in the laboratory under a 14L:10D photoperiod, 26-31 C, and 70-85% RH. The egg rolls were collected once a week for 3.5 wk from six oviposition jars, eggs counted, a nd transferred into nymph jars. There were 21 replicates. The number a nd percentage of adult that successfully emerged were determined for each replicate. This experiment was designed to determine if could be solely reared on fresh corn on the cob. The oviposition jars (Figure 3-2 A) and castone for the nymph jars were constructed as described for the oviposition jars in Test 3. The egg rolls were collected weekly for 7 wk from one oviposition jar and eggs present were counted (total: 7,092 eggs). Twelve egg rolls (Figure 3-2 B) with eggs were transferred into each nymph jar containing one fresh surface-sterilized corn cob placed on two plastic stands. One fresh corn cob was added every 3 4 d and st acked alternately over the old corn cobs. Old corn cobs were not removed because nymphs used them as shelter and were difficult to remove. To maintain high humidity, 50 ml of water was added to the dental cas tone each week. All oviposition and nymph jars were maintained in the laboratory under a 14L:10D photoperiod, 2685

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31C, and 70-85% RH. The number and percentage survival of was dete rmined after 8 wk. An improved method of rearing adults on corn and nymphs on grass in 7.6-L glass jars was created based on the results of Tests 1-4. The following methods were used to determine the success of the rearing method. Clean jars had fluon applied as previously described. Jars were se t over wax paper and a marker was used to trace around the bottom of the jar to create wax pape r circles. The paper circles were then cut in half, taped to a 23 cm 2 cm cardboard strip, and placed at the bottom of the jars (Figure 3-4A). Castone (453 g mixed in 200 mL water) was poured over the wax paper/cardboard assemblage, carefully making sure to cover any holes in the cardboard (Figure 3-4B). The castone was allowed to dry for 24 h. The egg rolls were created as described in Te st 1 and were replaced in oviposition jars every 2 d. Any adults on the egg rolls were care fully removed and egg rolls were placed into a separate jar containing castone. One air-dried, soil-free, 15.2-cm St. Augustinegrass plant was placed next to or on top of the egg rolls and jars were covered with chiffon nylon mesh held in place with a rubber band (Figure 3-5). A clear plastic shower cap was added to maintain RH>70%. One end of a pair of sc issors was used to pierce the s hower cap, creating five holes in a star design to allow ventilation. One fresh St. Augustinegrass plant wa s added twice per week to each jar. The older plant mate rial was left in the jars so nymphs could use it for shelter and molting. Nymphs were often distribute d or concentrated at the bottom of the jar, 86

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perhaps because of the higher RH near the castone bottom. As a result, new plant m aterial was placed underneath or directly next to older plant material. To determine the quality of the insects a nd success rate of this rearing method, a colony was reared for eight generations (Figure 3-2 A, 34, 3-5) before testing. Egg rolls containing ninth-ge neration eggs were removed as desc ribed and 400 eggs were counted, placed into a castoned jar, and held in a Percival growth chamber (model I36VLC8) under a constant temperature of 30 C and 14L:10D photoperiod. There were eight replicates for a total of eight jars and 3,200 eggs. Nymphs that emerged were maintained as prev iously described. Three days after the first sign of adult emergence, all live insects were removed, counted, and place d into vials containing 95% EtOH. All were then sexed and wing-typed using a dissecting microscope. Thirty brachypterus females were randomly select ed from each jar, mounted onto cardstock, and body length measured. The percentage survival wing length, and body length (of brachypterus females) were used to determine environmen tal stresses (crowding) and food quality. Body length comparisons were made of offspring, pa rent, and field-collected (merged from four populations in central FL) brachypterus females to determine the relative fitness of the colony after being reared for eight generations. Analyses of variance (ANOVA) were used to determine mean differences in body lengt h of emerged brachypterus females by replicate and between offspring, parent, and fi eld-collected brachypterus female Treatment means were analyzed using the Tukeys Studentized Range (HSD) test (SAS Institute, Inc. 2001). 87

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The percentage survival of that successfully developed from egg to adult in each replicate ranged from 61.1-100% (Table 3-1). The total average percentage survival from all replicates was 85%. Th e sex ratio was 1:1 and all that emerged were brachypterus. However, because few eggs (1-22) we re placed into each c ontainer, a larger-scale study was needed to determine if this me thod could be used for mass rearing of and to determine the best time of day to collect eggs. The greatest number of eggs was collected at 2300 h and the temperature was highest between 1500 and 2300 h (Table 3-2). This suggests that the best time to collect eggs from containers would be in the morning to minimize interfering with oviposition. Females took slightly longer to develop than males, 42.5 0.5 and 40.2 0.5 d, respectively. The ratio of males to females was 1:1 and the ra tio of brachypterus and macropterus adults that emerged was 7:1. Out of the 976 eggs collected, only 381 (39%) survived to the adult stage. It is possible that survival decreased with increa sing egg density. In the containers with egg densities of 41 and higher, we had problems w ith chinch bug nymphs squeezing through the cap into the water tube and drowning, suggesting there was competition for food. Also, it is possible that nymphs may have been harmed or accidentally discarded when grass material was changed. We speculated that larger rear ing containers, additional food, a nd leaving the grass material inside the chambers until the test is completed may increase the percentage survival. The 7.6-L glass oviposition jars worked well for obtaining large numbers of eggs and were more suitable for housing more insects than the containers used in Tests 1 and 2. 88

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However, of the 30,534 eggs collected and placed into nymph jars, only 1,011 (3.3%) survived to the adult stage (Table 3-3). Plants used in th is test m ay not have been washed thoroughly, so predators remained hidden in the plants. nymphs were not apparent after the first week and upon greater inspec tion, spiders were seen toward the top of some nymphal jars, in the egg rolls, or fell out as th e plants were pulled apart. Spiders were present in all of the nymphal chambers and in one case each, small earwigs, centipedes, and one Say was found. Anderson (2004) also had pr oblems with predator s invading greenhouse populations of spp., resulting in difficulty in rearing multiple generations. Grass in the jars containing >1000 nymphs (Table 3-3) quickly wilted and died within the first 2 wk of the te st, indicating that the 15.2-cm gr ass plant was not enough food to sustain the high number of and allow them to complete development to the adult stage. While the nymphal chambers in this te st were inadequate for use in mass rearing of the ovipositional chambers were superior to those used in tests one and two. The ovipositional chambers in this study allowed for more to be housed and thus obtain more eggs. nymphs could not complete development to the adult stage when reared solely on fresh corn cob. Out of 7,092 eggs, onl y 12 (0.17%) survived to the second instar after 2 months of being maintained on corn (Table 34). In addition, castone was difficult to remove in some of the jars because it would stick to the bottom. There are several factors that may influence an organisms chance to survive and multiply: 1) food, 2) weather, 3) other organisms, and 4) a place in which to live (Andrewartha 1965). The host plan t used in rearing may affect one or more of these categories, directly or indi rectly (Berlinger 1992). It is possible that the fresh corn cob did not meet the nutritional needs for for growth and development. 89

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The absence or imbalance of certain nutriti onal requireme nts may have prevented growth (Chapman 1969) or impaired the ability of to molt. In addition, host plant morphology may also af fect the insects microclimate (Berlinger 1992). Nymphs of tend to settle or hide in narrow places such as inside the sheaths of St. Augustinegrass. This allows them to be in maximum contact with their surroundings (thigmotropism). It is possible that it allows them to be in close c ontact with food and/or provides protection from environm ental factors or predators. In order to rear thigmotrophic insects (such as ) effectively, it is important to fulfill this requirement (Berlinger 1992). By eliminating the St. Augustinegrass from the nymphal diet and replacing it with fresh corn, this may have eliminated their place to live. This may have exposed them to the direct light in the reari ng room or stressed them, causing death or slowed growth. All adult generation-nine emerged after 5.5 wk when reared at a constant temperature of 30 0.1 C and 14L:10D photoperiod. Percenta ge survival of successfully emerged in each jar ranged from 56 79% (Table 3-5). Th e sex ratio in each jar was 1:1 with a 5:1 ratio of brachypterus to macropter us individuals. The body length of brachypterus females from each jar ranged from 3.67 0.1 to 3.88 0.2 mm and significant differences were observed (Table 3-1). Particularly, jars 5, 7, and 8 had brachypterus fema les with significantly longer body lengths than females fr om other jars. Jar 1 produced brachypterus females with the shortest body length. Mean body length for brachypterus female of fspring was similar to that of the 30 randomly chosen brachypterus female collected from the parent colony (3.77 0.01 and 3.83 0.03 mm, respectively, for mean body length; = 3.47; df = 1, 268; = 0.06). 90

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However, both parent and offs pring of laboratory-reared were significantly larger than field-co llected (3.32 0.02 mm for mean body length for field-collected ; = 187; df = 2, 323; < 0.0001). The size of brachypterus females that emerged in Test 5 indicates that a combination of fresh corn cob and St. Augustinegrass provides a suitable diet for rearing The St. Augustinegrass used in our study received a weekly nitrogen (N) supply at the highest recommended rate for Florida lawns (0.11 kg N/0.09 m2 per wk). Busey and Snyder (1993) suggested that greater host plant quality is associated with fa ster development, greater survival, and higher fecundity in Likewise, an increase in N fertilization is usually followed by an increase in populations in other phytophagous arthropods (Harre wijn 1970, Wermelinger et al. 1985, Berlinger 1992). Realized fecundity is positively related to female body size in some insect species (Speight 1994, Preziosi et al. 1996, Tammaru et al. 1996, Sopow and Quiring 1998). In addition, Dahms (1947) conducted studies with sorghum and found that higher rates of nitrogen (N) fertilization were associated with increased oviposition rates. It is possible that the laboratory-reared in this study were provi ded with higher quality turfgrass than is found in residential lawns in Florida. This would explain why the offspring from Test 5 were larger than the insect s that were collected from residential lawns in central Florida. Although larger than the field-collected specimens, brachypterus female offspring in Test 5 had body lengths comparable to those found by Cherry and Wilson (2003) in Florida, which had body lengths ranging from 3.2-4.0 mm. Time for development (~35 d) was sim ilar to that of field populations of in Florida (Kelsheimer and Kerr 1957) at 28C. This would suggest that by using Test 5 methods, laboratory-reared can be produced that are of high quality and comparable in 91

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development times to field-collected in Florida. This may be important if laboratory-reared colonies are used for developing baseline insect icide studies in place of fieldcollected insects of unknown age and quality. The large number of male and female brachypterus that emerged in Test 5 showed that environmental stress (e.g., climate, crowding, competition for food) was minimal after nine generations. Cherry and W ilson (2003) reported brachypterus female contain more eggs, as determined by dissection, a nd laid significantly more eggs per female than macropterus females. Fujisaki (1985) noted more macropterus Okajima (Heteroptera: Blissidae) individuals emer ged when placed under extremely crowded conditions. The number of macropterus that emerge in colonies could be used as an index for quality control in laboratory rearing. This work presents the first successful synchronized rearing method for Other spp. have been reared on gr ass under laboratory conditions, but with minimal success. Yamada et al. (1984) reared the oriental chinch bug, Okajima, on maize, Kentucky bluegrass, sorghum, and sugarcane. However, only 40% of insects in the second generation successfully surv ived to the adult stage. Pa rker and Randolph (1972) reared the common chinch bug, (Say), in 3.78-L car dboard cartons on alternating stacked layers of maize and sorghum stalk sections. Each carton could produce 8001000 chinch bugs (Parker and Randolph 1972). Ho wever, this method produced overlapping generations in each container and the authors di d not report how many generations were reared. Baker et al. (1981) attempted to rear the hairy chinch bug, Montandon, using Parker and Randolphs (1972) techniqu e, but early-instar mortality was high, which appeared to be associated with fungal grow th on the corn sections. The authors increased 92

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survival by surface-sterilizing corn sec tions with 2% sodium hypochlorite and frequently changing corn sections in containers. However, the authors still had an average of 17% mortality in the egg, 48.3% during the first and second instars, and 15.7% during the third through fifth instar stages. On ly 80% of third-fifth instar successfully developed to the adult stage (Baker et al. 1981). The total per centage survival from e gg to adult in their study was 20%. In this study, a combin ed total average of 67% of successfully emerged from the rearing containers. However, four repl icates had percentage su rvival rates ranging from 71-79%. Grass plants in nymphal jars sitting on the top shelf of the growth chamber dried faster than in jars located on lower shel ves, possibly because the fans were directly above the top shelf and increased air flow. Excess ai r flow may have dried out eggs or reduced the food supply to early-instar Using shower caps with out ventilation or placin g the upper shelf farther from the fans appeared to increase survival. Ho wever, regardless of reduced survival in half of the jars, the total percentage survival in in this study is the highest reported in any laboratory-rearing procedure published for spp. Problems I encountered while rearing previous generations included contamination with predators, egg parasitoids, mold/fungal growth, and safety issues regarding removal of dental castone (ie., shards) from colony jars. Contamination with predat ors and egg parasitoids can be greatly reduced by carefully vacuuming from field samples (not introducing other debris into colony jars) and thoroughly washing plan t material before placing it into nymph jars. Mold was greatly reduced on corn cobs after surf ace sterilization and rem oval of excess moisture before placing the corn into jars. Another means of reducing mold was by eliminating the addition of water to the dental castone. The addition of wate r is needed when using the containers employed in Tests 1 a nd 2. The depth of the castone used in these containers was 93

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small and so water readily evaporated from it. However, the thicker dent al castone in the 7.6-L glass jars d idnt require the additional water an d helped maintain high humidity. Also, the thicker dental castone in the glass jars absorbed excess moisture from grass in nymphal jars, which greatly reduced problems with mold. Overa ll, sanitation was found to be crucial to the success of rearing in the laboratory. Safety also was of concern because of the diffi culty in removing dental castone from jars. Dental castone can be difficult to remove without the use of the wax paper/cardboard assembly, but with this in place, jars can be set upside down and the dental castone will drop down and then can be folded in half and safely removed. If the dental castone doesnt fall the cardboard strip in the center can be pulled out to remove the dental castone so jars can be cleaned and reused. In terms of labor for Test 5 methods, it takes ~5.5 h per week to set up one oviposition jar and produce one jar of offspring (Figure 3-6). Not including supplies, the labor cost ($10 per hour) would be $55 so, if one jar of offspring produces 224-317 then the cost per insect is $0.17-0.24. For the first time, we can produce large numbers of of known age and generation for use in bioassays that are of hi gh quality in comparison to the field-collected which could be stressed from previous treatm ents with pesticides or because of a poorquality diet. Using reared insect s will help to reduce variability in insecticide bioassays and can be used to develop insecticide-susceptible colonies for use as a baseline in bioassays. This work also provides a key step in buildi ng a resistance-management program for colonies can now be selected for bifenthrin resistance (or any other insecticide) in the laboratory. Pure insecticide-sus ceptible and -resistant colonies can then be used in tests to 94

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determine mechanisms, cross-resistan ce, mode of inheritance, and stab ility of resistance, providing key information regarding the genetics of resistance in 95

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Table 3-1. The total number eggs in each replic ate at the s tart of Test 1* and the number of female and male that successfully emerged af ter 5.5 wk that were reared at 26-31C, 60-70% RH, and a 14L:10D photoperiod. Total number eggs Number adults emerged (: ) Percentage survival from egg to adult Replicate 1 1 1 (1:0) 100.0 2 13 12 (7:5) 92.3 3 6 5 (2:3) 83.3 4 3 3 (2:1) 100.0 5 10 7 (3:4) 70.0 6 9 8 (4:5) 88.9 7 6 6 (3:3) 100.0 8 18 11 (6:5) 61.1 9 6 6 (3:3) 100.0 10 11 8 (3:5) 72.7 11 22 22 (10:12) 100.0 12 21 18 (8:9) 85.7 Small-scale rearing of adults on corn and nymphs on grass. 96

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97 Table 3-2. Mean number ( SEM) of eggs collected at each 8-h interval in Test 2 and respective average temperature (C) and % RH ( SEM). Collection Mean number Average air Average Time (h) eggs SEM temp SEM (C) % RH SEM 0700 3.0 0.7 26.8 0.3 65.0 2.2 1500 16.5 3.5 31.2 0.5* 60.5 2.2 2300 29.2 5.1* 29.9 0.5* 65.6 2.9 Mean SEM within a column followed by are significantly different ( < 0.05) by TukeyKramer HSD test ( = 13.2; df = 2, 57; < 0.0001 for mean number of eggs; = 24.2; df = 2, 57; < 0.0001 for temperature; and = 1.32; df = 2, 57; = 0.27 for % RH). Assessment of time of day for oviposition.

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Table 3-3. The total number eggs in each replicat e at the s tart of Test 3* and the number of adults that successfully emerged afte r 6 wk that were reared at 26C, 6070% RH, and a 14L:10D photoperiod. Replicate Total no. eggs in jar Num ber adults emerged Percentage survival from egg to adult 1 620 33 5.3 2 1113 48 4.3 3 2484 119 4.8 4 1448 137 9.5 5 532 15 2.8 6 759 17 2.2 7 1897 101 5.3 8 2977 110 3.7 9 581 56 9.6 10 909 0 0 11 637 0 0 12 930 0 0 13 605 20 3.3 14 572 27 4.7 15 1842 91 4.9 16 1336 129 9.6 17 1312 73 5.6 18 2732 30 1.1 19 459 5 1.1 20 395 0 0 21 6394 0 0 Rearing nymphs on planted grass in builders sand and glass jars. 98

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Table 3-4. The total number eggs in each replicat e at the s tart of test four* and the number and stage of found after 8 wks that were reared at 26-31C, 70-85% RH, and a 14L:10D photoperiod. Total number of eggs in jar Num ber found alive Stage of development Replicate 1 1497 0 2 1215 0 3 953 8 2nd instar 4 1383 4 2nd instar 5 996 0 6 530 0 7 518 0 Corn only rearing method. 99

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Table 3-5. The number of em erged generation-nine adults, percentage survival, wing type, and comparison of mean ( SEM) body length (mm) of brachypterus females by replicate for test five that were reared at a constant 30C, 75 5% RH, and a 14L:10D photoperiod, using fresh corn co b, St. Augustinegrass, and glass jars. Jar Number adults Percentage Number of Body length in mm emerged : survival brachypterus:m acropterus ( SEM)* 1 258 (141:117) 64.0 218: 40 3.67 0.02ad 2 236 (121:115) 59.0 201: 35 3.76 0.03ac 3 296 (146:150) 74.0 249: 47 3.75 0.02ac 4 288 (150:138) 72.0 245: 43 3.74 0.02ac 5 317 (174:143) 79.2 253: 64 3.88 0.03bc 6 283 (147:136) 70.7 247: 36 3.75 0.02ac 7 224 (118:106) 56.0 198: 26 3.79 0.02bc 8 237 (136:101) 59.2 204: 33 3.84 0.02bc Mean SEM within a column followed by the same letter are not significantly different ( < 0.05) by Tukey-Kramer HSD test ( = 187; df = 2, 323; < 0.0001). Improved method using corn and grass. 100

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Figure 3-1. Experimental design of Tests 1 and 2: A) oviposition container used to maintain adults and collect eggs, a nd B) container used for nymph development. These containers were limited by the amount of food and number of that could be housed. 101

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Figure 3-2. A 7.6-L oviposition ja r (A) used for maintaining adults and collecting eggs, and (B) a partially unrolled egg ro ll used in Tests 3, 4, and 5 displaying This method worked best for housing adult adults and oviposition. 102

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Figure 3-3. A 7.6-L glass jar with grass planted in sterili zed builders sand for nymph developm ent used in Test 3. This method failed because of contamination with predators and possibly limited food source. 103

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Figure 3-4. A 7.6-L glass jar with A) wax paper and cardboard assem blage at the bottom, and B) completely constructed jar with dental cast one used in Test 5, which allowed dental castone to be removed easily and safely so jars could be cleaned and reused. 104

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Figure 3-5. 7.6-L glass ja r containing St. Augustinegrass for development of nym phs used in Test 5. Fresh grass is added twice per week and placed near the bottom of the jar. This method worked best for producing the highest number of adults. 105

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106 Fertilize and cut grass weekly 15 min Place collected adults into castoned jar with corn and egg rolls 15 min Place collected eggs into castoned jar with grass 5m i n Wash 2 jars 10 min Fluon and castone 2 jars 1 h Shuck, soak, rinse, and store 4 ears of corn 20 min Change corn and vacuum dead and eggs lying on bottom of jar 30 min Plant 11 St. Augustinegrass plugs 20 min Collect eggs and place new egg rolls into jar 15 min Place fresh, air-dried grass under old grass material <5 min Remove emerged adult 45 min Cut and wash 1 St. Augustinegrass plant 1 5m i n Figure 3-6 Flow chart of steps and approxima te time and labor required to rear one jar of in a synchronous laboratory syst em (test five) at a constant 30 C and 14L:10D photoperiod. Each jar could produce 224 to 317 adults if initiated with 400 eggs.

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CHAPTER 4 CONCENTR ATION-MORTALITY RESPONSES TO FIVE INSECTICIDES BY A SUSCEPTIBLE COLONY OF USING AN AIRBRUSH BIOASSAY The southern chinch bug, Barber, is the most dama ging insect pest of St. Augustinegrass, (Walt.) Kuntze, in Florida causing stress and death of turfgrass (Reinert and Kerr 1973, Reinert and Niemczyk 1982, Bruton et al. 1983). All life stages are present throughout the growing season in densities of up to 2,000 chinch bugs/0.1 m2 (Reinert and Kerr 1973). In northern Flor ida, three to four generations of may occur from March to October, but seven to ten genera tions per year may occur in southern Florida (Kerr 1966, Reinert and Kerr 1973). Control of is mainly achieved through inse cticide use and, because it is multivoltine, insecticides may be applied up to 12 times a year in Florida (Reinert 1978, Reinert and Niemczyk 1982). As a result of consistent insecticide selection, has developed resistance to organochlorines, organophosphates, carbamates, neonicoti noids, and pyrethroids (Kerr 1958, 1961; Reinert 1982a, 1982b; Reinert a nd Niemczyk 1982; Reinert and Portier 1983; Cherry and Nagata 2005, 2007; Chapter 2). Given the history of in developing insecticide resistance, it is important to implem ent resistance management strategies that can prolong the effectiveness of existing or new insecticides for this pest. As with any resistance management program it is important to obtain information on current insecticide susceptibility levels in populations so that baselines can be established and changes in susceptibility over time and in different locations can be detected. The most commonly used labor atory bioassay for evaluating insecticide efficacy against is the sprig-dip test (Fi gure 4-1) (Reinert and Portie r 1983; Cherry and Nagata 2005, 2007; Congdon and Buss 2006; Chapter 2). This method involves cutting sections of St. 107

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Augustinegrass stolons, dipping them into insec ticide solutions, allowing them to dry, and placing them into petri di shes containing ten adult The set up for the sprig-dip bioassay can be conducted quickly and it is inexpens ive. However, a large degree of variability in response occurs. Tests are usually conducted in differe nt laboratories under varying environmental conditions, or with field-collected of unknown age and/or from different locations (Reinert and Portier 1983; Cherry and Nagata 2005, 2007; Congdon and Buss 2006; Chapter 2). It would be beneficial to ev aluate the sprig-dip bioa ssay under standardized conditions to validate the use of the assay. In addition to variability in the sprig-dip bioassay, scoring multiple individuals in the same dish can be cumbersome when they are not a ll moribund. A standardized bioassay that could detect differences between male and female would also greatly aid in understanding how insecticide resistance develops in this pest (i.e., mode of inher itance, stability of resistance). Thus, bioassays that made scoring easier would be an important asset for resistance monitoring. The goal of this study was to 1) evaluate the sprig-dip bioassay under standardized conditions, 2) develop a bioassay that could be used for detecting insecticide susceptibility differences between male and female and 3) validate both bioassays and determine optimal exposure times and sample sizes to be used for each bioassay for selected insecticides. Commercially-obtained plugs of Palmetto St. Augustinegrass were cut in half, planted in 8.9-cm plastic cups filled with Farfard #2 potting soil (Conrad Farfard Inc., Agawam, MA), and roots were allowed to establish for 3-4 wk before use in experiments. Plants were maintained in a University of Florida gree nhouse in Gainesville, FL, and held under a 14L:10D photoperiod 108

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and day and night temperatures of 27 and 24 C, respectively. Plants were fertilized weekly with 20-20-20 water-soluble complete N source (NH4NO3) at 0.11 kg N/0.09 m2, and watered and cut to a height of 7.6 cm as needed. fifth instars and adults were co llected from St. Augustinegrass on the University of Florida campus using a modi fied Weed Eater Barracuda blower/vacuum (Electrolux Home Products, Augusta, GA) (Crocker 1993, Nagata and Cherry 1999, Congdon 2004). This area was not treated with insecticides within 1 yr of this study and preliminary tests indicated that this population was the most susceptible compared to 16 populations tested against bifenthrin in Chapter 2. Insects were sorted from the debris and placed into 7.6-L glass colony jars. Each jar contained two full-sized, surfacesterilized, fresh yellow corn cobs and 12 pieces (7.6 7.6-cm) of cotton diaper towel (Tiger Acce ssory Group, LLC, Lincolnshire, IL) rolled to 1.7-cm diameter, then provided as an oviposition substrate. Cott on rolls were collected after 1 wk and placed into 7.6-L glass jars containing Palmetto St. Augus tinegrass plants that were cut at the crown and washed. Two 15.2-cm plants were added weekly until adults emerged. Secondand third-generation males and females were used for this study. Formulated products commonly used for control in Florida were chosen to represent four classes of insecticides (Table 41). Five to eight concentrations plus a water control were tested for each insecticide to estab lish probit lines with mortality ranging from 5 to 95% for unsexed The range of concentrations test ed for each insecticide is shown in Table 4-1. All bioassays were conducted between 13:30 and 15:30 h and held in growth chambers with a constant temperat ure of 26C and a 14L:10D photoperiod. 109

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All treatm ents in the airbrush bioassays were applied using a single-action Paasche airbrush (Figure 4-2A) (Model: H-set, Paasche Airbrush Company, Harwood Heights, IL), using the nozzle provided to deliver a fine aerosol spray. Lesco Tracker spray indicator dye (Lesco, Inc., Strongsville, OH) was used in initial tests to ensure that all insecticides were sprayed to runoff. New attachments were us ed for each insecticid e to eliminate crosscontamination. Half the solution was sprayed onto plant material, plants were rotated 90, and the remainder of the solution was sprayed to provide uniform coverage. Airbrush parts were cleaned with acetone. To determine uptake time for systemic insectic ides, two Palmetto St. Augustinegrass plants planted in 8-cm plastic cups were placed into th e center of a 929-cm2 cardboard tray and sprayed with clothianidin (1, 3, or 7 d before bioassay) and allowed to dry at room temperature (25 2C) and a 14L:10D photoperiod. Sections 1 cm in length containing a single node were cut from treated plants and placed into each cel l of a BioServe bioassay tray (Figure 4-2B) (BAW128, Bio-Serve, Frenchtown, NJ) that had b een swabbed with unscented Bounce fabric softener (Procter & Gamble, Cincinnati, OH) to reduce static electricity. Control cells were also swabbed to verify that the fabric softener did not affect the insects. One adult (2 wk old) of unknown sex was introduced into each cell. Cells were sealed with BioServe perforated tray lids (BACV16) and all trays were placed into clos ed plastic containers (35.6 cm 26.7 cm) lined with moistened paper towels to maintain humidity. The number of dead was assessed after 4, 8, 24, 48, and 72 h. were scored as dead if they were on their backs or unable to walk. A total of 144 and nine concentrations were tested for each spray time. 110

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A side-by-side comparison of the airbrush and sprig-dip bioassays was perform ed using 280 for bifenthrin (TalstarOne) and 720 for imidacloprid (Merit 2F). The airbrush bioassays were perf ormed as previously described (e xcept that insects were scored after 24, 48, and 72 h) and the sprig-dip bioassay s were conducted as described in Chapter 2. Both bioassays were placed into closed plastic containers as previously described. Because response to insecticides in indi vidual insects cannot be determ ined when using the sprig-dip bioassay, insects in comparis on tests were unsexed. Plants were treated with cont act insecticides 1 d before th e bioassay, except trichlorfon. Because trichlorfon degrades rapidly, plants were only dried for 2 h before the bioassay. Systemic insecticides were applied 3 d before the bioassay to allow for root uptake. The bioassay was set up as previously described. Each treatment was replicated six times fo r a total of 96 insects per concentration ( = 672 for bifenthrin, carbaryl, and clothian idin; 864 for imidacloprid; and 576 for trichlorfon). The location of each insect ( on or off the plant) and the number of dead were assessed after 1, 4, 8, 24, 48, and 72 h. mortality was scor ed as previously described. Insects were sexed at the end of the experiment using a dissecting microscope. The LC50 and LC90 values, 95% confidence limits (CL), slopes of the regression lines, and concentration-response relationshi ps were estimated by probit an alysis. In addition, likelihood ratio tests to examine the hypothesis of paralle lism and equality of th e regression lines among individual replicates were used to determine va riability in the bioassays in the comparison test using PoloPlus (LeOra Software 2002). 111

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To determine the appropriate exposure time for insecticides used in all tests, differences between LC50 and LC90 values for the different scoring in tervals within each sex or bioassay were determined by the 95% CL of lethal con centration ratios (LCRs). LCR confidence limits (95%) that did not include 1.0 were considered significant ( < 0.05) (Robertson and Priesler 1992, Robertson et al. 2007). Conventionally, if the 95% confidence lim its of the lethal concentrations overlapped, then the lethal con centrations were not c onsidered significantly different. However, the ratio test has greater statistical power and lower Type I error rates, so this statistical test was used in this study (Wheeler et al. 2006, R obertson et al. 2007). Subsamples of the comparison test data were taken in order to determine if smaller sampling sizes could be used fo r the sprig-dip and airbrush bi oassays. This was done by taking the raw concentration-mortality data from the optimal exposure time (as described above) for both bioassays and for each insecticide tested (b ifenthrin and imidacloprid), and entering into columns in Mircrosoft Excel. In empty columns next to each data set, random numbers were assigned to raw data using the formula =RAND() a nd typing Ctrl + Enter. The formula was then dragged down each column, assigning random numbers to all cells in the adjacent column. Columns were then sorted (ascending to descen ding) and subsamples were chosen starting with the first cell and subsequent ce lls until the desired subsample was taken (i.e., sample of 10, 20, 30 for each concentration). Data sets were re -sorted for each subsample taken. Subsamples were analyzed using PoloPlus and LCRs (Robe rtson and Priesler 1992, Robertson et al. 2007) were used to determine signi ficant differences between LC50 and LC90 values compared to the original sample size used in the comparison test. For the airbrush bioassay, the significance of differences between LC50 and LC90 values for male and female recorded at 24, 48, and 72 h was determined by the 95% CL of 112

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LCRs at the LC50 and LC90 (Robertson and Priesler 1992, Robert son et al. 2007). In addition, an analysis of variance (ANOVA) was conducted to determine differences between male and female in their ability to locate plant material within the first hour of the airbrush bioassay. If significant, treatme nt means were analyzed using the Tukey-Kramer (HSD) test using Jmp (SAS Institute Inc. 2001). The concentration-mortality data for exposed to St. Augustinegrass treated with clothianidin 1, 3, and 7 d before bioassay are shown in Table 4-2. LC90 values obtained at the 4and 8-h scoring intervals for exposed to plants treated 1 d prior to testing were significantly higher than LC90 values from respective scoring in tervals for plants treated 3 and 7 d previously. However, LC values obtained from exposed to 3and 7-d-treated plants were not different. Because clothianidin and imidacloprid are simila r in water solubility, a 3-d interval from spray to test was chosen for both products in the airbrush bioassay. The sprig-dip bioassay produced significantly lower LC50 values at all sc oring intervals, as well as LC90 values at 48 and 72 h, compared to the airbrush bioassay for bifenthrin (Table 43). The slope values for the sprig-dip bioassay were also lower than those for the airbrush bioassay (1.2-1.4 and 2.1-2.3, respec tively). Hypothesis tests for e quality and parallelism of the regression lines for each replicate for the spri g-dip bioassay show that slopes were not significantly different, but the intercepts were (Fi gure 4-3 A and C). The intercept of a probit or logit regression should correspond with the response that occurs with no treatment (Robertson et al. 2007), but control mortality was not observed. Differences between intercepts could have 113

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been due to physical processes (i .e., absorption through the cuticle or gut, target site sensitivity, or excretion) (Robertson and Preisler 1992, R obertson et al. 2007). However, if physical processes were the cause, it woul d likely have shown up in th e airbrush bioassay, as well, because from the sam e colony, generation, and age were used for both bioassays and all assays were conducted at the same time unde r the same conditions. It is possible that variability between replicates in the sprig-dip bioassay occurre d due to the large degree of untreated surface area in petri dishes, resu lting in differences in the ability of to locate plant material. Regression lines for replicat es in the airbrush bioassay were more similar (Figure 4-3 B and D) and result s of the hypothesis tests for equa lity and parallelism of the regression lines for each replicate showed that slopes and intercepts were not significantly different (Figure 4-3 B and D). This suggest s that there was a more uniform response among insects in the airbrush bioassay th an in the sprigdip bioassay. Comparisons of LC50 and LC90 values within each bioassay to determine appropriate exposure time for bifenthrin show the response at 24 and 48 h was similar to 72-h values with use of the airbrush bioassay [LCR50 for 24 h: 0.8 (0.5-1.2), LCR50 for 48 h: 1.0 (0.7-1.4); LCR90 for 24 h: 0.9 (0.5-1.5), LCR90 for 48 h: 1.0 (0.6-1.7)]. Fo r the sprig-dip bioassay, the LC50 values for the different scoring intervals we re not different; however, the 24-h LC90 values were significantly higher than respec tive values for 48 and 72 h [LCR50 for 24 h: 0.5 (0.3-1.1), LCR50 for 48 h: 0.9 (0.4-1.9); LCR90 for 24 h: 0.3 (0.2-0.8), LCR90 for 48 h: 0.8 (0.4-1.6)]. This suggests that when using the airbrush method for te sting bifenthrin, assays can be run for 24 h to estimate reliable LC50 and LC90 values. However, when using the sprig-dip bioassay, bifenthrin assays should be run for a minimum of 48 h to generate both LC50 and LC90 values. Considering the fast action of pyrethroids, the longer test time required for the sprig-dip bioassay may be due 114

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to the larger untreated surface area in the petri d i shes, resulting in differences in the time needed for to locate plant material. This fact or may also explain the high variability observed between replicates (Figure 4-3 A and C). Alternatively, pyrethroids often act as repellents and thus may have caused to avoid the plant materi al in the petri dishes. However, in the airbrush bioassay, the close quarters in the trays gr eatly reduces this variable. The sprig-dip bioassay produced significantly lower LC50 values at the 48and 72-h scoring intervals and higher LC90 values at 24 h compared to the airbrush bioassay for imidacloprid (Table 4-3). However, were more susceptible to imidacloprid after longer exposure (48 and 72 h) in both bioassays (Table 4-3). The LCR results comparing 72-h LC values to respective results obtained at 24 h within each bioassay show both bioassays had significantly higher LC50 and LC90 values at 24 h compared to re spective 72-h values [sprig-dip: LCR50 for 24 h: 0.4 (0.3-0.6); LCR90 for 24 h: 0.2 (0.1-0.3)], [airbrush: LCR50 for 24 h: 0.5 (0.40.7); LCR90 for 24 h: 0.6 (0.4-0.9)]. Howe ver, 48-h LC values were similar to those for 72 h for both bioassays [sprig-dip: LCR50 for 48 h: 0.8 (0.6-1.0); LCR90 for 48 h: 0.8 (0.6-1.2)], [airbrush: LCR50 for 48 h: 0.8 (0.6-1.0); LCR90 for 48 h: 0.9 (0.6-1.3)]. This would suggest that assays should run for at least 48 h in both bioassays to account for increas ed susceptibility to imidacloprid and to obtain both LC50 and LC90 values. These data are similar to other reports of increased susceptibility to neonicotinoids after longer exposure times. Prabhaker et al. (2006) also reported increased susceptibility to the neonicotinoids acetamiprid and imidacloprid after 48 h compared to 24 h in populations of (Say) (Hemiptera: Cicadellidae). Results for comparisons of replicates within each bioassay show wide variability at all scoring intervals for the sprig-dip bioassay for imidacloprid (Figure 4-4 A and C). Hypothesis tests of equality and parallelism of the different regression lines for each replicate show that 115

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slopes and intercepts were signif icantly different at the 24-h scoring interval, but after 48 h slopes were similar while intercep ts remained di fferent (Figure 4-4 A and C). By contrast, the slopes and intercepts of regression lines for th e different replicates were similar and more consistent for the airbrush bioassay (Figure 44 B and D), suggesting that insects were more uniform in response and in their ability to come into contact with plant material. The subsamples taken of the comparison study with bifenthrin using the airbrush bioassay produced similar LC50 values and narrow CLs compared to the original sample size of 280 (Table 4-4). Subsample sizes from 70-210 had slopes ranging from 1.7-2.7. Slopes and intercepts of regression lines for each subsample were similar to those of the original sample size of 280 (hypothesis test of equality = accept: 2 = 11.5, df = 6, = 0.07; parallelism = accept: 2 = 3.5, df = 3, = 0.37). These findings suggest that when using the airbrush bioassay to test bifenthrin, a sample size of 70 coul d be used to determine a reliable LC50 value. However, for estimation of reliable LC90 values, CL limits widen greatly as sample sizes are reduced from 210 to 70 (Table 4-4). Thus, to avoid excessively wide confidence limits at higher probit mortality levels, a sample size of 210 would be best. For the sprig-dip bioassay, the subsamples ta ken of the comparison study with bifenthrin produced LC values that were not significantly different from the original sample size of 280 (Table 4-4). The slopes ranged from 1.3-1.9 and slopes and interc epts of regression lines for each subsample were similar to those of the or iginal sample size of 280 (hypothesis test of equality = accept: 2 = 2.3, df = 6, = 0.89; parallelism = accept: 2 = 2.1, df = 3, = 0.55). However, the subsamples only produced an LC50 value for one of the subsamples (210) (Table 44). Concentration-mortality lines could not be produced for subsample sizes of 140 and 70 116

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because variability was too high. Based on these results, a minim um of 210 could be used when testing bifenthrin using the sprig-dip bioassay for estimating LC50 and LC90 values. A subsample size of 90 was sufficient (Table 4-5) for estimation of LC50 and LC90 values for imidacloprid using the airbrush bioassay (Table 4-5). LC50 values obtained for all subsample sizes were similar to those of the original sample size (LC50 = 4.2 and LC90 = 20.9) and CL limits were relatively narrow (Table 4-5). The slopes ranged from 1.5-2.1 and slopes and intercepts of regression lines for each subsampl e were similar to those of the original sample of 720 (hypothesis test of equality = accept: 2 = 6.0, df = 14, = 0.97; parallelism = accept: 2 = 5.1, df = 7, = 0.65). Concentration-mortality data for the sprigdip bioassay also demonstrated similar LC50 and LC90 values compared to the data from the or iginal sample size of 720. The slopes were more variable, ranging from 1.6-2.4 and slopes and intercepts of regression lines for each subsample were similar to those of the 720-sa mple size (hypothesis test of equality = accept: 2 = 6.9, df = 14, = 0.94; parallelism = accept: 2 = 4.5, df = 7, = 0.94). However, PoloPlus was unable to provide estimates for the 180-sample size. This may have been an artifact of the subsample obtained or it may indi cate this is too small a sample for estimation of LC values for imidacloprid using the sprig-dip bioassay. If the latter condition is true, then a sample size of 270 should be used for estimation of LC50 as CL limits are still narrow at this sample size. A sample size of 360 should be used for estimation of LC90 values for imidacloprid because CL limits widen considerably when smaller sample sizes were used (Table 4-5). Robertson et al. (1984) determined that a sample size of 120 was the minimum necessary for calculating a reliable LC50 estimation, but for increased precision sample sizes of 240 or more are often necessary (Robertson et al. 1984, 2007; Robertson and Preisler 1992). However, the 117

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authors noted that further investigation is needed to explore other combinations of sample size and dose placement in different bioassays (Robert son et al. 2007). Cherry and Nagata (2007) reported LC50 values for exposed to imidacloprid fo r 24 h using the sprig-dip bioassay with sample sizes ranging from 120-360 In this study, LC values and sample sizes varied depending on the insecticid e tested and bioassay used. These subsample data indicate that when using the sprig-dip bioassay, the optimal exposure time is 48 h using a minimum of 270 when testing approximately 8 c oncentrations of imidacloprid for estimation of LC50 values and 360 for estimation of the LC90. In terms of cost for supplies for each bioassa y, the BioServe bioassa y trays (BAW128) and perforated tray lids (BACV16) us ed in the airbrush bioassay cost 3.5 per insect. The cost of petri dishes (cat. number 08757-12, Fisher Scientific, Pitts burgh, PA) and 70-mm Whatman filter paper (cat. number 1002 070, Whatman Interna tional Ltd, Maidstone, England) for use in the sprig-dip bioassay is 4.0 per insect. The ti me to set up each bioassay will vary depending on the number of concentrations tested and the number of replicat es at each time interval. Not including the making of serial di lutions or treatment of plant material, one person can set up a single replicate of the airbrush bioassay with eigh t concentrations in 1-2 h. For the sprig-dip bioassay, one replicate with the sa me number of concentrations could be set up in 30 min to 1 h. The set up times for both bioassays may be greatl y reduced if insects are anesthetized before introduction into tests; however, the effects of anesthetizing insects prio r to insecticide exposure currently are not known. While the sprig-dip bioassay takes less time to set up per replicate, these data indicate that larger sa mple sizes and/or exposure times (for bifenthrin) are required for estimation of LC50 and LC90 values. One will need to take in to consideration increased costs for 118

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collecting or rearing larger numbers of insects in addition to increased costs for supplies and labor when using the spring-dip bioassay. The BioServe bioassay trays were usef ul f or testing insecticides against and have been used successfully with Coleoptera (Lalitha et al. 2005), Lepidoptera (Bomford and Isman 1996, Simmonds and Stevenson 2001, Kokubun et al. 2003, Zoerb et al. 2003, Akhtar and Isman 2004, Oigiangbe et al. 2007), and Thysanopt era (Brown et al. 2003). The trays allowed easier observation of individual insects (including differences between males and females, behavior, recovery time) and reduced evaluation ti me compared to the sprig-dip bioassay. Prior bioassays involving sprig dips (Congdon and Buss 2004, 2006) were time consuming when scoring and observations of indi vidual insects were difficult. The BioServe perforated lids helped keep St. Augus tinegrass succulent for feeding for the duration of each test while allowing adequate ventilation. The use of Bounce fabric softener for static charge reduction did not appear to affect as there was at most 2 3% mortality observed in controls after 72 h. Male and female located plant material equally well in all tests within 1 h (Table 4-6). Due to variability in the time at which insects were in contact with and/or fed on plant material, concentration-mortality lines were only determined for the 24-, 48-, and 72-h intervals. LC50 and LC90 values were within the range of concentrations tested for each insecticide (Table 4-1) with the exception of bifenthrin. The LC90 values for females after 24, 48, and 72 h were greater (32.9, 31.9, and 30.5 g/ml, respectively, Table 4-7) than the highest concentration of bifenthrin us ed in this study (26 g/ml) i ndicating these are calculated estimates. Using a higher concen tration may have provided true LC90 values for females exposed to bifenthrin. 119

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Of the five insecticides tested using the ai rbrush bioassay, clothian idin and imidacloprid were m ost toxic to (regardless of sex), particular ly after 48 and 72 h, followed by bifenthrin (Table 4-7). Carbaryl and trichlorfon were the least toxic of the insecticides tested, which may in part be due to their short residual and/or volatility in turfgrass. Bifenthrin, carbaryl, and trichlorfon are often used for contro l of surfaceand/or subs urface-feeding pests in turf (Clark and Kenna 2001). Ne onicotinoids, such as clothianid in and imidacloprid, are known to be highly effective against piercing-sucking insects (Cahill et al. 1996; Elbe rt et al. 1996; Nauen et al. 1996, 1998; Tomizawa and Casida 2005; Magalhaes et al. 2008). The low affinity of neonicotinoids for vertebrate compared to in sect nicotinic acetylchol ine receptors (Tomizawa and Casida 2005), along with a l ong residual life, allows clothian idin and imidacloprid to be applied at lower rates compared to insecticides with shorter residual lives such as carbaryl and trichlorfon. The slope values obtained af ter each scoring interval fo r the contact insecticides (bifenthrin, carbaryl, and trichlor fon) are steep, suggesting that males and females responded uniformly to these products (Geor ghiou and Metcalf 1961; ffrench-Constant and Roush 1990; Prabhaker et al. 1996, 2006). However, the slope values obtained for the systemic insecticides (clothianidin and imidacloprid) we re lower and more va riable across scoring intervals for male and female particularly for imidacloprid. Alternatively, it is possible that the population that was tested was more heterogeneous in its response to these insecticides as compared to the contact insecticides (Table 4-6). LC50 values at all scoring in tervals showed that male were significantly more susceptible to bifenthrin, carbaryl, clothianidin, and imidacloprid than females (Table 4-7). However, the LC90 values for clothianidin are not different between male and female 120

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. Both sexes demonstrated an equal respons e to trichlorfon at all scoring intervals (Table 4-7). The baseline data generated using the airbrush bioassay showing that female are more tolerant to most of the insecticides tested is not unusual, given that females are significantly larger than male (Cherry and Wilson 2003). However, size cannot always be used as an indicator for predicting sexu al differences in susceptibility. For example, (Stl) (Hemiptera: Pentatomidae) male s were more tolerant to thiomethoxam despite being smaller than females (Nielson et al. 2008). Also, one sex may show more tolerance to one insecticide but not another (de Lane et al. 2001, Shearer and Usmani 2001). Sexual differences in insecticide susceptibili ty also occur in other insects, including L. (Hemiptera: Cimicidae) (Busvine and Lien 1961), (Ashmead) (Hymenoptera: Eulophidae) (Rathman et al. 1992), (Busck) (Lepidoptera: Tortricidae) (de Lame et al. 2001, Shearer and Usmani 2001), and (Nielson et al. 2008). This study is the first to document differences in insecticide susceptibi lity between male and female Analysis of LC50 and LC90 values for the different scoring intervals within each sex show that the responses of male and female after 24 and 48 h were similar to their responses after the 72-h interval for the contact insecticides bife nthrin, carbaryl and trichlorfon (Table 48). This suggests that when using th e airbrush method for test ing these insecticides, tests can be scored af ter 24 h to obtain both LC50 and LC90 values for male and female However, when testing the systemic inse cticides clothianidin and imidacloprid using the airbrush bioassay, tests shoul d be run for at least 48 h befo re scoring results. The LCR50 for clothianidin showed LC50 values after 24 h for females we re significantly hi gher than their 121

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122 respective 48-h and 72-h values (Table 4-9). LC50 and LC90 values recorded for males and females after 24 h were significantly hi gher for imidacloprid (Table 4-9). The results of this study i ndicate that when using the airbrush bioassay, contact insecticides should have an exposure time of 24 h while systemic ones should run for 48 h. These data are consistent with the results of the airbrush bioassay conducted in the comparison test, but it would be useful to compare th e two bioassays using other insecticides and populations because responses may differ (ffren ch-Constant and Roush 1990, Scharf et al. 1995, Studebaker and Kring 2003). As part of a resistance management program, it is recommended that the sprig-dip bioassay be used for detecti on of bifenthrin-resistant populations because it was more sensitive in detecting lower LC values in this study. The airb rush bioassay would be beneficial for use in studies concerning cro ss resistance, resistance mechanisms, mode-ofinheritance, and st ability of pyrethroi d resistance in In addition, the airbrush bioassay could be used for determining a dia gnostic dose for pyrethroidand imidaclopridresistant populations because this bioassay results in less variance.

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Table 4-1. Insecticides tested against a suscep tible colony of Class Compound Registered Recommended Range Company name label rate* used Carbamate Carbaryl Sevin SL 5.622 7496 g/ml 7.3-468 g/ml Bayer Environmental Science, Research Triangle Park, NC Neonicotinoid Clothianidin Arena 50 WDG 368 g/ml 0.7-23 g/ml Arysta Life Science, San Francisco, CA Imidacloprid Merit 2F 375 g/ml 0.7-93 g/ml Bayer Environmental Science, Research Triangle Park, NC 123Organophosphate Trichlorfon Dylox 80 T & O 976 1,464 g/ml 23-366 g/ml Bayer Environmental Scienc e, Research Triangle Park, NC Pyrethroid Bifenthrin TalstarOne 209 g/ml 0.8-26 g/ml FMC Corporation, Philadelphia, PA Label rates calculated using a sp ray application vol ume of 11.3 L/92.9 m2.

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Table 4-2. Concentration-mortality data (at LC50 and LC90) at different exposure times for a susceptible laboratory colony exposed to St. Augustinegrass treated with clothianidin 1, 3, and 7 d before bioassay. Treatment time (d) Exposure tim e (h) Slope SE LC50 (95% CL)aLCR50 (95% CL)b LC90 (95% CL)a LCR90 (95% CL)b x2 (df)c 144 4 1.0 0.2 14.5 (3.5.6) --236 (80.3,284) --10.0 (6)d 7 8 1.5 0.3 8.5 (1.4.9) --58.6 (24.4) --12.1 (6)d 24 1.3 0.4 1.7 (0.1.8) --15.7 (8.3.0) --5.7 (6)d 48 1.3 0.4 1.3 (0.02.0) --11.4 (5.4.5) --3.4 (6)d 72 1.5 0.5 1.3 (0.04.1) --9.3 (4.8.4) --1.1 (6)d 144 4 1.4 0.2 15.9 (5.1.1) 0.9 (0.4.8) 121(49.1,692) 1.9 (0.5.4) 13.3 (6)d 3 8 1.2 0.2 8.4 (0.5.9) 1.0 (0.4.5) 101 (35.3,773) 0.6 (0.1.7) 11.2 (6)d 24 1.9 0.6 2.0 (0.2.6) 0.8 (0.1.8) 9.3 (5.6.4) 1.7 (0.6.3) 3.1 (6)d 48 1.9 0.6 2.0 (0.2.6) 0.6 (0.1.4) 9.3 (5.6.4) 1.2 (0.4.1) 3.1 (6)d 124 72 1.9 0.7 1.8 (0.09.4) 0.8 (0.1.9) 8.4 (4.9.9) 1.1 (0.4.7) 1.7 (6)d 144 4 1.7 0.3 7.9 (4.7.7) 1.8 (0.5.4) 45.3 (28.0) 5.2 (1.6.4)* 4.7 (6)d 1 8 2.0 0.4 4.9 (2.8.3) 1.7 (0.8.4) 21.9 (14.2.6) 2.7 (1.1.3)* 4.3 (6)d 24 2.1 0.7 2.0 (0.2.5) 0.8 (0.1.5) 8.4 (5.3.3 1.9 (0.7.6) 1.1 (5)d 48 2.0 0.7 1.9 (0.1.3) 0.6 (0.1.5) 7.6 (4.8.8) 1.5 (0.6.7) 0.5 (5)d 72 2.1 0.7 1.9 (0.1.3) 0.7 (0.1.4) 7.6 (4.8.8) 1.2 (0.5.9) 0.5 (5)d a LC50 and LC90 values in g/mL (95% confidence limits). b Lethal concentration ratios with 95% conf idence limits indicating the fold-difference for males for each insecticide in compar ison to the respective female scoring interval at LC50 and LC90. Confidence limits that incl ude 1.0 indicate no significant difference from female scoring interval ( 0.05). Shows ratios that are significant. c Pearson chi-square statis tic (degrees of freedom). d Good fit of the data to the probit model ( > 0.05).

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Table 4-3. Comparison of concentration-mortality data (at LC50 and LC90) for a susceptible laboratory colony to bifenthrin and imidacloprid at 24, 48, and 72 h using the airbrush and sprig-dip bioassays. Bioassay used Insecticide tested Test tim e (h) Slope SE LC50 (95% CL)b LCR50 (95% CL)c LC90 (95% CL)b LCR90 (95% CL)c x2 (df)d Bifenthrin 24 1.2 0.2 1.4 (0.3-2.8) 0.2 (0.1.4)* 17.1 (7.6-221) 0.8 (0.4.7) 7.4 (4)e Sprig-dip 48 1.3 0.2 0.8 (0.11.6) 0.2 (0.1.3)* 7.6 (4.1-33.5) 0.4 (0.2.8)* 5.6 (4)e 72 1.4 0.2 0.8 (0.04.6) 0.2 (0.1.3)* 6.1 (3.0-68.4) 0.3 (0.2.6)* 9.1 (4)e Airbrush 24 2.3 0.2 5.6 (3.2-10.6) --20.2 (10.7-111) --12.6 (4) 48 2.2 0.2 4.8 (3.1-7.0) --17.6 (10.7-40.9) --6.7 (4)e 72 2.1 0.3 4.5 (1.7-7.4) --18.0 (10.6-78.8) --6.8 (4)e Sprig-dip Imidacloprid 24 1.1 0.1 5.9 (3.9-8.8) 1.0 (0.7.2) 82.4 (44.3223) 2.8 (1.6.5)* 12.0 (6) 48 1.8 0.1 3.1 (2.5-3.8) 0.7 (0.5.9)* 16.5 (13.1-22.1) 0.8 (0.5.1) 6.0 (6)e 72 1.7 0.1 2.5 (1.8-3.2) 0.7 (0.5.9)* 14.0 (10.3-21.1) 0.8 (0.5.1) 6.5 (6)e 24 1.9 0.1 6.1 (5 .2-7.1) --29.7 (23.4.9) --4.7 (6)e Airbrush 125 48 1.8 0.1 4.2 (3 .2-5.3) --21.1 (15.4-31.7) --7.8 (6)e 72 1.7 0.1 3.3 (2.6-4.1) --18.3 (13.7-27.0) --6.3 (6) e a = 280 were tested for each bioassay for bifenthr in and 720 for each bioassay for imidacloprid. b LC50 and LC90 values in g/mL (95% confidence limits). c Lethal concentration ratios with 95% confidence limits for the sprig dip bioassay indicating th e fold-difference for each test time in comparison to respective airbrush test times at LC50 and LC90. Confidence limits that include 1.0 indicate no significant difference from the respective airbrush test time ( 0.05). Indicates ratios that are significant. d Pearson chi-square statis tic (degrees of freedom). e Good fit of the data to the probit model ( > 0.05).

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Table 4-4. Comparison of subsampleda concentration-mortality data (at LC50 and LC90) for a susceptible laboratory colony exposed to bifenthrin using th e airbrush and sprig-dip bioassays. Bioassay used Sample size Slope SE LC50 (95% CL)b LCR50 (95% CL)c LC90 (95% CL)b LCR90 (95% CL)c x2 (df)d 280 1.3 0.2 0.8 (0.1.6) --7.6 (4.1.5) --5.6 (4)e Sprig-dip 210 1.3 0.2 0.9 (0.21.7) 0.9 (0.4.9) 8.7 (4.640.4) 0.9 (0.4.8) 4.2 (4)e 140 N/A N/A N/A N/A N/A N/A 70 N/A N/A N/A N/A N/A N/A Airbrush 280 2.3 0.2 5.6 (3.2.6) --20.2 (10.7) --12.6 (4) 210 2.7 0.3 5.9 (3.510.7) 0.9 (0.7.2) 17.6 (10.083.1) 1.1 (0.6.9) 10.0 (4) 140 2.4 0.4 7.5 (3.523.7) 0.7 (0.5.0) 25.0 (11.4852) 0.8 (0.4.5) 11.3 (4)e 70 1.7 0.4 3.8 (2.26.6) 1.5 (0.8.5) 21.6 (11.0108) 0.9 (0.3.5) 3.1 (4)e a Subsamples were selected from the comparison test using data from the 48 h sprig-dip and 24 h airbrush bioassays for bifenthri n. Scoring intervals were chosen based on the appropriate am ount of time needed to run each bioassay for bifenthrin. 126b LC50 and LC90 values in g/mL (95% confidence limits). c Lethal concentration ratios with 95% confidence limits for each bioassay indicating the fold-di fference for subsamples within each bioassay in comparison to respecti ve original sample size at LC50 and LC90. Confidence limits that incl ude 1.0 indicate no significant difference from the respective original sample size ( 0.05) d Pearson chi-square statis tic (degrees of freedom). e Good fit of the data to the probit model ( > 0.05).

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Table 4-5. Comparison of subsampleda comparison test concentration-mortality data (at LC50 and LC90) for a susceptible laboratory colony exposed to imidacloprid us ing the airbrush and sprig-dip bioassays. Bioassay used Sample size Slope SE LC50 (95% CL)b LCR50 (95% CL)c LC90 (95% CL)b LCR90 (95% CL)c x2 (df)d 720 1.8 0.1 3.1 (2.5.8) --16.5 (13.1.1) --6.0 (6)e Sprig-dip 630 1.8 0.1 3.2 2.3.1) 1.0 (0.7.3) 15.8 (11.524.1) 1.0 (0.7.5) 6.8 (6)e 540 1.6 0.1 3.0 (2.14.2) 1.0 (0.7.4) 18.3 (12.531.4) 0.9 (0.5.3) 7.6 (6)e 450 1.7 0.1 3.6 (2.35.0) 0.9 (0.6.2) 20.6 (13.439.0) 0.8 (0.5.2) 8.5 (6)e 360 1.7 0.2 3.4 (2.05.0) 0.9 (0.6.3) 18.8 (11.938.4) 0.9 (0.5.3) 8.2 (6)e 270 2.0 0.3 3.2 (0.87.0) 1.0 (0.6.4) 14.0 (6.5125) 1.2 (0.7.9) 24.8 (6) 180 2.0 0.3 N/A N/A N/A N/A N/A 90 2.4 0.6 4.3 (2.07.0) 0.7 (0.4.3) 14.9 (8.943.4) 1.1 (0.5.2) 3.7 (6)e 720 1.8 0.1 4.2 (3.3.3) --20.9 (15.4.7) --7.8 (6)e Airbrush 630 1.8 0.1 4.1 ( 3.15.4) 1.0 (0.8.3) 21.0 (14.735.0) 1.0 (0.6.4) 9.3 (6)e 127540 1.8 0.1 4.0 ( 3.24.8) 1.0 (0.8.3) 21.1 (16.229.5) 1.0 (0.6.4) 5.7 (6)e 450 1.8 0.1 4.1 (2.95.7) 1.0 (0.7.3) 20.4 (13.240.3) 1.0 (0.6.5) 10.6 (6)e 360 2.1 0.2 4.0 (3.35.0) 1.0 (0.8.3) 16.6 (12.524.4) 1.3 (0.8.9) 5.7 (6)e 270 1.8 0.2 4.2 (2.86.0) 1.0 (0.7.3) 22.5 (14.048.0) 0.9 (0.5.5) 7.1 (6)e 180 1.5 0.2 4.5 (3.06.5) 0.9 (0.6.4) 33.8 (20.574.1) 0.6 (0.3.2) 4.4 (6)e 90 1.8 0.3 4.2 (2.56.7) 1.0 (0.6.6) 21.8 (12.363.5) 1.0 (0.4.1) 1.6 (6)e a Subsamples were selected from the 48-h scoring intervals for both the sprig-dip and airbrush bi oassays for imidacloprid. Scor ing intervals were chosen based on the appropriate amount of time needed to run each bioassay for imidacloprid. b LC50 and LC90 values in g/mL (95% confidence limits). c Lethal concentration ratios with 95% confidence limits for each bioassay indicating the fold-di fference for subsamples within each bioassay in comparison to respecti ve original sample size at LC50 and LC90. Confidence limits that incl ude 1.0 indicate no significant difference from the respective original sample size ( 0.05). Shows ratios that are significant. d Pearson chi-square statis tic (degrees of freedom). e Good fit of the data to the probit model ( > 0.05).

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Table 4-6. The mean number SEM of male and female B. insula ris that located treated plant ma terial within 1 h of introductio n into the airbrush bioassay. Mean no. SEM Mean no. SEM -value df -value Insecticide Bifenthrin 0.34 0. 3 0.33 0.02 0.27 2, 670 0.76 Carbaryl 0.47 0.03 0.41 0.03 2.20 2, 670 0.14 Clothianidin 0.48 0.03 0.41 0.03 3.48 2, 670 0.05 Imidacloprid 0.50 0.02 0.46 0.02 1.49 2, 862 0.22 Trichlorfon 0.64 0.03 0.62 0.03 0.14 2, 574 0.70 128

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Table 4-7. Concentration-mortality data (at LC50 and LC90) compared for males and females from a susceptible laboratory colony treated with five inse cticides after 24, 48, and 72 h us ing the airbrush bioassay. Insecticide tested n sex Test time (h) Slope SE LC50 (95% CL)a LCR50 (95% CL)b LC90 (95% CL)a LCR90 (95% CL)bx2 (df)c 376 Males 24 2.5 0.2 6.4 (5.0.4) 0.5 (0.4.8)* 21.4 (15.136.6) 0.6 (0.31.3) 4.1 (4)dBifenthrin 48 2.4 0.3 6.0 (4.5.1) 0.6 (0.4.8) 20. 6 (13. 939.2) 0.6 (0.31.3) 5.4 (4)d 72 2.4 0.2 5.5 (4.3.2) 0.6 (0.4.7) 18. 6 (13. 032.3) 0.6 (0.40.9) 4.6 (4)d 296 Females 24 2.8 0.6 11.5 (9.2.6) --32.9 (21.1.6) --1.7 (3)d 48 2.7 0.6 10.7 (8.4.1) --31.9 (20.3.1) --2.2 ( 3 )d 72 2.6 0.3 9.7 (7.9.9) --30.5 (22.8.6) --2.2 ( 4 )d 334 Males 24 2.9 0.3 104 (88.03) 0.6 (0.5.8)* 287 (229391) 0.5 (0.30.7)* 1.0 (4)dCarbaryl 48 2.6 0.3 90.1 (59.5134) 0.6 (0.5.8)* 281 (178694) 0.7 (0.41.0) 8.6 (4) 72 2.6 0.3 88. 8 (59.7130) 0.6 (0.5.9)* 276 (178643) 0.7 (0.41.0) 8.0 (4) 338 Females 24 2.4 0.3 167 (121) --572 (361,410) --3.4 (3)d 129 48 2.7 0.3 142 (119) --424 (329) --2.1 (3 )d 72 2.7 0.4 139 (111) --407 (315) --1.0 ( 3 )d 344 Males 24 1.9 0.2 4.1 (3.3.0) 0.6 (0.4.9)* 18.8 (13.729.2) 0.6 (0.31.2) 3.4 (4)dClothianidin 48 2.0 0.2 3.7 (3.0.5) 0.7 (0.5.9) 16. 7 (12. 325.5) 0.7 (0.41.5) 2.0 (4)d 72 2.1 0.2 3.2 (2.2.3) 0.7 (0.5.9) 12. 1 (8.2 2.6) 0.6 (0.31.1) 4.5 (4)d 328 Females 24 1.8 0.2 6.4 (3.7. 6) --32. 6 (12. 7,619) --6.3 (3)d 48 2.1 0.3 5.4 (3.6.7) --22.2 (11.6) --4.0 (3)d 72 2.0 0.2 4.5 (2.8.7) --19.1 (9.5) --5.4 ( 3 )d 456 Males 24 2.5 0.2 10.5 (8.92.4) 0.6 (0.5.8)* 34.1 (27.145.9) 0.4 (0.30.7)* 2.5 (6)dImidacloprid 48 1.8 0.2 3.1 (2.1.3) 0.7 (0.5.9) 15. 7 (10. 330.9) 0.7 (0.41.1) 11.0 (6) 72 1.5 0.2 2.2 (1.3.2) 0.5 (0.4.8) 14.8 (9.12.9) 0.6 (0.30.9)* 11.0 (6) 408 Females 24 2.0 0.2 17.4 (14. 3.3) --74.7 (55.8) --5.3 (6)d

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Table 4-7. Continued. 48 1.9 0.2 4.6 (2.8. 9) --22.3 (13.4.4) --11.0 (5) 72 1.6 0.1 4.0 (3.0.2) --25.3 (18.4.5) --3.6 ( 6 )d Trichlorfon 277 Males 24 2.6 0.3 79.2 (34.1172) 1.0 (0.8.4) 244 (1273,202) 0.9 (0.61.5) 12.6 (3) 48 2.9 0.3 69. 6 (45.1102) 1.1 (0.8.4) 194 (127-480) 0.9 (0.61.4) 4.9 (3)d 72 3.1 0.3 64.3 (37.8102) 1.0 (0.8.4) 167 (105541) 0.9 (0.61.3) 6.9 (3) 299 Fem a les 24 2.4 0.4 74.7 (45. 8) --254 (161) --4.5 (3)d 48 2.5 0.2 64.7 (43.7. 2) --210 (144) --3.1 ( 3 )d 72 2.5 0.3 61.0 (37. 9.6) --193 (128) --4.1 ( 3 )d a LC50 and LC90 values in g/mL (95% confidence limits). b Lethal concentration ratios with 95% conf idence limits indicating the fold-differenc e for males for each in secticide in compar ison to the respective female scoring interval at LC50 and LC90. Confidence limits that incl ude 1.0 indicate no significant difference from female scoring interval ( 0.05). Shows ratios that are significant. 130c Pearson chi-square statis tic (degrees of freedom). d Good fit of the data to the probit model ( > 0.05).

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Table 4-8. Analysis of LC50 values for 24, 48, and 72 h within each sex to determine bioassay time for the contact insecticides bifenthrin, carbaryl, and trichlorfon. Insecticide Sex Time (hours) LCR50 a 95% CL LCR90 a 95% CL Bifenthrin M 24 0.9 (0.7-1.1) 0.9 (0.6-1.3) 48 0.9 (0.7-1.2) 0.9 (0.6-1.4) F 24 0.8 (0.6-1.1) 0.9 (0.5-1.5) 48 0.9 (0.7-1.2) 0.9 (0.5-1.5) Carbaryl M 24 0.8 (0.7-1.1) 1.0 (0.6-1.4) 48 1.0 (0.8-1.3) 1.0 (0.6-1.4) F 24 0.8 (0.6-1.0) 0.7 (0.4-1.1) 48 1.0 (0.7-1.2) 1.0 (0.6-1.5) 131Trichlorfon M 24 0.8 (0.6-1.0) 0.7 (0.4-1.0) 48 0.9 (0.7-1.2) 0.8 (0.5-1.2) F 24 0.8 (0.6-1.0) 0.8 (0.5-1.1) 48 0.9 (0.7-1.2) 0.9 (0.6-1.4) a Lethal concentration ratios with 95% conf idence limits indicating the fold-difference in 48 h scoring times in comparison to t he respective 24-h scoring interval with in each sex for each insecticide at LC50 and LC90. Confidence limits that include 1.0 indicate no significant difference from 24-h scoring interval ( 0.05). Shows ratios that are significant.

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132 Table 4-9. Analysis of LC90 values for 24, 48, and 72 h within each sex to determine bioassay time for the systemic insecticides clothianidin and imidacloprid.Insecticide Sex Time LCR50 95% CL LCR90 95% CL Clothianidin M 24 0.8 (0.6-1.0) 0.6 (0.4-1.0) 48 0.9 (0.6-1.1) 0.7 (0.4-1.2) F 24 0.7 (0.5-0.9)* 0.6 (0.3-1.0) 48 0.8 (0.6-1.0) 0.7 (0.4-1.2) Imidacloprid M 24 0.2 (0.1-0.3)* 0.4 (0.2-0.6)* 48 0.7 (0.5-1.0) 0.9 (0.5-1.4) F 24 0.2 (0.1-0.3)* 0.3 (0.2-0.5)* 48 0.9 (0.6-1.2) 1.2 (0.7-1.9) a Lethal concentration ratios with 95% conf idence limits indicating the fold-difference in 48-h scoring times in comparison to t he respective 24-h scoring interval with in each sex for each insecticide at LC50 and LC90. Confidence limits that include 1.0 indicate no significant difference from 24 h scoring interval ( 0.05). Shows ratios that are significant.

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Figure 4-1. The sprig-dip bioassay conventionall y used for testing insecticides against 133

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134 Figure 4-2. The (A) Paasche airbrush and (B) BioSer ve bioassay tray and lid used in the airbrush bioassay. Photos by C. Vzquez.

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135 Equality: accept Equality: reject 2=23.3; df=6; =0.001 2=4.6; df=6; =0.60 Parallelism : accept Parallelism : accept 2=5.2; df=3; =0.16 2=4.4; df=3; = 0. 22 Equality: accept Equa lity: reject 2=16.8; df=6; =0.01 2=4.1; df=6; =0.66 Parallelis m: accept Parallelism : accept 2=2.9; df=3; =0.41 2=4.1; df=3; = 0. 2 5 Figure. 4-3. The differences in variability between replicates of bifenthrin ( =280) for the (A) sprig-dip bioassay after 24 h, (B) airbrush bi oassay after 24 h, (C) sprig-dip bioassay after 48 h, and (D) airbrush bioassay after 48 h. Each regression line within a graph represents one replicate. The results of hypothesis tests for equality and parallelism of the regression lines among individual rep licates at each time interval are also shown.

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136 Equality: accept 2=15; df=14; =0.39 Parallelism: accept 2=5 ; df=7; =0.64 Equality: accept 2=10; df=14; =0.74 Parallelism: accept 2=1 ; df=7; =0.99 Equality: reject 2=155; df=14; <0.05 Parallelism: reject 2=21 ; df=7; <0.05 Equa lity: rej ect 2=54; df=14; <0.05 Parallelism: accept 2=11 ; df=7; =0.15 Figure. 4-4. The differences in variability between replicates of imidacloprid ( = 720) for the (A) sprig-dip bioassay after 24 h, (B) airb rush bioassay after 24 h, (C) sprig-dip bioassay after 48 h, and (D) ai rbrush bioassay after 48 h. Each regression line within a graph represents one replicate. The results of hypothesis tests for equality and parallelism of the regression lines among individual replicat es at each time interval are also shown.

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CHAPTER 5 CONCLUSIONS The goals of this dissertation were to 1) sam ple select populations in 2006 and 2008 in northern and central Florida to describe th eir susceptibility to bifenthrin, document new locations of bifenthrin resistan ce, and evaluate another pyrethroid, permethrin (Chapter 2), 2) develop a synchronous rearing method for that produces insects of known age and generation (Chapter 3) and 3) develop an impr oved bioassay that could be used for detecting insecticide susceptibility differences between male and female evaluate and validate both the sprig-dip and the new bioassay under st andardized conditions, and determine optimal exposure times and sample sizes to be used for each bioassay for selected insecticides (Chapter 4). The results of Chapter 2 show that bifenthrin resistance continues to be problematic, is becoming more widespread, and that there is a positive relationship between insecticide application and the development of bifenthrin resistance. Given the high number of insecticide applications observed in this study, resistance is likely to continue to spread into surrounding areas within the state unless management tactics are changed. In addi tion, the occurrence of cross resistance to other pyrethroi ds is evident from my data ( population JC to permethrin) and that of Cherry and Nagata (2007). Floridas warm climate and high number of pests increases the need for lawn care professionals, and results in greater use of pesticides compared to other states (Short et al. 1982). Ol kowski et al. (1978) reported pest icide use in the landscape is usually the result of response to aesthetic damage rather than a reaction to medical problems or economic losses. While homeowners find St. Augustinegrass damage aesth etically displeasing, it can create economic losses when sod needs to be replaced or multiple in secticide applications are required to gain control of damaging populations. One of the challenges that we 137

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face in dealing with the resistance problem in Flor ida will be to change the mindset of lawn care professionals, homeowners, a nd homeowner associations. Potter (1993) suggested that unn ecessary or excessive use of pesticides can increase problems with thatch and pests by reducing be neficial organisms already present in the landscape, encouraging the development of resi stance, or enhanced microbial degradation. Potter et al. (1990a, 1990b) demonstrated that Kentucky bluegra ss plots treated with either chlordane or carbofuran greatly reduced earthworm numbers and resulted in increased thatch compared to untreated contro ls. Reinert (1978) observed populations remained low in Florida St. Augustinegrass lawns that had an abundance of natural enemies and had not been treated with insecticides. C onversely, the author reported populations reached outbreak densities on insecticide-treated lawns (Reine rt 1978). It is clear from the many cases of resistance that have been reported over the last several decades (Wolfenbarger 1953; Kerr 1958, 1961; Reinert 1982a, 1982b; Reinert and Niemczyk 1982; Reinert and Portier 1983; Cherry and Nagata 2005, 2007) that many of Florida lawns ca n be considered high maintenance and receive considerable amounts of pesticides. Florida is second only to Calif ornia in terms of employment impacts of the turfgrass industry, providing 83,944 jobs in 2002 (Haydu et al. 2006). Considering the number of housing units in Florida increased from ~3.9 million in 1980 to 8.5 million in 2006 (an increase of 118%), the demand for quality turf and maintenan ce has likely increased (Haydu et al. 2005). In addition to meeting the demands of homeowners for high quality turf, lawn-care companies may also face high turnover of employees. New employees may not have developed the proper skills or been properly traine d to monitor and manage damage in lawns. Fothergill (1982) conducted a study in Massachusetts and found that the damage rate in lawns greatly increased as 138

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the length of employee service decreased. Furthe rm ore, the author noted that damage rate was more dependent on the experience and level of training employees received during and prior to their being allowed to monitor and treat lawns without direct supervis ion (Fothergill 1982). Another issue that is likely adding to resistance problems in Florida lawns is the use of insecticides by sod growers, lawn-care companies, and homeowners for control. Often the same active ingredients are available to all users at th e same time year round (personal observation). While current pesticide use by homeowners is not known, Lipsey (1980) conducted a survey in Florida and found that ho meowners used over 2,000,000 lbs of pesticides in a 12-month period during 1978-1979. Currently, pyrethroids, carbamates, neonicotinoids, and organophosphates are used for control in Florida. Carbamate (propoxur) and organophosphate (chlorpyrifos) re sistance was reported in the 1970s and 80s (Reinert and Niemczyk 1982, Reinert and Port ier 1983). Cross-resistance patterns and the stability of propoxur and chlorpyrifos resistance in are not known, making it unclear as to their effects on the current use of the carbamate, car baryl, and the organophosphate, trichlorfon. Meanwhile, sod growers, lawn-care companies, a nd homeowners continue to use the same active ingredients, and quite possibly increase the rate of insecticide resistance development in The problem of encroachment of on to neighboring St. Augustinegrass lawns may also be an important factor in the devel opment of resistance. Encroachment was observed in almost all of the lawns I collected from in 2006 and 2008 and the results of chapter 2 indicate that an individual lawn may represent a single population. If the latter is true, theoretical studies suggest that the evolution of insecticide resi stance may occur more rapidly in small, subdivided populations rather than large ones (Wright 1931, Crow and Kimura 1970, 139

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Roush and Daly 1990). Also, effects of imm igra tion on insecticide-resistant arthropods has been well discussed and show that immigration of susc eptible individuals into treated areas can slow resistance development by increasing the frequenc y of susceptible alleles in a treated population (Comins 1977, Georghiou and Taylor 1977a, Cur tis et al. 1978, Tayl or and Georghiou 1979, Tabashnik and Croft 1982, Roush and Daly 1990, Tabashnik 1990). Alternately, emigration of resistant individuals from treated areas speeds th e resistance development in the untreated area (Comins 1977). Sutherst and Comins (1979) indicat ed that acaricide-resis tant cattle ticks, (Acari: Ixodidae), in Au stralia were spread primarily by emigration. Immigration (or emigration) of between lawns is poorly understood. In addition, flight patterns ha ve not been evaluated in and so it is unclear how far they can travel in order to find a new food source. Methods to measure immigration/emigration and wing polymorphism warrants further investigatio n to understand their impact on resistance development in Cherry (2001a) documented that m acroptery is greatest in denser populations. The use of aggregation or se x pheromone traps for determining increases in macropterus for monitoring population increases in lawns would be useful. Traps and dye-marked may also be of use to investigat e movement patterns between lawns. Mark-release-recapture programs are frequently used for inve stigating animal populations (Southwood 1971) and dyes have been used for ma rking termites in studies involving foraging populations of termites (Lai 1977, Lai 1977 et al., Su and Scheffrahn 1988, Grace 1990, Jones 1990). Laboratory studies using test 5 rearing methods (chapter 3) could be used to facilitate studies in identifying aggregation and/or sex pherom ones. Aggregation pheromones have been identified for se veral heteroptera including Motschulsky and spp. (Hemiptera: Pentatomidae) (Is hiwatari 1976, Aldrich et al. 1991), 140

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(Dallas) and Stl (Hemiptera: Lygaeid ae) (Aller and Caldwell 2008), (Dallas) (Hemiptera: Core idae) (Leal et al. 1994), and Thunberg and (Westwood) (Hemiptera: Alydidae) (Numata et al. 1990, Ventura and Panizzi 2003). Also, sex pheromo nes have been found in heteroptera including (Green) and (Risso) (Hemiptera: Pseudococcidae) (Zada et al. 2004, Zhang et al. 2004, Zhang and Nie 2005, Zhang and Amalin 2005), and (Schrank) and (Rondani) (Hemiptera: Aphididae) (Campbell et al. 1990; Dawson et al. 1987, 1988, 1989, 1990). In addition to encroachment issues, Reinert (1 982b) speculated that th e tropical climate in south Florida, the high number of generations pe r year, all life stages being present each month of the year, and the monocultu re of St. Augustinegrass in residential lawns along Floridas southeastern coast may influence the development of insecticide resistance in He also reported that it was not unc ommon for lawns to be treated six to twelve times per year in some areas, with lawns possibly receiving less than recommended rates and exposing to sublethal doses each year (Reinert 1982b). These observations were made during the 1980s when insecticide resistance was almost exclus ively in the southeastern coast of Florida. The increase in housing development in Florida has helped incr ease the number of neighborhoods that are also a monoculture of St. Augustinegrass (personal observation). The results of chapter 2 and the data from Cherry and Nagata (2005, 2007) show that insecticide resistance is no longer restricted to south Florida and can be a pr oblem in all areas of the state (south, central, north) although to varying degrees of severity. Several factors can influence the selection of resistance to insecticid es in field populations of insects, including genetic (f requency and number of R alleles, dominance of R alleles, past 141

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selection by other chemicals), biol ogical (biotic, behavioral/ecol ogical), and operational factors (nature and persistence of insecticide, relationship to ea rlier used chemicals, number of applications, application methods ) (Georghiou and Taylor 1986). While the genetic factors are currently unknown, the development of a synchronous rearing method (test 5, Chapter 3) and the development of the airbrush bioassay and improved sprig dip bioassay (Chapter 4) will be useful tools for insecticide resistance studies in In chapter 3, test 5 proved to be the best method for synchronized rearing of For the first time, colonies of known age and genera tion can now be selected in the laboratory for bifenthrin resistance over multip le generations (or any other insecticide one chooses). Pure insecticide-susceptible and -res istant colonies can then be used in mode-ofinheritance studies and charac terization of mechanisms. Mo de-of-inheritance tests with susceptible and resistant males and virgin fema les would provide key information regarding the genetics of resistance in First, one could determine if resistance is dominant or recessive. If resistance in was shown to be dominant, a ro tation strategy as part of a resistance management program would be ineffectiv e. Second, one could determine if resistance is sex-linked or autosomal (Georghiou and Sa ito 1983). Comparison of XX and XY individuals of the F1 progeny could determine whether or not a sex chromosomal resistance factor had been involved in the parental R strain. This information could help to determine the rate of evolution of resistance and which management strate gies to use (Georghiou and Saito 1983). The rearing methods outlined in test 5 (Chapter 3) will be useful when characterizing mechanisms of resistance in because pure insecticide resistant and susceptible colonies can be developed. This is important because it may be difficult to distinguish if different responses (e.g., to a pesticide with or without a synerg ist) are due to a physiological 142

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resistance mechanism or the differences that can occur between differe nt populations (Scott 1990). In addition, the u se of pure colonies will allow for determining if a putative mechanism is really the one th at determines resistance (e.g., one that has been identified by bioassay in a resistant vs. susceptible insect). The expression of the mechanism in reciprocal F1 (resistant susceptible) progeny sh ould be correlated with the le vel of resistance seen in the bioassay (Scott 1990). There are other ways that the test 5 rearing me thod can be used. First, studies determining the number of eggs laid per labo ratory-reared female and mating habits would provide additional insight into the biology of (ie., how long to hold each generation for testing). Second, the effect of density on size and wing polymorphi sm in tests using laboratory colonies along with studies of field populations, woul d be of use in determining population dynamics in St. Augustinegrass lawns. Sasaki et al. (2002) found that environmental factors such as high temperat ure, long photoperiod, and crow ding during nymphal development stimulated the production and increase in Hidaka (Hemiptera: Lygaeidae). Third, as previously mentioned, st udies determining the presence of aggregation and/or sex pheromones would be useful for de velopment of monitoring techniques. Fourth, rearing studies could be conduc ted with natural enemies of Quality could be reared as a food or oviposition source and for use in studies with spp. as well as Last, it would be of benefit to determine the presence of gut symbionts in Various true bugs in the order Hemipter a contain large masses of bacteria in the alimentary tract and are thought to have a symbiotic association with them (Brues 1946). Forbes (1892) found the presence of bacteria in the alimentary canal of the common chinch bug, and later found them in other Hemiptera. Glasgow (1914) discovered that specific 143

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types of bacteria were charac teristic of each species he ex ami ned. Later, Kuskop (1924) suggested that bacteria are passe d from parent to offspring by entering the egg from a surface contamination at the time of oviposition, and reach the alimentary canal before hatching, accumulating in the caeca to which they are gene rally confined. If gut symbionts exist in there may be ways to alter the relationship and potentially lead to novel approaches to pest management. Several studies have demonstrat ed that when experimentally deprived of the symbiont, host stinkbugs suffer retarded growth and high mortality (Buchner 1965, Abe et al. 1995, Fukatsu and Hosokawa 2002, Hosokawa et al. 2006). Monitoring for resistance is considered essent ial to insecticide and acaricide resistance management (Dennehy and Granett 1984, Staetz 1985, Roush and Mill er 1986, Denholm 1990). In chapter 4, I developed an airbrush bioassay fo r testing contact and systemic insecticides and evaluated both the airbrush and the sprig-di p bioassay under more standardized conditions. These bioassays will be useful tools for detect ion and monitoring of insecticide resistance in Furthermore, I recommended that the sp rig-dip bioassay be used for detection of bifenthrin-resistant populations because it was more sensitive than the airbrush bioassay in detecting lower LC va lues. The airbrush bioassay would be better than the sprig-dip bioassay for use in studies concerning cross re sistance, mechanisms, mode-of-inheritance, and stability of pyrethroid (and other chemical classes) resistance in Future studies using the ai rbrush bioassay could include penetration studies, crossresistance patterns, insecticide synergists, enzyme assays, metabol ic detoxication, and target site sensitivity (Matsumura and Brown 1963; Plapp and Hoyer 1968; Scott and Georghiou 1986; Scott 1990; Bull and Patterson 1993; Sc harf et al. 1998a, 1998b, 1999, 2000a, 2000b, 2001; Scharf and Siegfried 1999; Wu et al. 1998; Li u and Yue 2000; Miota et al. 2000; Valles et al. 144

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2000; Ahmad et al. 2006). In addition, the airbru sh bioassay could be used for determ ining a diagnostic dose for pyrethroi dand imidacloprid-resistant populations because the results of chapter 4 demonstrate this bioassay re sults in less variance. Also, the airbrush bioassay has the added benefit of distinguishing differences between males and females (Chapter 4), and documenting behavioral differences could be of im portance when monitoring and documenting resistance in populations. In addition, I also addressed questions about sample size and duration of the airbrush and sprig-dip bioassays for response of to bifenthrin and imidacloprid (Chapter 4). Robertson et al. (1984) determined that a sample size of 120 was the minimum necessary for calculating a reliable LC50 estimation, but for increased precision sample sizes of 240 or more are often necessary (Robertson et al. 1984, 2007; Robertson and Preisler 1992). However, the authors noted that further investigation is needed to explore other combinations of sample size and dose placement in different bioassays (Robert son et al. 2007). Tabashnik et al. (1993) evaluated the duration of bioassays for (L.) (Lepidoptera: Plutellidae) against the microbial insecticide, Berliner. The authors demonstrated that bioassays for against could be run using shorter time intervals and a single concentration with little loss of inform ation compared to the standard bioassays (Tabashnik et al. 1993). The re sults of comparison tests and s ubsampled data in chapter 4 indicated that smaller sample si zes could be used when testing bifenthrin for a shorter duration compared to the sprig-dip bioassay. While sma ller sample sizes could be used for testing imidacloprid, the airbrush bioassa y required a similar duration to that of the sprig-dip bioassay. Future studies measuring wing polymorphism and the weight of the insects in each of the bioassays to determine differences in efficacy woul d be of value because size is not always an 145

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146 indicator for greater response to an insecticide. In additi on, studies should be conducted to determine if either bioassay closely mimics response of to insecticides under field conditions. Nonetheless, the development of the airbrush bioassay and synchronous rearing method provide valuable t ools that can be used to further investigate biology, population dynamics, response to in secticides, and how insecticide resistance develops in this pest. A greater understanding of how insecticide resist ance develops in will provide researchers, chemical companies, and lawn-car e companies the means to make responsible and sound decisions for resistance management of

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LIST OF REFERENCES Symbiont of brown-winged green bug, Scott. Jpn. J. Appl. Entomol. Zool. 39: 109-115. Delayed cuticular penetration and enhanced metabolism of deltamethrin in pyrethroid-resistant strains of from China and Pakistan. Pest Manage. Sci. 62: 805-810. Feeding responses of speci alist herbivores to plant extracts and pure allelochemicals: effects of prolonged exposure. Entomol. Exp. Appl. 111: 201-208. Identification and attractiveness of a ma jor pheromone component for nearctic spp. stink bugs (Heteroptera: Pentatomidae). Environ. Entomol. 20: 477-483. An investigation of the possible presence of an aggregation pheromone in the milkweed bugs, and Physiol. Entomol. 4: 287-290. Imidacloprid: Human health assessment and ecological risk assessment-fina l report. Syracuse Environ. Res. Assoc., Inc., New York, SERA TR 05-43-24-03a. Evaluation of cool and warm season grasses for resistance to multiple chinch bug species. M. S. University of Nebraska, NE. Introduction to the study of an imal populations. Phoenix Science Series, third impression. University of Chicago Press, Chicago and London. Floratam: a new disease-resistant St Augustinegrass. Circ. L-1146, Univ. TX Agric. Exp. Stn. Laboratory rearing of the hairy chinch bug. Environ. Entomol. 10: 226-229. A new species of Leptoglossus: a new and varieties. Bull. Brooklyn Entomol. Soc. 13: 35-39. Turfgrass: science and culture. Prentice-Hall, Englewood Cliffs, NJ. The role of turfgrasses in environmental protection and their benefits to humans. J. Environ. Qual. 23: 452-460. Management practices that reduce runoff transport of nutrients and pesticides fr om turfgrass, pp. 133-150. In M. T. Nett, M. J. Carroll, B. P. 147

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Horgan, and A. M. Petrovic (eds.), The fate of nutrients and pesticides in the urban environment. Am. Che m. Soc. Washington, D. C. Importance of host plant or diet on th e rearing of insects and mites, pp. 237-251. In T. E. Anderson and N. C. Leppla (eds.), Advances in insect rearing for research and pest management. Westview Press. Boulder, CO. Chinch bug control on St. Augustinegrass. Proc. Fla. State Hortic. Soc. 37: 216-219. Controlling chinch bugs with calcium cyanide. Univ. Fla. Agric. Exp. Stn. Bull. 362. 2 pp. Desensitization of fifth instar to azadirachtin and neem. Entomol. Exp. Appl. 81: 307-313. Handbook of turfgrass insect pests. Entomological Society of America, Lanham, MD. Home gardeners lawn insect control guide. Fla. Univ. Agric. Ext. Serv. Cir. No. 213. Improved detection of insecticide resistance through conventional and molecular tec hniques. Annu. Rev. Entomol. 32: 145-162. Influence of a short exposure to teflubenzuron residues on th e predation of thrips by (Acari: Phytoseiidae) and (Hemiptera: Anthocoridae). Pest Manag. Sci. 59: 1255-1259. Insect dietary. Harvard University Press. Cambridge, MA. Combined resistance in St. Augustinegrass to the southern chinch bug and the St. Augustinegrass decline strain of panicum mosaic virus. Plant Dis. 67: 171-172. Endosymbiosis of animals with plant microorganisms. Interscience, New York, NY. Characterization of pyrethroid resistance in a strain of the German cockroach (Dictyoptera: Blatte llidae). J. Econ. Entomol. 86: 20-25. Insect damage of lawns checked. Mississippi Farm Res. 21: 1-7. What is Floratam? Proc. Fla. State Hortic. Soc. 92: 228-232. 148

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Morphological identification of St. Augus tinegrass cultivars. Crop Sci. 26: 2832. Inheritance of host plant adaption in the southern chin ch bug (Hemiptera: Lygaeidae). Ann. Entomol. Soc. Am. 83: 563-567. Polyploid D. L. germplasm resistance to the polyploidy damaging population southern chinch bug (Hem iptera: Lygaeidae). Crop Sci. 30: 588593. Registration of FX-10 St. A ugustinegrass. Crop Sci. 33: 214-215. Genetic diversity and vulnerability of St. Augustinegrass. Crop Sci. 35: 322327. St. Augustinegrass, (Walt.) Kuntze, pp. 309-330. In M. D. Casler and R. R. Duncan (eds.), Bi ology, breeding, and genetics of turfgrasses. John Wiley and Sons, Inc., Hoboken, NJ. Southern chinch bug (Hemipte ra: Heteroptera: Lygaeidae) overcomes resistance in St. Augustineg rass. J. Econ. Entomol. 80: 608-611. Vulnerability of St. Augustin egrass to the southern chinch bug. Proc. Fla. State Hortic. Soc. 101: 132-135. Population outbreak of the southern chinch bug is regulated by fertilization. Int. Turf. Soc. Res. J. 7: 353-357. Resistance bioassay fr om southern chinch bug (Heteroptera: Lygaeidae) excret a. J. Econ. Entomol. 85: 2032-2038. Pest management attitudes and practices of Florida superintendents and lawn care professionals Fla. Turf Dige st. Sept./Oct.: 22-27. Recommended methods for measurement of pest resistance to pesticides. FAO Plant Production and Protection paper no. 21. FAO United Nations, Rome, Italy. Methods for measuring insectic ide susceptibility levels in bed-bugs, cone-nosed bugs, fleas and lice. Bull. WHO 24: 509-517. Insecticide resistance in : current status and implications for management. Proc. Brighton Crop Prot. Conf. 2B-3: 75-81. The sex attractant of the damson-hop aphid (Homoptera: Aphididae). J. Chem. Ecol. 16: 3455-3464. 149

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Some chinch bugs arent. W eeds Trees and Turf 7: 31-32. Variety evaluations in St. A ugustinegrass for resistance to the southern lawn chinch bug. Tex. Agric. Exp. Stn. Prog. Rep. PR-3374C. An example of cross-resistance to pyrethroids in DDT-resistant Pestic. Sci. 8: 618-624. Cabbage looper: effect of temperature on the toxicity of insecticides in the laboratory. J. Econ. Entomol. 66: 339-341. Chemistry of organophosphorus insecticides, pp. 913-917. In R. Krieger (ed.), Handbook of pesticide toxicology, 2nd ed. Academic Press, San Diego, CA. The insects-structure and functi on. English University Press, London. Seasonal wing polymorphism in southern chinch bugs (Hemiptera: Lygaeidae). Fla. Entomol. 84: 737-739. Spatial distribution of southern ch inch bugs (Hemiptera: Lygaeidae) in St. Augustinegrass. Fla. Entomol. 84: 151-153. Interrelationship of ants (Hymenoptera: Formicidae) and southern chinch bugs (Hemiptera: Lygaeidae) in Florida lawns. J. Entomol. Sci. 36: 411-415. Ovipositional preference and survival of southern chinch bugs ( Barber) on different grasses. Int. Turf. Soc. J. 8: 981-986. Development of resistance in southern chinch bugs (Hemiptera: Lygaeidae) to the insecticid e bifenthrin. Fla. Entomol. 88: 219-221. Resistance to two classes of insecticides in southern chinch bugs (Hemiptera: Lygaeidae). Fla. Entomol. 90: 431-434. Morphology and fertility of wing polymorphic adults of southern chinch bugs (Hemiptera: Lygaei dae). J. Entomol. Sci. 38: 688-691. Effects of larval size on predation by Fla. Entomol. 63: 146-151. Fundamentals of turfgrass management, 2nd ed. John Wiley and Sons, Inc., Hoboken, NJ. Choosing the right grass to fit the environment, pp. 99-113. In A. R. Leslie (ed.), Handbook of in tegrated pest management for turfgrass and ornamentals. Lewis P ublishers, Boca Raton, FL. 150

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Lawn and turf: management and environm ental issues of turfgrass pesticides, pp. 203-241. In R. Krieger (ed.), Handbook of pesticide toxicology, 2nd ed. Academic Press, San Diego, CA. Phosphorus in drainage water from sugarcane in the everglades agricultural area as affected by drainage rate. J. Environ. Qual. 23: 121-126. Insecticide resistance. In: J. M. Owens, M. K. Rust, and D. A. Reierson (eds.), Understanding and controlling the German cockroach. Oxford Univ. Press, Oxford. The development of insecticide resistan ce in the presence of immigration. J. Theor. Biol. 64: 177-197. Southern chinch bug, Barber (Heteroptera: Blissidae), management in St. Augustinegrass. M.S. th esis. University of Florida, Florida. Southern chinch bug control, 2002. Arthropod Management Tests, vol 29. Report: L13. Southern chinch bug control, 2004. Arthropod Management Tests, vol 31. Report: L9. The role of phosphorus in the eutr ophication of receiving waters: A review. J. Environ. Qual. 27: 261. Chemical control of southern chinch bug in St. Augustinegrass. Int. Turf. Soc. Res. J. 7: 358-365. Bioassay of St. Augustinegrass lines for resistance to southern chinch bugs (H emiptera: Lygaeidae) and to St. Augustinegrass decline virus. J. Econ. Entomol. 75: 515-516. St. Augustinegrass antibiosis to southern chin ch bug (Hemiptera: Lygaeidae) and to St. Augustine declin e strain of mosaic virus. J. Econ. Entomol. 82: 1729-1732. Local and regional resist ance to fenvalerate in Foerster (Homoptera: Psyllidae) in western North America. Can. Entomol. 121: 121-129. An introduction to population genetics theory. Harper and Row, New York, NY. Selection for and against insecticide resistance and possible methods of inhibiting the evolution of resistance in mosquitoes. Ecol. Entomol. 3: 273-287. 151

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Oviposition and longevity of chinch bugs on seedling growing in nutrient solution. J. Econ. Entomol. 40: 841-845. Association of thatch with populations of hairy chinch bug (Hemiptera: Lygaeidae) in turf. J. Econ. Entom ol. 83: 2370-2374. The aphid sex pheromone. Pure Appl. Chem. 61: 555-558. Identification of an aphid se x pheromone. Nature 325: 614-616. The sex pheromone of the greenbug, Entomol. Exp. Appl. 48: 91-93. Aphid semiochemicals: a review and recent advances on the sex pheromone. J. Chem. Ecol. 16: 3019-3030. Sex-related differences in the tolerance of Oriental fruit moth ( ) to organophosphate insecticides. Pest Manag. Sci. 57: 827-832. Monitoring and interpreting changes in in secticide resistance. Funct. Ecol. 4: 601-608. Monitoring dicofol-resistant spider mites (Acari: Tetranychidae) in California co tton. J. Econ. Entomol. 77: 1386-1392. Gahan as an egg parasite of the hairy chinch bug. J. Econ. Entomol. 30: 376. Physiological and growth responses of St. Augustinegrass cultivars to salinity. HortScience 28: 46-48. Carbamate insecticides, pp. 1087-1106. In R. Krieger (ed.), Handbook of pesticide toxicology, 2nd ed. Academic Press, San Diego, CA. Controlling chinch bugs on St. Augustine grass lawns. Auburn Uni. Agric. Exp. Stn. Prog. Rprt. No.7. Imidacloprid a new systemic insecticide. Pflanzenschu tz-Nachr. Bayer 44: 113-136. Resistance management with chloronicotinyl insecticides using imidacloprid as an example. Pflanzenschutz-Nachr. Bayer 49: 5-55. 152

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Capsuletransmitted gut symbiotic bacterium of the Japanese common plataspid stinkbug Appl. Environ. Microbiol. 68: 389-396. The magnitude of the resistance problem, pp. 14 43. In E. H. Glass et al. (eds.), Pesticide resistance: strategies and tactics for management. Nat. Acad. Press, Washington, D. C. A bioassay method and results of laboratory evaluation of insecticides against adult mosquitoes. Mosq. News 21: 328-337. Pest resistance to pesticides. Plenum Press, New York, NY. Genetic and biological infl uences in the evolution of insecticide resi stance. J. Econ. Entomol. 70: 319-323. Operational influences in the evolution of insecticide resistance. J. Econ. Entomol. 70: 653-658. Factors influencing the ev olution of resistance, pp. 157 169. In E. H. Glass et al. (eds.), Pesticide resistance: strate gies and tactics for management. Nat. Acad. Press, Washington, D. C. The gastric caeca and caecal bacteria of the Heteroptera. Biol. Bull. 26: 101-1701. Mark-recapture studies with (Isoptera: Rhinotermitidae). Sociobiology 16: 297-303. Effects of temperature on pyrethroid toxicity to colorado potato beetle (Coleopteran: Chrysomelidae). J. Econ. Entomol. 79: 588-591. Glycolipids as receptors for crystal toxin. Science 307: 922-925. Insensitive acetylcholinesterase as sites for resistance to organophosphates and carbamates in insects: insensitiv e acetylcholinesterase confers resistance in Lepidopter a, pp. 221-238. In I. Ishaaya (ed.), Biochemical sites of insecticide action and resistan ce. Springer-Verlag, Berlin. Southern branch insect detecti on, evaluation and prediction report 1983. Southeastern Branch of the Entomological Soci ety of America, vol 8. Entomol. Soc. Am. College Park, MD. Reproduction of the aphid related to the mineral nutrition of potato plants. Entomol. Exp. Appl. 13: 307-319. 154

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171 BIOGRAPHICAL SKETCH Julie Cara Congdon Vzquez was born in 1970 in St. Petersburg, Florida. After spending most of her childhood in Gainesville, she moved to Elma, Washington, and attended Elma High School. After enjoying 10 years in Washington, she returned to Gainesville to pursue a college degree. She enrolled at Santa Fe Community Co llege and developed an interest in entomology after taking several honors courses. In 1998, Cara entered the University of Florida as an undergraduate entomology major. While at the Un iversity of Florida, Cara gained practical experience in both pest control and research by working for the Florida Pest Control and Chemical Company, the University of Florid as Entomology and Nematology Department (urban entomology labora tory), and United States Department of Agriculture (USDA). Cara received her bachelors degree in ento mology and nematology with a specialization in urban pest management in May 2001. Afterwar ds, she was hired by the FMC Corporation as a summer intern and provided technical and sales s upport to golf course superintendents, pest management professionals, and distributors. Cara started her graduate studies in August 2001 at the University of Florida under the guidance of Dr. Eileen A. Buss. During her masters research Cara developed a fondness for southern chinch bugs. She completed her Master of Science degree in May 2004 and immediately started work on her Ph.D. She is a member of the Entomological Society of America, Entomology and Nematology Student Organization (ENSO), Florida Entomological Society, Florida Turfgr ass Association, Gamma Sigma Delta, and the Urban Entomological Society (UES). After comp leting her dissertation, Cara will work as a Research Scientist for Scynexis, Inc., in the Research Triangle Park, NC.