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Southern Chinch Bug, Blissus insularis Barber (Heteroptera: Blissidae), Management in St. Augustinegrass


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SOUTHERN CHINCH BUG, Blissus insularis BARBER (HETEROPTERA: BLISSIDAE), MANAGEMENT IN ST. AUGUSTINEGRASS By JULIE CARA CONGDON A THESIS PRESENTED TO THE GRADUATE SCHOOL OF THE UNIVERSITY OF FLORIDA IN PARTIAL FULFILLMENT OF THE REQUIREMENTS FOR THE DEGREE OF MASTER OF SCIENCE UNIVERSITY OF FLORIDA 2004

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Copyright 2004 by Julie Cara Congdon

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This thesis is dedicated to an extraordinary woman, my grandmother, Margaret Congdon.

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iv ACKNOWLEDGMENTS I am deeply grateful to my major advisor, Dr. Eileen Buss, for her guidance, support, and encouragement while obtaining this degree. I also thank the members of my committee, Dr. Laurie Trenholm and Dr. Philip Koehler, for their valuable advice, expertise, and review of this thesis. I thank Dr. Ronald Cherry for his technical advice and the use of the blower/vacuum. I also thank Drs. Kris Braman, Frederick Baxendale, and Grady Miller, for their advice on my research. Special thanks go to Dr. William Crow and his wife Laurel for “donating” their lawn to science. I thank Dr. Jerry Butler and Dr. Oscar Liburd for the use of laboratory supplies. I acknowledge and thank Dr. Susan Halbert and Julieta Brambila from the Division of Plant Industry for their assistance in obtaining literature and specimen identifications. This research could not have been completed without the assistance of several individuals. I am grateful to Paul Ruppert, Lois Wood, Mike Wang, Kathryn Barbara, and Jay Cee Turner for their help with data collection. Special thanks go to Brian Owens and the employees at the G.C. Horn Memorial Turfgrass Field Laboratory for their advice, use of supplies, and assistance with field projects. I thank Gil Marshall for his patience and assistance with formula calculations. I also thank Jan Weinbrecht and Jason Haugh for their advice, use of equipment, and assistance during the irrigation experiment. Many thanks go to Becky Griffin, the Homeowners of Hickory Forest, and Colonnades On Top of the World, for helping me obtain B. insularis for this research. I acknowledge

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v and thank the Bayer Corporation, Entomos, FMC Corporation, Syngenta Crop Protection, Inc., and Southern Agricultural Insecticides, Inc., for donating products for this research. I acknowledge and thank Lyle Buss for his assistance with photography. I thank Matthew Aubuchon for assistance with SAS programming. Many thanks go to the faculty, students, and staff of the Entomology and Nematology Department for their advice, guidance, and friendship. Special thanks are given to my family and friends for all of their love and support. I am especially thankful to my fianc, Ricky Vazquez, for his assistance in data collection, statistical analysis, editing of this thesis, and in his confidence that I would succeed in obtaining this degree.

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vi TABLE OF CONTENTS page ACKNOWLEDGMENTS.................................................................................................iv LIST OF TABLES...........................................................................................................viii LIST OF FIGURES...........................................................................................................ix ABSTRACT....................................................................................................................... .x CHAPTER 1 LITERATURE REVIEW.............................................................................................1 Chinch Bugs..................................................................................................................2 Management Practices..................................................................................................5 Chemical Control...................................................................................................5 Biological Control.................................................................................................5 Big-eyed bugs.................................................................................................6 Geocoris punctipes .........................................................................................7 Host Plant Resistance............................................................................................8 Cultural Control.....................................................................................................8 Objectives..................................................................................................................... 9 2 SAMPLING METHODS FOR THE SOUTHERN CHINCH BUG, Blissus insularis BARBER (HETEROPTERA: BLISSIDAE)..............................................................10 Introduction.................................................................................................................10 Materials and Methods...............................................................................................11 Results and Discussion...............................................................................................13 3 SOUTHERN CHINCH BUG, Blissus insularis (HETEROPTERA: BLISSIDAE), INTEGRATED PEST MANAGEMENT (IPM)........................................................18 Introduction.................................................................................................................18 Materials and Methods...............................................................................................19 Insect Collection and Colony Maintenance.........................................................19 Host Plant Resistance..........................................................................................20 Predation Assay...................................................................................................20 Insecticidal Control.............................................................................................22

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vii Results and Discussion...............................................................................................24 4 ST. AUGUSTINEGRASS GROWTH RESPONSE TO THREE LEVELS OF IRRIGATION AND Blissus insularis DENSITY......................................................32 Introduction.................................................................................................................32 Materials and Methods...............................................................................................33 Insect Collection and Maintenance.....................................................................33 Results and Discussion...............................................................................................35 LIST OF REFERENCES...................................................................................................40 BIOGRAPHICAL SKETCH.............................................................................................47

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viii LIST OF TABLES Table page 2-1. Mean ( SEM) number and percentage of Blissus insularis recovered from 15.2 cm pots of ‘Palmetto’ St. Augustinegrass......................................................................17 3-1. Mean percent mortality of Blissus insularis ( SEM) after feeding on Floratine or FHSA 115 St. Augustinegrass...............................................................................27 3-2. Mean percent mortality of Blissus insularis ( SEM) killed by different rates of Bioblitz and diazinon at 2 and 4 days after treatment..............................................28 3-3. Mean percent survival of Blissus insularis ( SEM) at 1 and 4 wk post treatment..29 3-4. Mean percent survival of Blissus insularis ( SEM) at 1 and 4 wk post treatment..30 4-1. Mean dry weight (mg) ( SEM) of grass clippings by week....................................38 4-2. Mean root weight ( SEM) of St. Augustinegrass after 8 wk...................................39

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ix LIST OF FIGURES Figure page 2-1. St. Augustinegrass grown in 15.2 cm diameter clay pot enclosed in a chiffon mesh cage........................................................................................................................... 16 2-2. Equipment used to recover Blissus insularis from test plants...................................16 3-1. Methods used in pesticide field trials........................................................................31 4-1. Lysimeter used in experiment....................................................................................37

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x Abstract of Thesis Presented to the Graduate School of the Universi t y of F l orida in Pa r ti a l F u l f illm e nt o f the Requirements for the Degree of Master of Science SOUTHERN CHINCH BUG, Blissus insularis BARBER, (HETEROPTERA: BLISSIDAE), MANAGEMENT IN ST. AUGUSTINEGRASS By Julie Cara Congdon May 2004 Chair: Eileen A. Buss Major Department: Entomology and Nematology The southern chinch bug, Blissus insularis Barber, is the most destructive insect pest of St. Augustinegrass ( Stenotaphrum secundatum [Walt.] Kuntze). The purpose of this research was to examine the effectiveness of different components of an IPM program for B. insularis including sampling techniques, host plant resistance, biological, chemical, and cultural controls. The optimal sampling method for B. insularis in a greenhouse experiment was flotation, compared to using a hand-held vacuum or berlese funnel. The large blower/vacuum was nearly as accurate at extracting B. insularis so it was used to collect field populations for use in experiments and flotation was used to recover insects from test plants in the laboratory. In a no-choice test, more than 62% of the B. insularis successfully survived on both Floratine and FHSA-115. During the 3 wk experiment feeding, mating, and molting were observed. The different life stages of B. insularis had similar mortality after feeding on either variety, suggesting that neither variety was

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xi resistant to B. insularis Fifth instar and adult Geocoris punctipes (Say) consumed a similar number of B. insularis When pooled together, fifth instar and adult G. punctipes consumed significantly more first-third instar (57%) B. insularis than fourth-fifth instar or adult stages (17.2 and 16.0% respectively). This study evaluated the efficacy of professional and non-professional (homeowner) products. In the laboratory insecticide assay, diazinon (100 ppm) killed significantly more B. insularis adults on sprigs than any other diazinon or Bioblitz treatment 2 and 4 days after treatment. Homeowner products containing bifenthrin, carbaryl, deltamethrin, and -cyhalothrin achieved over 80% control 1 wk post-treatment in the field test. All of the professional products were statistically different from the control 1 wk after treatment, although carbaryl was the only treatment to kill all B. insularis at 1 wk. Stunted growth from B insularis feeding was observed from grass clippings collected after 1 wk. Dry weights of grass clippings collected from the different B. insularis densities were different from one another except during weeks two, six, and seven. During this time, the dry weights of grass clippings recovered from treatments containing 200 B. insularis were lower than dry weights of grass clippings collected from treatments containing 0 or 30 B. insularis Blissus insularis feeding in densities of 30 and 200 lowered root weight. At five weeks, dry weight of grass clippings was lowest in treatments with low irrigation. At week seven, dry weights of grass clippings collected from treatments with low irrigation were lower than clippings from treatments with high irrigation levels. Irrigation did not impact root growth. Irrigation and B. insularis density did not interact to affect plant growth.

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1 CHAPTER 1 LITERATURE REVIEW Turfgrass production and maintenance are valuable parts of the Florida economy (Hodges et al. 1994). In 1992, residential lawns accounted for 75% of the 4.4 million acres of turfgrass used and maintained throughout the state. Consumers spent $5 billion on turfgrass maintenance or roughly $1,200 per acre. Sales from products and services by turfgrass producers and commercial distributors totaled $6.5 billion. The demand for high quality turfgrass is growing with Florida’s increasing annual population of almost 2% (U.S. Census Bureau 2003). Common grasses used in Florida include bahiagrass ( Paspalum notatum Flugge), bermudagrass [ Cynodon dactylon ( L.) Pers], centipedegrass [ Eremochloa ophiuroides (Munro) Hack], seashore paspalum ( Paspalum vaginatum Swartz), and zoysiagrass ( zoysia spp). However, of the >6.3 million residences in Florida (U.S. Census Bureau 2000), the most widely used grass is St. Augustinegrass ( Stenotaphrum secundatum [Walt.] Kuntze). St. Augustinegrass is a warm-season, coarse-textured, aggressive, and stoloniferous grass that is widely used in lawns in the coastal regions of the United States (Turgeon 1996). Most St. Augustinegrass cultivars have good salt and shade tolerance and establish easily from sprigs or sod. Its aggressive growth habit gives it good recuperative capability, but it is prone to thatch buildup (Potter 1998). Cultivars of St. Augustinegrass that are commonly marketed include ‘Floratam’, ‘Palmetto’, ‘Delmar’, ‘Floratine’, and ‘Seville’ (White and Busey 1987, McCarty and Cisar 1997, Trenholm et al. 2000).

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2 Several insect pests attack St. Augustinegrass in Florida including tropical sod webworms [ Herpetogramma phaeopteralis (Guenee)], armyworms ( Spodoptera spp.), grass loopers ( Mocis spp.), mole crickets ( Scapteriscus spp.), rhodesgrass mealybugs [ Antonina graminis (Maskell)], and southern chinch bugs ( Blissus insularis Barber). Sod webworms feed primarily at night. Newly hatched larvae skeletonize the grass blades, while older larvae chew on grass blades near the soil surface (Buss and Caldwell 2001). Armyworms first skeletonize the grass blades and later create bare spots. Mature larvae feed during the day and night (Buss 2001). Grass loopers chew on the grass blades, resulting in ragged, yellow to brown patches of grass (Sprenkel 2003). Mole crickets damage turfgrass by their tunneling, which uproots and dries out grass plants. Feeding from rhodesgrass mealybugs discolors grass and heavy infestations give the appearance that excess fertilizer has ‘caked’ around the grass nodes (Buss 2001). The southern chinch bug is the most destructive insect pest of St. Augustinegrass and can cause extensive damage or kill entire lawns (Reinert and Kerr 1973, Bruton et al. 1983). Chinch Bugs Many Blissus spp. are important pests that attack plants in the family Graminae. Blissus spp. feed on the phloem and xylem near the base of the plant where the grass meristem occurs (Painter 1928). Salivary sheaths are deposited along feeding tracks within the tissues, which is characteristic of feeding on phloem tissue bundles (Backus et al. 1988). Their feeding can cause wilting, chlorosis, stunting, and death through clogging of vascular transport systems (Painter 1928, Negron and Riley 1990, Spike et al. 1991). In particular, B. insularis prefer open sunny areas of St. Augustinegrass, especially areas with abundant thatch (Reinert and Kerr 1973), where they suck fluids from the crown and stem of grasses with their needle-like mouthparts. As the grass dies,

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3 the insects feed at the edge of the dead spots, thus enlarging the damaged area. Blissus insularis activity begins between March and April (Kerr 1966) and continues until October. Damage is usually first noticed in drought-stressed areas along sidewalks, pavements, or in poorly irrigated areas. At least 15 Blissus species occur in the United States and Canada; however, only four are considered damaging to turfgrass, including the buffalograss chinch bug, ( B. occiduus Barber); common chinch bug, [ B. leucopterus leucopterus (Say)]; hairy chinch bug, ( B. l. hirtus Montandon); and southern chinch bug, ( B. insularis ) (Brandenburg and Villani 1995, Sweet 2000). Blissus occiduus is a serious pest of buffalograss in Arizona, California, Colorado, Kansas, Montana, Nebraska, New Mexico, Oklahoma, and parts of Canada (Baxendale et al. 1999, Vittum et al. 1999, Carstens 2003). Blissus leucopterus leucopterus has caused severe damage to wheat, spelt, corn, millet, pearl millet, rye, barley, sorghum, grain sorghum, rice, and Sudan grass (Sweet 2000). Blissus leucopterus leucopterus can be found in Colorado, the Great Plains, New Mexico, and Nebraska eastward to the Atlantic Ocean (Webster 1907, Leonard 1966, Khuhro 1994). Blissus leucopterus hirtus is a major pest of cool-season grasses throughout the Northeast, parts of the Midwest, into the mid-Atlantic states as far south as Virginia, and west to Minnesota (Vittum et al. 1999). Blissus insularis occurs in Florida, Alabama, Georgia, Louisiana, Mississippi, North and South Carolina, Texas, California, and Mexico (Henry and Froeschner 1988, Sweet 2000). Adult B. insularis are small and somewhat elongate insects measuring about 3 mm long and 1 mm wide (Leonard 1968). Adult females are slightly larger than males. The head is usually narrower than the posterior margin of the pronotum (Miller 1971). Wings

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4 are white with a distinctive triangular-shaped 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 long-winged forms (macropterous), short-winged forms (brachypterous), or both (Webster 1907). In Florida, macroptery is greatest during the summer and fall (Cherry 2001a). Young nymphs are as small as 1.0 mm, are reddishorange with a white band across the dorsal side of the abdomen, and become black in color as they mature. Eggs are laid singly or a few at a time in sheaths, soft soil, or in other protected areas. The eggs are white when first laid but turn bright orange just before hatching. Development from egg to adult requires about 13 wk at 21 C and 5 wk at 28 C (Potter 1998). All life stages are present year-round 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). Blissus insularis move between lawns mainly by walking and large numbers have been observed crawling across sidewalks and driveways bordering heavily infested lawns (Kerr 1966). All life stages are distributed vertically through the turf thatch and into the upper organic layer of the soil. Densities of 500-1,000 B. insularis /0.1 m2 are common and infestations of more than 2,000/0.1 m2 have been reported (Reinert and Kerr 1973). Light to moderate infestations are extremely aggregated in small areas in the lawn rather than evenly distributed (Cherry 2001b). Blissus insularis also attack other lawn grasses including bahiagrass, bermudagrass, centipedegrass, and zoysiagrass, but most of the injury to these has occurred near heavily infested St. Augustinegrass (Kerr 1966). Other hosts include crabgrass, torpedograss, and Pangolagrass (Slater and Baranowski 1990, Brandenburg and Villani 1995).

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5 Management Practices Chemical Control Control of B. insularis has historically been achieved using insecticides such as tobacco dust, calcium cyanide, nicotine sulfate, DDT, parathion, dieldrin, aldrin, chlordane, chlorpyrifos, and diazinon (Watson and Bratley 1929a and b; Kelsheimer 1952, Wolfenbarger 1953, Kerr 1956, Brogdon and Kerr 1961). Up to six insecticidal applications per year may be needed on Florida lawns to control damaging populations of B. insularis (Reinert 1978). With such reliance on chemical use, this insect has developed resistance to organophosphates and organochlorines (Kerr 1958, 1961; Reinert & Niemczyk 1982; Reinert 1982; Reinert and Portier 1983). Currently, pyrethroids are most commonly available to homeowners and turfgrass professionals for controlling B. insularis outbreaks. A lack of rotation with other chemical formulations increases the risk of B. insularis developing resistance to pyrethroids as well. Some alternatives to chemicals are biological control, host-plant resistance, and cultural control. Biological Control Parasites, predators, and pathogens are found in association with Blissus spp. The parasitic wasp, Eumicrosoma benefica Gahan (Hymenoptera: Scelionidae), attacks the eggs of B. leucopterus (Say) in Kansas, reportedly reducing ‘broods’ of chinch bugs by as much as 50% (McColloch and Yuasa 1914). Eumicrosoma benefica also parasitized B. insularis eggs in Florida and field tests showed an average abundance of 34.5 wasps/0.1 m2 in lawns containing B. insularis populations of 90.0/0.1 m2 (Reinert 1972a). Along with E. benefica predatory insects [e.g., Pagasa pallipes Stal (Heteroptera: Nabidae), Xylocoris vicarius (Reuter) (Heteroptera: Anthocoridae), Lasiochilus pallidulus Reuter (Heteroptera: Anthocoridae), Sinea spp. (Heteroptera: Reduviidae), Labidura riparia

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6 Pallas, Geocoris bullatus (Say) and Geocoris uliginosus (Say) (Heteroptera: Geocoridae)] were observed feeding on B. insularis in Florida (Reinert 1978). The native fire ant, Solenopsis geminata (F.), and a spider, Lycosa sp., were also observed attacking B. insularis (Reinert 1978). However, the red imported fire ant, Solenopsis invicta Buren, was unable to suppress B. insularis populations (Cherry 2001c). Beauveria globulifera (Spegazzini) Picard and Metarhizium anisopliae (Metch.) Sorokin were unsuccessful in controlling B. insularis in field tests (Kerr 1958). Beauveria bassiana (Balsamo) Vuillemin was pathogenic on all life stages of B. insularis ; however, success appeared limited to high humidity, B. insularis populations, and moisture levels (Reinert 1978). Big-eyed bugs Big-eyed bugs Geocoris spp. are very abundant and widely distributed (Readio and Sweet 1982). These generalist predators attack a wide range of prey species, including various insect and mite pests of agricultural crops such as cotton, soybean, strawberry, peanut, turfgrass, vegetable, sugarbeet, alfalfa, and tobacco (McGregor and McDonough 1917, Knowlton 1935, York 1944, Champlain and Sholdt 1967a and b, Mead 1972, Crocker and Whitcomb 1980). They specifically have been observed attacking aphids, plant bugs, eggs and larvae of the bollworm [ Heliothis zea (Boddie)], flea beetles ( Phyllotreta spp.), false chinch bugs [ Nysius ericae (Schilling)], southern green stink bugs [ Nezara viridula (L.)], and green peach aphids [ Myzus persicae (Sulzer)] (Bell and Whitcomb 1964, Crocker and Whitcomb 1980). Geocoris spp. will feed on eggs, immatures, or adults of relatively small or passive prey. They attack by inserting their proboscis into the head, thorax, or abdomen and tend to walk about with the prey suspended on the labium (Sweet 2000).

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7 Although Geocoris spp. are predators, they also require water from plant sources (York 1944). In laboratory colonies, Geocoris spp. can temporarily be maintained on plant food alone, but prey are required for proper development and fecundity (Stoner 1970). Geocoris bullatus once thought to be a turfgrass pest in Connecticut, feeds on lawn insects such as B. l. hirtus (Dunbar 1971). Geocoris uliginosus had densities as high as 16.0/0.1 m2 when high B. insularis populations were present and fed on an average of 9.6 3.3 (mean SEM) nymphs in 5 d under laboratory conditions (Reinert 1978). The most common species in Florida are G. uliginosus G. bullatus and G. punctipes (Say) (Mead 1972). Geocoris punctipes Geocoris punctipes are small oblong Lygaeids having the head broader than long, large prominent eyes that curve backward and overlap the front of the pronotum, and tylus that has a longitudinal groove (Mead 1972). They have relatively short antennae that are slightly enlarged at the tip. Geocoris punctipes adults are about 0.5 cm long and are silver-gray in color. Nymphs resemble adults but lack fully developed wings. Cigarshaped eggs are white to tan in color with a distinctive red spot and are laid singly on leaves and stems of many crops. Eggs hatch in approximately 1 wk. Geocoris spp. have five instars, each of which lasts from 4 to 6 d. Adults live for about 1 mo and a female can lay up to 300 eggs during her lifetime. Several generations may occur throughout the season. Adult and immature G. punctipes feed by sucking juices from their prey through needle-like mouthparts. They are primarily active in the morning (Dumas et al. 1962). These insects are often confused for Blissus spp. Geocoris spp. are commonly found throughout the southern United States.

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8 Host Plant Resistance Host resistant grasses are used to suppress pest damage. There are three types of resistance that can occur: antibiosis, antixenosis, or tolerance. Antibiosis affects the biology of the pest insect and often results in increased mortality, or reduced oviposition and longevity. With antixenosis, the pest shows non-preference for a resistant grass compared to a susceptible one. When a host resistant grass shows tolerance, the grass is able to withstand or recover from damage caused by an insect. Floratam was released as a chinch bug resistant cultivar in 1973 by the University of Florida and Texas A&M. Floratam is the most widely produced and used cultivar of St. Augustinegrass and accounts for 80% of sod production in Florida (Haydu et al. 1998). It has a very coarse leaf texture, poor cold and shade tolerance, is resistant to the St. Augustine decline virus, and successfully minimized B. insularis problems for years (Busey 1979, Trenholm et al. 2000). The mode of resistance was antibiosis (Reinert and Dudeck 1974). However, B. insularis can now survive and reproduce on this variety (Busey and Center 1987; Busey 1990a and b). Research is being conducted to develop new cultivars of St. Augustinegrass that are resistant to B. insularis Cultural Control Cultural practices may influence the susceptibility of St. Augustinegrass to B. insularis Over-fertilizing, over-watering, and improper mowing can cause lawn grasses to develop a thick thatch layer (Trenholm et al. 2000). Thatch is a layer of accumulated dead leaf blades, stolons, and roots between the live plant and soil. Accumulation of thatch often makes the lawn more susceptible to pests (Trenholm et al. 2001), possibly by protecting them from predation and environmental stress. The abundance of B. l. hirtus was found to be closely linked to thatch thickness in lawns (Davis and Smitley 1990).

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9 Excessive thatch (exceeding 2.5 cm) may need to be professionally removed by vertical mowing (Trenholm et al. 2000). Heavy fertilization may cause increased growth, thatch, susceptibility to pests such as B. insularis and reduced tolerance to environmental stresses (Kerr 1966, Busey and Snyder 1993, Trenholm et al. 2001). Excess irrigation may lead to problems such as a shallow root system, increased thatch, and increased pest problems (Trenholm et al. 2003). Reduced fertilization and proper irrigation practices may help to reduce B. insularis problems (Potter 1998, Sweet 2000). St. Augustinegrass should be mowed to a height of 8 10 cm (Trenholm et al. 2000). Proper cultural practices promote healthy grass, which may be able to better tolerate chinch bug damage. Objectives An integrated pest management program (IPM) is needed for the southern chinch bug to reduce the turfgrass industry’s sole reliance on pesticides. The purpose of this study was to evaluate the effectiveness of different components of an IPM program for B. insularis including: 1) sampling techniques, 2) the potential of ‘FHSA-115’ as a chinch bug resistant variety of St. Augustinegrass, 3) B. insularis predation by G. punctipes 4) several homeowner, professional, and experimental insecticides labeled for chinch bug control, and 5) St. Augustinegrass growth response to three levels of irrigation and B. insularis density.

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10 CHAPTER 2 SAMPLING METHODS FOR THE SOUTHERN CHINCH BUG, Blissus insularis BARBER (HETEROPTERA: BLISSIDAE) Introduction A successful integrated pest management (IPM) program largely depends on the development of an effective and efficient sampling technique (Hutchins 1994). The proper use of such techniques provides a means by which to make informed decisions, such as establishing more accurate economic and/or aesthetic thresholds (Southwood 1978), evaluating product trials, and designing innovative monitoring or management strategies. However, the researcher must have an understanding of the biology of the insect population and environmental factors (i.e. temperature, humidity, light), which may affect activity periods of the insect and critical thresholds (Horn 1988). Effective sampling also depends on insect distribution and behavior (Trmla 1982, Standon 2000). Insects with clumped distributions that live in dense plant material, such as the southern chinch bug, Blissus insularis may be somewhat difficult to accurately sample (Cherry 2001b). Blissus insularis is one of the most important pests of St. Augustinegrass (Kerr 1966, Crocker 1993, Cherry and Nagata 1997). Infestations usually occur in open, sunny, and drought-stressed areas near sidewalks and driveways. Nymphs and adults live in the thatch and suck fluids from the grass with their needle-like mouthparts, resulting in yellow to brown patches in the lawn. Newly hatched B. insularis nymphs often wedge themselves in between the grass blades at the nodes to feed (Kerr

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11 1966), making it difficult to recover them without physically pulling the blades back to expose the insects. Techniques used to extract turfgrass insects include flotation, irritant sampling, and vacuuming. Although some studies have examined the effectiveness of various techniques used in sampling turfgrass pests, including chinch bugs (Short & Koehler 1979, Majeau et al. 2000), vacuum extraction was not included. Our objective for this study was to determine the most accurate and efficient method for specifically extracting B. insularis from St. Augustinegrass. Materials and Methods Blissus insularis were mass-collected from St. Augustinegrass lawns in Gainesville, FL (Alachua Co.), using a modified blower/vacuum (Electrolux Home Products, Augusta, GA) and transported in a mesh-covered bucket to the laboratory. Insects were provided fresh cuttings of Palmetto St. Augustinegrass as needed and maintained in the laboratory <10 d until being aspirated into plastic vials (2.5 10.2 cm) containing a coneshaped, moistened, 70 mm Whatman filter paper to prevent being damaged from aspiration. Vials were closed using a foam cap to prevent insect escape. Twenty 15.2 cm diameter clay pots of Palmetto St. Augustinegrass were established in Arrendondo fine sand (loamy, siliceous, hypothermic, Grossarenic Paleudalt) and maintained under ambient conditions in a greenhouse at the University of Florida (Gainesville, FL). Plants were fertilized every 2 wk and watered as needed. The thatch layer was <0.6 cm. Plants were maintained at a 7.6 cm height. Twenty fourthfifth instars and five adult B. insularis were transferred from vials using a camel-hair brush into the center of each pot and caged on 13 May 2003 (Fig. 2-1). Insects were maintained in caged pots for 7 d. Plants were arranged in a complete randomized design

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12 (CRD). After 7 d, four sampling methods (flotation; large, gas powered vacuum; small, battery operated vacuum; or berlese funnel) were used to extract the insects (n = 5 pots per treatment). The first method involved floating B. insularis from the plant material. Blissus insularis were forced from plant debris using a modified version of the flotation technique used in extracting Solenopsis invicta colonies from soil (Banks et al. 1981). Potted plants were placed into 19 liter buckets, water was added to each bucket with 1.3 cm diameter PVC tubing, brass connectors, and 7.6 cm long 0.6 cm diameter rubber tubing (Fig. 2-2 A). Warm water (~38 C) slowly filled up to the edge of each pot, was reduced to a fast drip, and was turned off after about half of the grass blades were submerged. Insects were collected as they crawled up the grass blades, were placed into vials of 80% EtOH, and counted. Pots were submerged for ~2 h. The second method used was a large, gas-powered blower/vacuum (Electrolux Home Products, Augusta, GA) equipped with a 0.8 m long hose attachment (Fig. 2-2 B). Insects were collected into a 12.7 cm diameter knitting-ring with a chiffon mesh that covered the vacuum attachment’s intake hose. Each pot of grass was vacuumed for 1 min, samples were placed into plastic bags, and the number of chinch bugs collected were counted in the laboratory. A smaller, hand-held, modified Black & Decker Dust-Buster vacuum (Bioquip Products, Rancho Dominguez, CA) equipped with a 12.7 cm long 3.8 cm diameter hose attached to a removable collecting chamber (12.7 cm long 5.1 cm diameter) was used as the third sampling method (Fig. 2-2 C). The vacuum was powered by a portable, 12-

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13 volt DC battery pack (Bioquip Products, Rancho Dominguez, CA). Each pot was vacuumed as described above. The last method involved placing infested plants into berlese funnels using 40 wattage bulbs. Potted plants were placed sideways into the berlese funnels and were left in the pots to minimize debris falling into the collection containers (Fig. 2-2 D). It was assumed that any living chinch bugs would fall into the funnel and container of 80% EtOH as the soil and plants dried within 48 h. The total number of B. insularis collected using each method was counted. The mean number of chinch bugs recovered was analyzed using an analysis of variance (ANOVA, P 0.05) and treatment means were compared using the Tukey-Kramer HSD multiple comparison test (Jmp SAS Institute Inc. 2001). Results and Discussion Flotation was the most effective technique for extracting a known number of B. insularis from pots of grass (Table 2-1). Different versions of this method are commonly used in B. insularis experiments. Under field conditions, a large metal cylinder is placed 3–5 cm into the soil near damaged turfgrass, filled with water, and any chinch bugs present float to the surface within 3–10 min (Kerr 1966; Reinert 1972b, 1982). For laboratory or greenhouse studies, cores or pots of grass are submerged in buckets of water, insects crawl up the grass blades, and are collected (Nagata and Cherry 1999, Richmond and Shetlar 2000). In this experiment, B. insularis were forced to escape slowly increasing water levels and could float on the water if they fell off the plant material. Even if completely submerged for considerable periods ( 4 h), chinch bugs were expected to survive (Britcher 1903, Janes et al. 1935). However, this method did

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14 not recover all of the insects. Some mortality may have occurred during the experiment or some insects may have remained in the plant material. Because soil particles were heavy and fell to the bottom of the bucket, very little debris interfered with observation of B. insularis movement on the plants or water. Although flotation was somewhat laborintensive, it extracted the highest number of B. insularis and was the only method where variability among treatments was low. The number of B. insularis recovered by flotation compared to the large blower/vacuum was not statistically significant (22.0 and 15.2 respectively, Table 2-1). Vacuums similar to the one used in this experiment have successfully been used for collecting chinch bugs for laboratory assays and from field insecticide plots (Crocker and Simpson 1981, Crocker 1993, Nagata and Cherry 1999). Standen (2000) compared the effectiveness of pitfall traps and a D-vac suction trap combined with a lightweight swish net and reported the highest number of Hemipterans being collected in D-vac samples. Several different vacuums (whereby insects are sucked into a layer of fabric that is stretched across an intake hose and released into a different container) or D-vac machines are used to collect insects (Crocker and Simpson 1981, Crocker 1993, Nagata and Cherry 1999, Cherry 2001b). Vacuum extraction of insects may be faster and at least as effective as the flotation method (Crocker 1993). The number of B. insularis recovered by the large blower/vacuum compared to the smaller, hand-held vacuum was not significant (Table 2-1). However, the smaller handheld vacuum recovered significantly fewer B. insularis than the flotation method (10.0 and 3.4 respectively). Additional sampling time may be required for the small, hand-held vacuum to be effective in recovering known numbers of B. insularis However, for its

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15 convenient size, the hand-held vacuum might work well for rapid detection of chinch bugs in infested lawns. For the berlese funnel method, soil or grass samples may be placed into funnels and as actively moving insects escape the light and heat of lowwattage bulbs, they drop into containers of alcohol (Niemczyk et al. 1992, Heng-Moss et al. 2002). The lowest recovery of B. insularis occurred with the berlese funnel (Table 21). Additional testing and improved methods (i.e. different bulb wattage, plant arrangement, funnel design) for the berlese funnel technique may improve its effectiveness in recovering B. insularis As part of developing an IPM program, several of these techniques would be useful under the proper circumstances. The flotation method and both vacuums tested in this study could be used to detect the presence of B. insularis in the lawn. However, when conducting experiments using a known number of B. insularis the flotation method or large blower/vacuum may be best to use since these techniques recovered the highest number of insects from infested pots of St. Augustinegrass.

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16 Figure 2-1. St. Augustinegrass grown in 15.2 cm diameter clay pot enclosed in a chiffon mesh cage. Figure 2-2. Equipment used to recover Blissus insularis from test plants. A. Flotation method. B. Large blower/vacuum. C. Hand-held vacuum. D. Berlese funnel. B C A D

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17 Table 2-1. Mean ( SEM) number and percentage of Blissus insularis recovered from 15.2 cm pots of ‘Palmetto’ St. Augustinegrass. 1 Sampling method Mean ( SEM) number of B. insularis recovered 2 Percentage of B. insularis recovered Flotation 22.0 1.0 c 88.0 Large blower/vacuum 15.2 2.6 bc 61.0 Hand-held vacuum 10.0 2.6 ab 40.0 Berlese funnel 3.40 1.3 a 14.0 1 25 B. insularis were placed into each pot. 2 Means SEM followed by the same letter were not significantly different ( P 0.05) by the Tukey-Kramer HSD multiple comparison test. ANOVA statistics: n = 5 reps; F = 15.16; df = 3, 19; P < 0.0001.

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18 CHAPTER 3 SOUTHERN CHINCH BUG, Blissus insularis (HETEROPTERA: BLISSIDAE), INTEGRATED PEST MANAGEMENT (IPM) Introduction St. Augustinegrass is the most widely used turfgrass in the >6.3 million lawns in Florida (U.S. Census Bureau 2000). Its primary insect pest is the southern chinch bug, Blissus insularis which sucks fluid from the crown and stems of the grass. Control of this pest has historically been achieved by use of up to six insecticide applications per year (Reinert 1978, Cherry 2001c). Organophosphates, such as chlorpyrifos and diazinon, were routinely used to control and prevent outbreaks, but are no longer available because of the Food Quality Protection Act. The primary insecticides remaining for urban turfgrass use are pyrethroids. Given the history of B. insularis resistance to organophosphates (Kerr 1958, 1961; Reinert and Niemczyk 1982; Reinert 1982; Reinert and Portier 1983), eventual resistance to pyrethroids is possible with repeated use without rotation with another pesticide class. Other problems from overuse or misuse of pesticides include drift, run-off, ground water contamination, and non-target effects. An integrated pest management program (IPM) is needed for this pest. Biological control of turfgrass pests has been underutilized in the United States, but both parasitism and predation of B. insularis have been observed (Beyers 1924, Wilson 1929, Kerr 1966). Eumicrosoma benefica are chinch bug egg parasitoids in Florida (Reinert 1972a). Reinert (1978) also observed Geocoris spp. (Heteroptera: Lygaeidae) feeding on B. insularis Geocoris spp. are generalist predators that occur in

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19 grass systems, and feed on other chinch bug species, such as the buffalograss chinch bug, Blissus occiduus (Heng-Moss et al. 1998, Carstens 2003). However, little is known about the effectiveness of these biological control agents in suppressing B. insularis populations. Host plant resistance has been an effective tool against B. insularis but new resistant cultivars are needed. Floratam, a cultivar of St. Augustinegrass, was released in 1973 by the University of Florida and Texas A&M (Busey 1979, Trenholm et al. 2000). It successfully minimized B. insularis problems for years and is still the most widely produced cultivar of St. Augustinegrass (Haydu et al. 1998). The mode of resistance was antibiosis (Reinert and Dudeck 1974). However, B. insularis can now survive and reproduce on this variety (Busey and Center 1987; Busey 1990a and b). The objective of this study was to examine several components of an IPM program including host plant resistance, predation by G. punctipes and the efficacy of several professional and non-professional (homeowner) insecticides. Materials and Methods Insect Collection and Colony Maintenance Blissus insularis used in the following experiments were collected from St. Augustinegrass lawns in Gainesville, Alachua Co., FL., using a modified Weed Eater Barracuda blower/vacuum (Electrolux Home Products, Augusta, GA) and transported in a mesh-covered bucket to the laboratory. Insects were provided fresh cuttings of Palmetto St. Augustinegrass as needed and maintained in the laboratory 7-14 d, until being aspirated into plastic vials (2.5 10.2 cm) containing a cone-shaped, moistened, 70 mm Whatman filter paper, and a foam cap.

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20 Host Plant Resistance Blissus insularis survival on an experimental variety (FHSA-115) and susceptible cultivar of St. Augustinegrass (Floratine) was tested in a laboratory assay. Terminal sprigs of FHSA-115 and Floratine St. Augustinegrass were cut from established plots at the G.C. Horn Memorial Turfgrass Field Laboratory at the University of Florida (Gainesville, FL), transported on ice to the laboratory, and refrigerated (<12 h). All sprigs were between 5.0-6.4 cm in length, with three leaflets and one node. Lab assay. The arena consisted of a cone-shaped, moistened, 70 mm Whatman filter paper at the bottom of a plastic vial (2.5 10.2 cm), one grass sprig (either FHSA115 or Floratine), and a foam cap to prevent insects from escaping. Ten B. insularis (either first-second instar, third-fourth instar, fifth instar, or adult) were transferred into each vial using a camel-hair brush. Certain instars were grouped because of the difficulty separating and distinguishing between life stages. One uninfested sprig of each variety served as a control to ensure that grasses did not die from factors other than B. insularis feeding. Sprigs were replaced every 7 d. There were 10 replicates in a complete randomized design (CRD). All vials were kept at 80 F ( 2 ) and a photoperiod of 11:13 (L:D). The number of dead B insularis were counted daily for 3 wk. An analysis of variance (ANOVA) was conducted to determine B. insularis mortality differences between grasses. Treatment means were analyzed using Tukey’s Studentized Range (HSD) test (SAS Institute Inc. 2000). Predation Assay Geocoris punctipes were acquired from a laboratory colony maintained by Entomos, LLC. (Gainesville, FL), and immediately separated to avoid cannibalism.

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21 Individuals were transferred with a camel-hair brush to petri dishes containing four Entomos food beads, a piece of moistened cotton ball, and one-fourth of an 11 cm Whatman filter paper. After 24 h of feeding on the food beads, G. punctipes were starved 24 h before bioassay, but were provided moistened cotton balls. Blissus insularis were collected as previously described. Bioassay. A cone-shaped, moistened, 70 mm Whatman filter paper was placed at the bottom of each plastic vial (2.5 10.2 cm) and one Palmetto St. Augustinegrass sprig was added to the vial. Sprigs of Palmetto St. Augustinegrass were cut from established pots maintained in the Landscape Entomology greenhouse at the University of Florida (Gainesville, FL) and refrigerated until use (<12 h). All sprigs were between 5.0-6.4 cm in length, with three leaflets and one node. Twenty B. insularis from three different age groups (either first-third instar, fourth-fifth instar, or adult) were transferred into each vial using a camel-hair brush. Certain instars were grouped because of the difficulty separating and distinguishing between life stages. Insects were allowed to acclimate for 24 h before the bioassay. Either one adult or one fifth-instar G. punctipes was transferred into each vial using a camel-hair paintbrush. Control vials containing first–third B. insularis and no G punctipes were set up to ensure B. insularis survival. Vials were kept in the laboratory at 80 F and 12:12 h (L:D). There were ten replicates in a CRD. After 24 h, G. punctipes were removed from vials. The number of B. insularis live, injured, or dead were counted, and percentage mortality were determined. Percent mortality was square root arcsine transformed. An analysis of variance (ANOVA) was conducted to determine if mean mortality of a particular age group was

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22 greater when compared to others. Treatment means were analyzed using Tukey’s Studentized Range (HSD) test (SAS Institute, Inc. 2000). Insecticidal Control Lab assay. Different concentrations of an experimental biorational product (Bioblitz, Jentree Canada, INC.) and an industry standard (diazinon, Spectracide Group, Division of United Industries Corporation) were tested in the laboratory. All treatments were applied on 27 Nov 2002 and replicated ten times in a CRD. Floratine St. Augustinegrass sprigs were cut from established plots at the G.C. Horn Memorial Turfgrass Field Laboratory in Gainesville, FL, and dipped in 1, 10, or 100 ppm Bioblitz or diazinon. Control sprigs were dipped in water. For the bioassay, a cone-shaped, moistened, 70 mm Whatman filter paper was placed at the bottom of each plastic vial (2.5 10.2 cm), and one dry, treated sprig was added to the vial. All sprigs were between 5.0-6.4 cm in length, with three leaflets and one node. Blissus insularis were collected and maintained as previously described. Ten healthy B. insularis adults were transferred into each vial using a camel-hair brush. All vials were kept at 80 F ( 2 ) and a photoperiod of 12:12 (L:D). The number of live B. insularis were counted 2 and 4 d after treatment (DAT). Percent mortality was square root arcsine transformed. An analysis of variance (ANOVA) was conducted to determine B. insularis mortality differences between treatments. Treatment means were analyzed using Tukey’s Studentized Range (HSD) test (SAS Institute, Inc. 2000). Field tests. The efficacy of several professional and non-professional (homeowner) insecticides were field-tested against B. insularis Two schedule 80 PVC rings (15 cm

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23 diameter ca. 20 cm high) were placed 10 cm into the greenest sections of established 1 m2 Floralawn St. Augustinegrass plots at the G.C. Horn Memorial Turfgrass Field Laboratory at the University of Florida, Gainesville, FL (Fig. 3-1 A). Grass was maintained at a height of 7.6 cm and vacuumed to ensure that no B. insularis were present before ring establishment. Twenty fourth-fifth instar and ten adult B. insularis were placed in the center of each ring and allowed to acclimate for 24 h. Rings were covered with chiffon mesh to prevent insect escape and predation during the experiment. Homeowner products were applied in June 2003 [n = 4, CRD] and professional products were applied in September 2003 (n = 5, CRD design). Homeowner products included bifenthrin (Scotts MaxGuard Insect Protection with Turf Builder Fertilizer 243-10, The Scotts Company, Marysville, OH), carbaryl (GardenTech Sevin Concentrate Bug Killer, GardenTech Lexington, KY), cyfluthrin (Bayer Advanced Lawn and Garden Multi-Insect Killer Ready to Use Spray, Bayer Advanced LLC, Birmingham, AL), deltamethrin (Southern Ag Mole Cricket and Chinch Bug Lawn Insect Control, Southern Agricultural Insecticides, Inc., Palmetto, FL), -cyhalothrin (Spectracide Triazicide Soil & Turf Insect Killer Granules, Realex Corporation, Spectrum Brands, St. Louis, MO), and permethrin (Real Kill Multi-Purpose Insect Killer Concentrate Realex Corporation, Spectrum Brands, St. Louis, MO). Professional products included bifenthrin (Talstar F, FMC Corp., Philadelphia, PA), carbaryl (Sevin SL, Bayer Environmental Science, Montvale, NJ), cyfluthrin (Tempo SC Ultra, Bayer Environmental Science, Montvale, NJ), cypermethrin (Demon TC, Syngenta Crop Protection, Inc., Greensboro, NC), deltamethrin (Deltagard T & O, Bayer Environmental Science, Montvale, NJ), -cyhalothrin (Scimitar CS, Syngenta Crop

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24 Protection, Inc., Greensboro, NC), and permethrin (Astro FMC Corp., Philadelphia, PA). Control plots were untreated. All treatments were applied at the label rate. An 80% active non-ionic surfactant (Amway APSA 80, Amway Phils., L.L.C.) was added to Scimitar CS and Demon TC according to label recommendation. Liquid formulations were sprayed uniformly onto the grass using a hand-held spray bottle and granular insecticides were applied by hand and watered according to label recommendations. During the course of the experiment, irrigation was applied every morning in amounts corresponding to average monthly evapotranspiration (ET) rates for Florida. June treatments received 0.42 cm and September treatments received 0.31 cm. Soil cores were removed at 1 and 4 wk post-treatment and surviving insects were removed by flotation (Fig. 3-2 B). Percent mortality was square root arcsine transformed. An analysis of variance (ANOVA) was conducted to determine B. insularis mortality differences between treatments. Treatment means were analyzed using Tukey’s Studentized Range (HSD) test (SAS Institute, Inc. 2000). Results and Discussion More than 62% of the B. insularis successfully survived on both Floratine and FHSA-115 during the 3 wk experiment with feeding, mating, and molting observed. The different life stages of B. insularis had similar mortality after feeding on either variety (Table 3-1), suggesting that neither variety was resistant to B. insularis Fifth instar and adult G. punctipes consumed a similar number of B. insularis ( F = 0.15; df = 1, 78; P = 0.7). When pooled together, fifth instar and adult G. punctipes consumed significantly more first-third instar B. insularis compared with the other chinch

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25 bug life stages. These results are similar to reports of G. uliginosus feeding on B. occiduus where feeding was highest in first–fourth instars (Carstens 2003). After 24 h, the mean percentage ( SEM) of first–third instar, fourth–fifth instar, or adult B. insularis killed by G. punctipes were 57.0 0.4, 17.2 0.3, and 16.0 0.4, respectively (n = 20; F = 37.89; df = 2, 57; P < 0.0001). A small percentage (5.3%) of B. insularis died in the controls, possibly from handling. In the laboratory insecticide assay, diazinon (100 ppm) killed significantly more B. insularis adults on sprigs than any other diazinon or Bioblitz treatment 2 and 4 DAT (Table 3-2). Diazinon at the 1 and 10 ppm rates and Bioblitz at the 100 ppm rate were also statistically different from the untreated control at 4 DAT (Table 3-2). However, B. insularis mortality was likely too low for industry standards. Some mortality occurred in the untreated control, possibly from handling. No phytotoxicity was observed. Homeowner products containing bifenthrin, carbaryl, deltamethrin, and cyhalothrin achieved over 80% control 1 wk post-treatment in the field test (Table 3-3). Carbaryl killed significantly more B. insularis than any other treatment 4 wk after treatment. Although plots treated with bifenthrin, deltamethrin, and permethrin had less than 10% B. insularis survival, these treatments did not differ from the control (Table 32). All of the professional products were statistically different from the control 1 wk after treatment, although carbaryl was the only treatment to kill all B. insularis at 1 wk (Table 3-4). Though most product labels recommended watering in after application, the label for carbaryl did not. The effectiveness of carbaryl may be reduced by irrigation and/or rainfall within 24 h of application. Plots treated with bifenthrin had few live B. insularis at 1 wk and complete mortality at 4 wk post-treatment.

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26 The results from this research provide groundwork for the development of an IPM program for B. insularis An IPM program utilizing biological, host plant resistance, and cultural control could help reduce the amount of insecticides used to control B. insularis

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27 Table 3-1. Mean percent mortality of Blissus insularis ( SEM) after feeding on Floratine or FHSA 115 St. Augustinegrass. % Mortality SEM 1 Life Stage Floratine FHSA 115 1st-2nd instar 15.4 7.3* 23.4 4.7* 3rd-4th instar 23.0 4.7* 37.7 8.1* 5th instar 21.0 4.1* 32.0 4.2* Adult 23.0 5.8* 32.3 5.1* 1 Percent SEM followed by are not significantly different ( P 0.05) by ANOVA (n = 20 B. insularis of each respective age per vial; F = 2.02; df = 7, 72; P = 0.06).

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28 Table 3-2. Mean percent mortality of Blissus insularis ( SEM) killed by different rates of Bioblitz and diazinon at 2 and 4 days after treatment. % Mortality 1 Treatment Rate 2 DAT 4 DAT Untreated control ---12.0 0.7 23.0 0.6 Bioblitz 1 ppm 23.0 0.7 33.0 0.3 10 ppm 19.0 0.9 34.0 0.4 100 ppm 18.0 0.7 38.0 0.2 Diazinon 1 ppm 14.0 0.8 40.0 0.3 10 ppm 16.0 0.8 54.0 0.3 100 ppm 82.0 0.5 97.0 0.08 1 Percent SEM followed by are significantly different ( P 0.05) using Tukey’s Studentized Range (HSD) test (n = 10 B. insularis of each respective age per vial; 2 DAT: F = 8.28; df = 6, 63; P < 0.0001; 4DAT: F = 23.0; df = 6, 63; P < 0.0001).

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29 Table 3-3. Mean percent survival of Blissus insularis ( SEM) at 1 and 4 wk post treatment. Treatment Rate 1 wk 4 wk Control ---76.7 3.6 18.0 8.3 Bifenthrin 152.5 kg/ha 18.0 8.3* 1 6.7 4.1 Carbaryl 38.2 L/ha 3.3 1.4* 0* Cyfluthrin 19.1 L/ha 55.9 12.6 20.8 5.7 Deltamethrin 146.4 kg/ha 2.5 1.6* 5.0 1.7 -cyhalothrin 146.4 kg/ha 9.0 2.1* 21.6 4.4 Permethrin 19.1 L/ha 39.0 10.4 8.3 5.0 1 Percent SEM followed by are significantly different ( P 0.05) using Tukey’s Studentized Range (HSD) test (n = 4 reps. 1 wk: F = 13.23; df = 6, 21; P < 0.0001; 4 wk: F = 4.30; df = 6, 21; P = 0.006).

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30 Table 3-4. Mean percent survival of Blissus insularis ( SEM) at 1 and 4 wk post treatment. Treatment Rate 1 wk 4 wk Control --76.0 5.5 26.6 11.4 Bifenthrin 1.6 L/ha 1.0 0.7 1 0 Carbaryl 18.78 L/ha 0 0 Cypermethrin 2.1 L/ha 1.0 0.7 0.7 1.0 Cyfluthrin 0.86 L/ha 34.0 11.2 9.3 4.0 Deltamethrin 146 kg/ha 3.0 2.0 1.3 1.0 -cyhalothrin 1.5 L/ha 14.0 3.3 4.5 3.2 Permethrin 2.5 L/ha 7.3 4.4 2.7 2.7 1 Percent SEM followed by are significantly different ( P 0.05) using Tukey’s Studentized Range (HSD) test (n = 5 reps. 1 wk: F = 26.28; df = 7, 32; P < 0.0001; 4 wk: F = 6.37; df = 7, 32; P < 0.001).

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31 Figure 3-1. Methods used in pesticide field trials. A. PVC rings used in pesticide field trials. B. Flotation of St. Augustinegrass post-treatment. A B

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32 CHAPTER 4 ST. AUGUSTINEGRASS GROWTH RESPONSE TO THREE LEVELS OF IRRIGATION AND Blissus insularis DENSITY Introduction Blissus insularis an important pest of St. Augustinegrass, has been reported to cause more damage and be more abundant in sunny, open, drought-stressed areas of lawns (Kuitert and Nutter 1952, Reinert and Kerr 1973). Particularly susceptible areas include turf near sidewalks, pavements, or in poorly irrigated areas. The nymphs and adults live in the thatch and suck fluids from the crown and stem of grasses. This feeding results in brown, dead patches of turf that are aesthetically displeasing and allow weed encroachment. Because B. insularis populations are aggregated with up to 2,000/0.1 m2 (Reinert and Kerr 1973), turfgrass managers attempt to prevent outbreaks with frequent insecticide applications. Currently, 20-25 B. insularis /0.1 m2 is considered enough to warrant a treatment (Short et al. 1982). Moisture has been reported to have a “marked but paradoxical” effect on B. insularis (Kerr 1966). Adequately irrigated turf may be attractive to B. insularis or easier to feed on. But rapidly growing grass may withstand the effects of feeding, and excess moisture may actually suppress populations (Braman 1995). However, little is known about the response of St. Augustinegrass to B. insularis feeding and the interaction with irrigation. The objective of this research was to quantify St. Augustinegrass growth response to three levels each of irrigation and B. insularis densities.

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33 Materials and Methods Insect Collection and Maintenance Blissus insularis were collected from St. Augustinegrass lawns in Ocala, FL (Marion Co.), using a modified Weed Eater Barracuda blower/vacuum (Electrolux Home Products, Augusta, GA) and transported in mesh-covered buckets containing fresh cuttings of Palmetto St. Augustinegrass to the laboratory. Grass was replaced as needed and buckets were maintained at 80 F and 13:11 (L:D) for <2 wk. Just before bioassay, B. insularis were aspirated into plastic vials (2.5 10.2 cm) containing a cone-shaped, moistened, 70 mm Whatman filter paper and a foam cap. Bioassay. Forty-five sewer polyvinyl chloride (PVC) pipes (15.2 cm diameter 43.2 cm long) fitted with sewer caps were filled with 7.6 cm of river rock, a piece of #4 Whatman filter paper, and Arrendondo fine sand up to 1.3 cm from the top (Fig. 4-1). Holes were drilled into the sewer caps to allow drainage and the filter paper layer prevented root and soil migration into the rock layer. Soil was allowed to settle for 24 h. Forty-five 15.2 cm diameter plugs of Palmetto St. Augustinegrass were transplanted from pots onto the top of the lysimeters on 14 Aug 2003 and lysimeters were placed on reinforced metal platforms in a climate-controlled greenhouse at the G. C. Horn Memorial Turfgrass Field Laboratory at the University of Florida in Gainesville, FL. Daytime and nighttime temperatures were 27 C and 24 C, respectively. Plants were fertilized with 16-4-8 water soluble complete N source NH4NO3 at 0.5 lb N/1000 ft2 and allowed to establish for 1 mo before bioassay. Treatments included irrigation [low (30%), medium (60%), or high (100%) saturation] and either 0, 30, or 200 fourth-fifth instar B. insularis There were five

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34 replicates in a CRD. A mesh cage was placed 5.0 cm into the soil and around the plants, such that the grass blades could be maintained at 7.6 cm height. Lysimeters were completely saturated with 1,500 ml of water and allowed to drain for 24 h. After 24 h, insects were transferred from vials using a camel-hair brush and cages were closed with nylon. Each lysimeter was weighed to determine its initial saturation. Amounts of water to apply at respective irrigation levels were determined by replacing some fraction of the water used in evapotranspiration (ET). This was determined gravimetrically by the following: ET = Wmax – Wmin Wneeded = deficit irrigation level ETcontrol Wmax = Wmin + Wneeded Wmin and Wmax were the lysimeter weights before and after water was applied. Wneeded represented the water amount applied to lysimeters. ETcontrol represented the mean of 100% irrigation levels of ET. ET rates were measured and plants were irrigated weekly. After 2 mo, B. insularis were removed from lysimeters using a small, hand-held Black & Decker Dust-Buster vacuum (Bioquip Products, Rancho Dominguez, CA) equipped with a 12.7 cm long 3.8 cm diameter hose attached to a removable collecting chamber (12.7 cm long 5.1 cm diameter). The vacuum was powered by a portable 12volt DC battery pack (Bioquip Products, Rancho Dominguez, CA). Each lysimeter was vacuumed for 2 min, samples were placed into plastic bags, and the number of B. insularis collected were counted in the laboratory. To collect root data, lysimeters were emptied onto a screen table and the roots were removed just below the crown. Crown contents were placed into buckets, covered

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35 with mesh, and brought to the laboratory. Remaining B. insularis in the crown material were removed by flotation and counted. Soil was washed off roots, and roots were transported to the laboratory in paper bags. Roots were rinsed again in the laboratory using a #20 standard testing sieve (Fisher Scientific Co.) and non-root debris were removed. Wet and dry weights of roots were recorded. For initial dry weights, roots were dried for 48 h in a Blue M Stabil-Therm mechanical convection oven with Pro-Set II control at 55 C. The dry ash procedure was done to obtain the percent organic matter of the roots. Dry roots were placed in 150 ml Erlenmeyer beakers and baked at 450 C in a Fisher Scientific Isotemp Muffle furnace (model #550-58). Dry ash weights were subtracted from initial dry weights to obtain the percent organic matter of roots. Grass blades were cut weekly to a 7.6 cm height, and clippings were collected and transported to the laboratory in paper bags in a cooler to obtain fresh and dry weights. For dry weights, grass blades were dried for 24 h in a Blue M Stabil-Therm mechanical convection oven with Pro-Set II control at 55 C. An analysis of variance (ANOVA) was conducted to evaluate differences between treatment means in root weights and weekly grass clippings. Data were analyzed as a two-way factorial complete randomized design with irrigation and B. insularis as main factors using the Student-Newman-Keuls test (SAS Institute, Inc. 2000). Results and Discussion Stunted growth from B insularis feeding was observed from grass clippings collected after 1 wk (Table 4-1). Dry weights of grass clippings collected from the different B. insularis densities were different from one another except during weeks two, six, and seven (Table 4-1). During weeks two, six, and seven, the dry weight of grass

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36 clippings recovered from treatments containing 200 B. insularis were lower than dry weights of grass clippings collected from treatments containing 0 or 30 B. insularis Blissus insularis feeding from densities of 30 and 200 lowered root weight as shown in Table 4-2. Research on other Blissus spp. reports their feeding can cause wilting, chlorosis, stunting, and death through clogging of vascular transport systems (Painter 1928, Negron and Riley 1990, Spike et al. 1991). Beyer (1924) observed that large numbers of B. insularis created a ‘dwarfed condition’ of St. Augustinegrass, eventually leading to plant death. The dry weights of grass clippings collected from treatments with different irrigation levels were different during weeks five and seven. At week five, the dry weight of grass clippings collected from treatments with low irrigation were lower than the dry weights of grass clippings collected from treatments containing medium or high irrigation (Table 4-1). At week seven, dry weights of grass clippings collected from treatments with low irrigation were lower than clippings from treatments with high irrigation levels. Irrigation did not affect root weight (Table 4-2). The interaction between irrigation and B. insularis was not significant for grass clipping ( F = 0.75; df = 4, 39; P = 0.57) or root weight data ( F = 0.54; df = 4, 39; P = 0.71). It is possible that B. insularis populations are found first in drought-stressed areas in the lawn because of an increased chance of survival. Heavy rainfall and irrigation have been observed to drown early instar Blissus spp. (Luginbill 1922, Beyer 1924, Wilson 1929, Kuitert and Nutter 1952). Also, environmental factors such as temperature and light may play a role in where B. insularis outbreaks occur first in the lawn.

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37 Figure 4-1. Lysimeter used in experiment. River rock filled up to sewer cap and filter paper placed on top of rock Arrendondo soil St. Augustinegrass enclosed in mesh cage

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38Table 4-1. Mean dry weight (mg) ( SEM) of grass clippings by week. 1 Main Effect Level 1 wk 2 wk 3 wk 4 wk 5 wk 6 wk 7 wk 8 wk B. insularis 0 400.0 20.0a 320.0 20.0a 270.0 60.0a 180.0 30.0a 170.0 30.0a 130.0 20.0a 110.0 20.0a 110.0 20.0a 30 330.0 20.0b 280.0 20.0a 190.0 60.0b 140.0 40.0b 140.0 50.0b 110.0 30.0a 100.0 20.0a 80.0 20.0b 200 230.0 20.0c 140.0 10.0b 60.0 10.0c 40.0 3.00c 20.0 4.00c 30.0 6.00b 20.0 3.00b 20.0 4.00c F -value 17.98 34.78 59.14 37.89 72.86 38.25 30.61 26.03 P -value < 0.0001 < 0.0001 < 0.0001 < 0.0001 <0.0001 <0.0001 <0.0001 <0.0001 Irrigation low 330.0 30.0a 240.0 20.0a 160.0 10.0a 110.0 0a 90.0 1.00b 70.0 0a 60.0 0.30b 60.0 2.00a med 310.0 20.0a 250.0 30.0a 180.0 10.0a 120.0 5.00a 120.0 10.0a 100.0 3.00a 80.0 3.00ba 70.0 10.0a high 320.0 20.0a 250.0 20.0a 190.0 10.0a 130.0 4.00a 130.0 4.00a 100.0 6.00a 90.0 3.00a 70.0 4.00a F -value 0.39 0.12 2.12 1.55 6.88 2.90 3.18 26.0 P -value 0.68 0.88 0.13 0.23 0.003 < 0.07 < 0.05 0.10 1 Means SEM within a column followed by the same letter are not significantly different (P 0.05) by the Student-Newman-Kuels test.

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39 Table 4-2. Mean root weight ( SEM) of St. Augustinegrass after 8 wk. 1 Mean root weight SEM followed by are significantly different (P 0.05) using Student-Newman-Kuels test. ( B. insularis : F = 7.33; df = 2, 42; P = 0.002. Irrigation: F = 1.81; df = 2, 42; P = 0.18). Treatment Level Mean root wt (mg) B. insularis 0 2100.0 140.0 30 1600.0 170.0 1 200 1300.0 130.0 Irrigation low 1700.0 200.0 med 1400.0 130.0 high 1800.0 150.0

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40 LIST OF REFERENCES Backus, E. A., W. B. Hunter, and C. N. Arne. 1988. Technique for staining leafhopper salivary sheaths and eggs within unsectioned plant tissue. J. Econ. Entomol. 81: 1819-1823. Banks, W. A., C. S. Lofgren, D. P. Jouvenaz, C. E. Stringer, P. M. Bishop, D. F. Williams, D. P. Wojcik, and B. M. Glancey. 1981. Techniques for collecting, rearing, and handling imported fire ants. U.S. Dept. Agric. Tech. AAT-S-21. Baxendale, F. P., T. M. Heng-Moss, and T. P. Riordan. 1999. Blissus occiduus (Hemiptera: Lygaeidae): A chinch bug pest new to buffalograss turf. J. Econ. Entomol. 92: 1172-1176. Bell, K. O., and W. H. Whitcomb. 1964. Field studies on egg predators of the bollworm, Heliothis zea (Boddie). Fla. Entomol. 47: 171-180. Beyers, A. H. 1924. Chinch bug control on St. Augustinegrass. Proc. Fla. State Hort. Soc. 37: 216-219. Braman, S. K. 1995. Insect/plant stress interactions, pp. 112-114. In R. L. Brandenburg and M. G. Villani (eds.), Handbook of turfgrass insect pests. Entomological Society of America, Lanham, MD. Brandenburg, R. L. and M. G. Villani. 1995. Handbook of turfgrass insect pests. Entomological Society of America, Lanham, MD. Britcher, W. H. 1903. The chinch bug in Maine. Maine Agr. Exp. Sta. Bull. No. 91. Brogdon, J. E. and S. H. Kerr. 1961. Home gardner’s lawn insect control guide. Fla. Univ. Agric. Ext. Serv. Cir. No. 213. Bruton, B. D., R. W. Toler & J. A. Reinert. 1983. Combined resistance in St. Augustinegrass to the southern chinch bug and the St. Augustinegrass decline strain of panicum mosaic virus. Plant Disease. 67: 171-172. Busey, P. 1979. What is Floratam? Proc. Fla. State Hort. Soc. 92: 228-232. Busey, P. 1990a. Inheritance of host adaptation in the southern chinch bug (Hemiptera: Lygaeidae). Ann. Entomol. Soc. Am. 83: 563-567.

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41 Busey, P. 1990b. Polyploid Stenotaphrum D. L. germplasm resistance to the polyploid damaging population southern chinch bug (Hemiptera: Lygaeidae). Crop Sci. 30: 588-593. Busey, P. and B. Center. 1987. Southern chinch bug (Hemiptera: Heteroptera: Lygaeidae) overcomes resistance in St. Augustinegrass. J. Econ. Entomol. 80: 608611. Busey, P. and G. H. Snyder. 1993. Population outbreak of the southern chinch bug is regulated by fertilization. Int. Turf. Soc. Res. J. 7: 353-357. Buss, E. A. 2001. Insect pest management on golf courses. Florida Cooperative Extension Service, Institute of Food and Agricultural Sciences, University of Florida, Gainesville. ENY351. Buss, E. A. and D. L. Caldwell. 2001. Biology and management of tropical sod webworms. Florida Cooperative Extension Service, Institute of Food and Agricultural Sciences, University of Florida, Gainesville. ENY318. Carstens, J. D. 2003. Influence of buffalograss management practices on the chinch bug, Blissus occiduus Barber, and its natural enemies. Master’s Thesis. University of Nebraska, Lincoln. Champlain, R. A. and L. L. Sholdt. 1967a. Life history of Geocoris punctipes (Hemiptera: Lygaeidae) in the laboratory. Ann. Entomol. Soc. Am. 60 (5): 881883. Champlain, R. A. and L. L. Sholdt. 1967b. Temperature range for development of immature stages of Geocoris punctipes (Hemiptera: Lygaeidae). Ann. Entomol. Soc. Am. 60 (60): 883-885. Cherry, R. H. 2001a. Seasonal wing polymorphism in southern chinch bugs (Hemiptera: Lygaeidae). Fla. Entomol. 84: 737-739. Cherry, R. H. 2001b. Spatial distribution of southern chinch bugs (Hemiptera: Lygaeidae) in St. Augustinegrass. Fla. Entomol. 84: 151-153. Cherry, R. H. 2001c. Interrelationship of ants (Hymenoptera: Formicidae) and southern chinch bugs (Hemiptera: Lygaeidae) in Florida lawns. J. Entomol. Sci. 36 (4): 411415. Cherry, R. H. and R. T. Nagata. 1997. Ovipositional preference and survival of southern chinch bugs ( Blissus insularis Barber) on different grasses. Int. Turf. Soc. J. 8: 981-986. Crocker, R. L. 1993. Chemical control of southern chinch bug in St. Augustinegrass. Int. Turf. Soc. Res. J. 7: 358-363.

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42 Crocker R. L. and W. H. Whitcomb. 1980. Feeding niches of the big-eyed bugs Geocoris bullatus G. punctipes and G. uliginosus (Hemiptera: Lygaeidae: Geocorinae). Entomol. Soc. Am. 9 (5): 508-513. Crocker, R. L. and C. L. Simpson. 1981. Pesticide screening test for the southern chinch bug. J. Econ. Entomol. 74: 730-731. Davis, M. G. K., and D. R. Smitley. 1990. Association of thatch with populations of hairy chinch bug (Hemiptera: Lygaeidae) in turf. J. Econ. Entomol. 83: 2370-2374. Dumas, B. A., W. P. Boyer, and W. H. Whitcomb. 1962. Effect of time of day on surveys of predaceous insects in field crops. Fla. Entomol. 45: 121-128. Dunbar, D. M. 1971. Big-eyed bugs in Connecticut lawns. Conn. Agric. Exp. Stn. Cir. No. 244. Haydu, J. J., L. N. Satterthwaite, and J. L. Cisar. 1998. An economic and agronomic profile of Florida’s sod industry in 1996. Food and Resource Economics Department, Agricultural Experiment Stations and Cooperative Extension Service, Institute of Food and Agricultural Sciences, University of Florida, Gainesville. Heng-Moss, T. M., F. P. Baxendale, and T. P. Riordan. 1998. Beneficial arthropods associated with buffalograss. J. Econ. Entomol. 91(5): 1167-1172. Heng-Moss, T. M., F. P. Baxendale, T. P. Riordan, and J. E. Foster. 2002. Evaluation of buffalograss germplasm for resistance to Blissus occiduus (Hemiptera: Lygaeidae). J. Econ. Entomol. 95: 1054-1058. Henry, T. J. and R. C. Froeschner [eds.]. 1988. Catalog of the Heteroptera or True Bugs of Canada and the continental United States. E. J. Brill, Leiden. Hodges, A. W., J. J. Haydu, P. J. van Blokland, and A. P. Bell. 1994. Contribution of the turfgrass industry to Florida’s economy, 1991-92: A value-added approach. Economics Report ER 94-1, Food and Resource Economics Department Institute of Food and Agricultural Sciences, University of Florida, Gainesville. Horn, D. J. 1988. Ecological approach to pest management. Guilford Press, New York. Hutchins, S. 1994. Techniques for sampling arthropods in integrated pest management, pp. 73-98. In Larry P. Pedigo and G. David Buntin (eds.), Handbook of sampling methods for arthropods in agriculture. CRC Press, Inc., Boca Raton, Florida. Janes, M. J., A. Hager, and G. E. Carman. 1935. Preliminary studies on starvation and drowning of the chinch bug, Blissus leucopterus (Say). J. Econ. Entomol. 28: 638–646. Kelsheimer, E. G. 1952. Insects and other pests of lawns and turf. Fla. Agric. Exp. Sta. Cir. S-42.

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43 Kerr, S. H. 1956. Chinch bug control on lawns in Florida. J. Econ. Entomol. 49: 83-85. Kerr, S. H. 1958. Tests on chinch bugs and the current status of controls. Proc. Fla. Hort. Soc. 71: 400-403. Kerr, S. H. 1961. Lawn chinch bug research. Proc. Univ. Fla. Turf. Man. Conf. 9: 211221. Kerr, S. H. 1966. Biology of the lawn chinch bug, Blissus insularis Fla. Entomol. 49(1): 9-18. Khuhro, R. D. 1994. Biological studies on the chinch bug, Blissus leucopterus leucopterus (Say) in northeast Mississippi. Ph.D. dissertation. Mississippi State University, Starkville. Knowlton, G. F. 1935. Beet leafhopper predator studies. Proc. Utah Acad. Sci. Arts. Letters 12: 255-60. Kuitert, L. C., and G. C. Nutter. 1952. Chinch bug control and subsequent renovation of St. Augustinegrass lawns. Univ. Fla. Agric. Exp. Sta. Cir. S-50. Leonard, D. E. 1966. Biosystematics of the “leucopterus complex” of the genus Blissus (Heteroptera: Lygaeidae). Conn. Agric. Exp. Stn. Bull. No. 677. Leonard, D. E. 1968. A revision of the genus Blissus (Heteroptera: Lygaeidae). Ann. Entomol. Soc. Am. 61: 239-250. Luginbill, P. 1922. Bionomics of the chinch bug. U. S. Dept. Agric. Bull. No. 1016. Majeau, G., J. Brodeur, and Y. Carrire. 2000. Sequential sampling plans for the hairy chinch bug (Hemiptera: Lygaeidae). J. Econ. Entomol. 93: 834-839. McCarty, L. B. and J. L. Cisar. 1997. St. Augustinegrass for Florida lawns, pp. 12-13. In Kathleen C. Ruppert and Robert J. Black (eds.), Florida lawn handbook, 2nd ed. Department of Environmental Horticulture, Institute of Food and Agricultural Sciences, University of Florida, Gainesville. McColloch, J. W., and N. Yuasa. 1914. A parasite of the chinch bug egg. J. Econ. Entomol. 14: 219-227. McGregor, E. A., and F. L. McDonough. 1917. The red spider on cotton. U. S. Dept. Agric. Bull. No. 416. Mead, F. W. 1972. Key to the species of big-eyed bugs, Geocoris spp., in Florida (Hemiptera: Lygaeidae). Fla. Dept. Agric. Cons. Serv., Division of Plant Industry. ENY Cir. No.121.

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44 Miller, N. C. E. 1971. The biology of the Heteroptera, 2nd ed. Classey Ltd, Hampton, UK. Nagata, R. T. and R. H. Cherry. 1999. Survival of different life stages of the southern chinch bug (Hemiptera: Lygaeidae) following insecticidal applications. J. Entomol. Sci. 34: 126–131. Negrn, J. F., and T. J. Riley. 1990. Long-term effects of chinch bug (Hemiptera: Lygaeidae) feeding on corn. J. Econ. Entomol. 83: 618-620. Niemczyk, H.D., R. A. J. Taylor, M. P. Tolley, and K. T. Power. 1992. Physiological time-driven model for predicting first generation of the hairy chinch bug (Hemiptera: Lygaeidae) on turfgrass in Ohio. J. Econ. Entomol. 85: 821-829. Painter, R. H. 1928. Notes on the injury to plant cells in chinch bug feeding. Ann. Entomol. Soc. Am. 21: 232-241. Potter, D. 1998. Destructive Turfgrass Insects: Biology, Diagnosis, and Control. Ann Arbor Press. Chelsea, MI. Readio, J. and M. H. Sweet. 1982. A review of the Geocorinae of the United States East of the 100th Meridian (Hemiptera: Lygaeidae). Misc. Publ. Entomol. Soc. Am. 12: 1-91. Reinert, J. A. 1972a. New distribution and host record for the parasitoid Eumicrosoma benefica. Fla. Entomol. 55(3): 143-144. Reinert, J. A. 1972b. Control of the southern chinch bug, Blissus insularis in south Florida. Fla. Entomol. 55: 231-235. Reinert, J. A. 1978. Natural enemy complex of the southern chinch bug in Florida. Ann. Entomol. Soc. Am. 71: 728-731. Reinert, J. A. 1982. Carbamate and synthetic pyrethroid insecticides for control of organophosphate-resistant southern chinch bugs (Heteroptera: Lygaeidae). J. Econ. Entomol. 75: 716-718. Reinert, J. A. and S. H. Kerr. 1973. Bionomics and control of lawn chinch bug. Bull. Entomol. Soc. Am. 19: 91-92. Reinert, J. A. and A. E. Dudeck. 1974. Southern chinch bug resistance in St. Augustinegrass. J. Econ. Entomol. 67: 275-277. Reinert, J. A. and H. D. Niemczyk. 1982. Insecticide resistance in epigeal insect pests of turfgrass. II. Southern chinch bug resistance to organophosphates in Florida, pp. 77-80. In H. D. Niemczyk and B. G. Joyner (eds.), Advances in turfgrass entomology. Hammer Graphics, Piqua, OH.

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45 Reinert, J. A. and K. Portier. 1983. Distribution and characterization of organophosphate-resistant southern chinch bugs (Heteroptera: Lygaeidae) in Florida. J. Econ. Entomol. 76: 1187-1190. Richmond, D. S. and D. J. Shetlar. 2000. Hairy chinch bug (Hemiptera: Lygaeidae) damage, population density, and movement in relation to the incidence of perennial ryegrass infected by Neotyphodium endophytes. J. Econ. Entomol. 93: 1167-1172. SAS Institute, Inc. 2000. SAS user’s guide statistics, version 8.2. SAS Institute Inc., Cary, NC SAS Institute, Inc. 2001. Jmp In: Statistical discovery software. SAS Institute Inc., Cary, N.C. Short, D. E. and P. G. Koehler. 1979. A sampling technique for mole crickets and other pests in turfgrass and pasture. Fla. Entomol. 62: 282-283. Short, D. E., J. A. Reinert and R. A. Atilano. 1982. Integrated pest management for urban turfgrass culture Florida. pp. 25-30. In H.D. Niemcyk and B.G. Joyner (eds.) Advances in turfgrass entomology. Hammer Graphics, Inc., OH. Slater, J. A., and R. M. Baranowski. 1990. Lygaeidae of Florida (Hemiptera: Heteroptera). Vol 14. Arthropods of Florida and neighboring land areas. Fla. Dept. Agric. Cons. Serv., Gainesville, Florida. Southwood, T. R. E. 1978. Ecological methods. Halsted, New York. Spike, B. P., R. J. Wright, S. Danielson, and D. W. Stanley-Samuelson. 1991. The fatty acid compositions of phospholipids and triacyglycerols from two chinch bug species Blissus leucopterus leucopterus and Blissus iowensis (Hemiptera: Lygaeidae) are similar in the characteristic dipteran pattern. Comp. Biochem. Physiol. 998: 799-802. Sprenkel, R. K. 2003. Insect management in pasture. Florida Cooperative Extension Service, Institute of Food and Agricultural Sciences, University of Florida. ENY402. Standen, V. 2000. The adequacy of collecting techniques for estimating species richness of grassland invertebrates. J. Appl. Ecol. 37: 884-893. Stoner, A. 1970. Plant feeding by a predaceous insect, Geocoris punctipes J. Econ. Entomol. 63: 289-298. Sweet, M. H. 2000. Seed and chinch bug (Lygaeoidea), pp. 143-264. In Carl W. Schaefer and Antonio Ricardo Paninzzi (eds.), Heteroptera of economic importance. CRC Press LLC, Boca Raton, Florida.

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46 Trmla, T. 1982. Evaluation of five methods of sampling field layer arthropods, particularly the leafhopper community in grasslands. Annales Entomologicae Fennici, 48: 1-16. Trenholm, L. E., J. L. Cisar & J. B. Unruh. 2000. St. Augustinegrass for Florida lawns. Florida Cooperative Extension Service, Institute of Food and Agricultural Sciences, University of Florida, Gainesville. ENH5. Trenholm, L. E., J. L. Cisar & J. B. Unruh. 2001. Thatch and its control in Florida lawns. Florida Cooperative Extension Service, Institute of Food and Agricultural Sciences, University of Florida, Gainesville. ENH12. Trenholm, L. E., J. L. Cisar & J. B. Unruh. 2003. Let your lawn tell you when to water. Florida Cooperative Extension Service, Institute of Food and Agricultural Sciences, University of Florida, Gainesville. ENH63. Turgeon, A. J. 1996. Turfgrass management, 4th ed. Prentice Hall, Upper Saddle River, NJ. U.S. Census Bureau. 2000. General Housing Characteristics: 2000. Found in Census 2000 Summary File 1, Matrices H3, H4, H5, H6, H7, and H16. U.S. Census Bureau. 2003. State rankings: Statistical abstract of the United States. http://www.census.gov/statab/ranks/rank01.html, March 2004. Vittum, P. J., M. G. Villani, and H. Tashiro. 1999. Turfgrass insects of the United States and Canada, 2nd ed. Comstock Publishing Assoc., Ithaca. Watson, J. R. and H. E. Bratley. 1929a. The chinch bug on St. Augustinegrass lawns. Fla. Agric. Exp. Sta. Press Bull. No. 409. Watson, J. R.,and H. E. Bratley. 1929b. The chinch bug on St. Augustinegrass. Fla. Agric. Exp. Sta. Bull. No. 209: 18-20. Webster, F. M. 1907. The chinch bug. U. S. Dept. Agric. Bull. No. 69. White, W. W. and P. Busey. 1987. History of turfgrass production in Florida. Proc. Fla. State Hort. Soc. 100: 167-174. Wilson, R. N. 1929. The chinch bug in relation to St. Augustinegrass. U. S. Dept. Agric. Cir. No. 51. Wolfenbarger, D. O. 1953. Insect and mite control problems on lawn and golf grasses. Fla. Entomol. 36: 9-12. York, G. T. 1944. Food studies of Geocoris spp., predators of the beet leafhopper. J. Econ. Entomol. 37: 25-29.

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47 BIOGRAPHICAL SKETCH Julie Cara Congdon was born on October 25, 1970, in St. Petersburg, Florida. 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 College and after taking several honors courses developed an interest in entomology. In 1998, Cara entered the University of Florida as an undergraduate entomology major. While at the University 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 Florida’s Entomology and Nematology Department (urban entomology laboratory), United States Department of Agriculture (USDA), and FMC Corporation. She is a research associate for the Division of Plant Industry and is a member of the Entomological Society of America, Florida Entomological Society, Florida Turfgrass Association, Certified Pest Control Operators of Florida, Entomology and Nematology Student Organization (ENSO) and the Urban Entomological Society (UES). Cara plans to pursue a Ph.D. in entomology at the University of Florida.


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Title: Southern Chinch Bug, Blissus insularis Barber (Heteroptera: Blissidae), Management in St. Augustinegrass
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Copyright Date: 2008

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Permanent Link: http://ufdc.ufl.edu/UFE0004889/00001

Material Information

Title: Southern Chinch Bug, Blissus insularis Barber (Heteroptera: Blissidae), Management in St. Augustinegrass
Physical Description: Mixed Material
Copyright Date: 2008

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Source Institution: University of Florida
Holding Location: University of Florida
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SOUTHERN CHINCH BUG, Blissus insularis BARBER
(HETEROPTERA: BLISSIDAE), MANAGEMENT IN ST. AUGUSTINEGRASS


















By

JULIE CARA CONGDON


A THESIS PRESENTED TO THE GRADUATE SCHOOL
OF THE UNIVERSITY OF FLORIDA IN PARTIAL FULFILLMENT
OF THE REQUIREMENTS FOR THE DEGREE OF
MASTER OF SCIENCE

UNIVERSITY OF FLORIDA


2004

































Copyright 2004

by

Julie Cara Congdon

































This thesis is dedicated to an extraordinary woman, my grandmother, Margaret Congdon.















ACKNOWLEDGMENTS

I am deeply grateful to my major advisor, Dr. Eileen Buss, for her guidance,

support, and encouragement while obtaining this degree. I also thank the members of my

committee, Dr. Laurie Trenholm and Dr. Philip Koehler, for their valuable advice,

expertise, and review of this thesis.

I thank Dr. Ronald Cherry for his technical advice and the use of the

blower/vacuum. I also thank Drs. Kris Braman, Frederick Baxendale, and Grady Miller,

for their advice on my research. Special thanks go to Dr. William Crow and his wife

Laurel for "donating" their lawn to science. I thank Dr. Jerry Butler and Dr. Oscar

Liburd for the use of laboratory supplies. I acknowledge and thank Dr. Susan Halbert

and Julieta Brambila from the Division of Plant Industry for their assistance in obtaining

literature and specimen identifications.

This research could not have been completed without the assistance of several

individuals. I am grateful to Paul Ruppert, Lois Wood, Mike Wang, Kathryn Barbara,

and Jay Cee Turner for their help with data collection. Special thanks go to Brian Owens

and the employees at the G.C. Horn Memorial Turfgrass Field Laboratory for their

advice, use of supplies, and assistance with field projects. I thank Gil Marshall for his

patience and assistance with formula calculations. I also thank Jan Weinbrecht and Jason

Haugh for their advice, use of equipment, and assistance during the irrigation experiment.

Many thanks go to Becky Griffin, the Homeowners of Hickory Forest, and Colonnades

On Top of the World, for helping me obtain B. insularis for this research. I acknowledge









and thank the Bayer Corporation, Entomos, FMC Corporation, Syngenta Crop Protection,

Inc., and Southern Agricultural Insecticides, Inc., for donating products for this research.

I acknowledge and thank Lyle Buss for his assistance with photography. I thank

Matthew Aubuchon for assistance with SAS programming. Many thanks go to the

faculty, students, and staff of the Entomology and Nematology Department for their

advice, guidance, and friendship. Special thanks are given to my family and friends for

all of their love and support. I am especially thankful to my fiance, Ricky Vazquez, for

his assistance in data collection, statistical analysis, editing of this thesis, and in his

confidence that I would succeed in obtaining this degree.
















TABLE OF CONTENTS

page

ACKNOW LEDGM ENTS ........................................ iv

LIST OF TA BLES ................... ................................................... ............ viii

LIST OF FIGURES .. .............. ............................. ix

ABSTRACT ............... ................................................ ........ x

CHAPTER

1 L ITE R A TU R E R E V IE W ................................................................. ............... 1

Chinch Bugs..................... ................... ........ .2
M anagem ent Practices ............................................................................5
Chemical Control....................................... ........5
Biological Control .................................................... ........5
Big-eyed bugs....................6................................................
G eocoris punctipes ............... .................. ...... ..... ....... .. .. .......... .7
H ost Plant R resistance .............................. ..... ......... ............ ..8
Cultural Control ............................................. ............. ........ 8
Objectives ......................................................... ..................9

2 SAMPLING METHODS FOR THE SOUTHERN CHINCH BUG, Blissus insularis
BARBER (HETEROPTERA: BLISSIDAE)............................................................ 10

Introduction............ .................... ........ .....................10
M materials and M methods ................................................................ ............... 11.....
Results and Discussion ............. .. ... ........ .. ...................13

3 SOUTHERN CHINCH BUG, Blissus insularis (HETEROPTERA: BLISSIDAE),
INTEGRATED PEST MANAGEMENT (IPM)....................................................18

Introduction ...................................... .......... ........... .18
M materials and M methods ........................................19
Insect Collection and Colony Maintenance................................................... 19
H ost Plant R resistance .............................. .............................20
Predation Assay ...... ......... ........ ............. ..................20
Insecticidal Control .............................. ...... ...... .............. 22









R results and D discussion ....................................................... 24

4 ST. AUGUSTINEGRASS GROWTH RESPONSE TO THREE LEVELS OF
IRRIGATION AND Blissus insularis DENSITY ........................32

Introduction............... .................. .......... .... ...................... 32
M materials and M methods ............................................................33
Insect Collection and M maintenance .......................................33
Results and Discussion ........................ ................. ..... .. .35

LIST O F REFEREN CE S ........................................................................................ ........40

BIOGRAPHICAL SKETCH .............. .. ...... ......... .......... 47
















LIST OF TABLES


Table page

2-1. Mean ( SEM) number and percentage of Blissus insularis recovered from 15.2 cm
pots of 'Palmetto' St. Augustinegrass ............................................17

3-1. Mean percent mortality of Blissus insularis ( SEM) after feeding on Floratine or
FH SA 115 St. Augustinegrass. ................................ ............... 27

3-2. Mean percent mortality of Blissus insularis ( SEM) killed by different rates of
Bioblitz and diazinon at 2 and 4 days after treatment............... ....... .........28

3-3. Mean percent survival of Blissus insularis (I SEM) at 1 and 4 wk post treatment ..29

3-4. Mean percent survival of Blissus insularis (I SEM) at 1 and 4 wk post treatment ..30

4-1. Mean dry weight (mg) ( SEM) of grass clippings by week .................................38

4-2. Mean root weight ( SEM) of St. Augustinegrass after 8 wk ............... ...............39
















LIST OF FIGURES

Figure page

2-1. St. Augustinegrass grown in 15.2 cm diameter clay pot enclosed in a chiffon mesh
cage.......................................................... ........ ....... ......... 16

2-2. Equipment used to recover Blissus insularis from test plants................................. 16

3-1. Methods used in pesticide field trials .......... ..........................31

4-1. Lysimeter used in experiment..............................................................................37
















Abstract of Thesis Presented to the Graduate School
of the University of Florida in Partial Fulfillment of the
Requirements for the Degree of Master of Science

SOUTHERN CHINCH BUG, Blissus insularis BARBER,
(HETEROPTERA: BLISSIDAE), MANAGEMENT IN ST. AUGUSTINEGRASS

By

Julie Cara Congdon

May 2004

Chair: Eileen A. Buss
Major Department: Entomology and Nematology

The southern chinch bug, Blissus insularis Barber, is the most destructive insect

pest of St. Augustinegrass (Stenotaphrum secundatum [Walt.] Kuntze). The purpose of

this research was to examine the effectiveness of different components of an IPM

program for B. insularis, including sampling techniques, host plant resistance, biological,

chemical, and cultural controls.

The optimal sampling method for B. insularis in a greenhouse experiment was

flotation, compared to using a hand-held vacuum or berlese funnel. The large

blower/vacuum was nearly as accurate at extracting B. insularis, so it was used to collect

field populations for use in experiments and flotation was used to recover insects from

test plants in the laboratory. In a no-choice test, more than 62% of the B. insularis

successfully survived on both 'Floratine' and 'FHSA-115'. During the 3 wk experiment

feeding, mating, and molting were observed. The different life stages of B. insularis had

similar mortality after feeding on either variety, suggesting that neither variety was









resistant to B. insularis. Fifth instar and adult Geocoris punctipes (Say) consumed a

similar number of B. insularis. When pooled together, fifth instar and adult G. punctipes

consumed significantly more first-third instar (57%) B. insularis than fourth-fifth instar

or adult stages (17.2 and 16.0% respectively).

This study evaluated the efficacy of professional and non-professional

(homeowner) products. In the laboratory insecticide assay, diazinon (100 ppm) killed

significantly more B. insularis adults on sprigs than any other diazinon or Bioblitz

treatment 2 and 4 days after treatment. Homeowner products containing bifenthrin,

carbaryl, deltamethrin, and X-cyhalothrin achieved over 80% control 1 wk post-treatment

in the field test. All of the professional products were statistically different from the

control 1 wk after treatment, although carbaryl was the only treatment to kill all B.

insularis at 1 wk.

Stunted growth from B. insularis feeding was observed from grass clippings

collected after 1 wk. Dry weights of grass clippings collected from the different B.

insularis densities were different from one another except during weeks two, six, and

seven. During this time, the dry weights of grass clippings recovered from treatments

containing 200 B. insularis were lower than dry weights of grass clippings collected from

treatments containing 0 or 30 B. insularis. Blissus insularis feeding in densities of 30 and

200 lowered root weight. At five weeks, dry weight of grass clippings was lowest in

treatments with low irrigation. At week seven, dry weights of grass clippings collected

from treatments with low irrigation were lower than clippings from treatments with high

irrigation levels. Irrigation did not impact root growth. Irrigation and B. insularis

density did not interact to affect plant growth.














CHAPTER 1
LITERATURE REVIEW

Turfgrass production and maintenance are valuable parts of the Florida economy

(Hodges et al. 1994). In 1992, residential lawns accounted for 75% of the 4.4 million

acres of turfgrass used and maintained throughout the state. Consumers spent $5 billion

on turfgrass maintenance or roughly $1,200 per acre. Sales from products and services

by turfgrass producers and commercial distributors totaled $6.5 billion. The demand for

high quality turfgrass is growing with Florida's increasing annual population of almost

2% (U.S. Census Bureau 2003). Common grasses used in Florida include bahiagrass

(Paspalum notatum Flugge), bermudagrass [Cynodon dactylon ( L.) Pers], centipedegrass

[Eremochloa ophiuroides (Munro) Hack], seashore paspalum (Paspalum vaginatum

Swartz), and zoysiagrass (zoysia spp). However, of the >6.3 million residences in Florida

(U.S. Census Bureau 2000), the most widely used grass is St. Augustinegrass

(Stenotaphrum secundatum [Walt.] Kuntze).

St. Augustinegrass is a warm-season, coarse-textured, aggressive, and stoloniferous

grass that is widely used in lawns in the coastal regions of the United States (Turgeon

1996). Most St. Augustinegrass cultivars have good salt and shade tolerance and

establish easily from sprigs or sod. Its aggressive growth habit gives it good recuperative

capability, but it is prone to thatch buildup (Potter 1998). Cultivars of St. Augustinegrass

that are commonly marketed include 'Floratam', 'Palmetto', 'Delmar', 'Floratine', and

'Seville' (White and Busey 1987, McCarty and Cisar 1997, Trenholm et al. 2000).









Several insect pests attack St. Augustinegrass in Florida including tropical sod

webworms [Herpetogramma phaeopteralis (Guenee)], armyworms (Spodoptera spp.),

grass loopers (Mocis spp.), mole crickets (Scapteriscus spp.), rhodesgrass mealybugs

[Antonina graminis (Maskell)], and southern chinch bugs (Blissus insularis Barber). Sod

webworms feed primarily at night. Newly hatched larvae skeletonize the grass blades,

while older larvae chew on grass blades near the soil surface (Buss and Caldwell 2001).

Armyworms first skeletonize the grass blades and later create bare spots. Mature larvae

feed during the day and night (Buss 2001). Grass loopers chew on the grass blades,

resulting in ragged, yellow to brown patches of grass (Sprenkel 2003). Mole crickets

damage turfgrass by their tunneling, which uproots and dries out grass plants. Feeding

from rhodesgrass mealybugs discolors grass and heavy infestations give the appearance

that excess fertilizer has 'caked' around the grass nodes (Buss 2001). The southern

chinch bug is the most destructive insect pest of St. Augustinegrass and can cause

extensive damage or kill entire lawns (Reinert and Kerr 1973, Bruton et al. 1983).

Chinch Bugs

Many Blissus spp. are important pests that attack plants in the family Graminae.

Blissus spp. feed on the phloem and xylem near the base of the plant where the grass

meristem occurs (Painter 1928). Salivary sheaths are deposited along feeding tracks

within the tissues, which is characteristic of feeding on phloem tissue bundles (Backus et

al. 1988). Their feeding can cause wilting, chlorosis, stunting, and death through

clogging of vascular transport systems (Painter 1928, Negron and Riley 1990, Spike et al.

1991). In particular, B. insularis prefer open sunny areas of St. Augustinegrass,

especially areas with abundant thatch (Reinert and Kerr 1973), where they suck fluids

from the crown and stem of grasses with their needle-like mouthparts. As the grass dies,









the insects feed at the edge of the dead spots, thus enlarging the damaged area. Blissus

insularis activity begins between March and April (Kerr 1966) and continues until

October. Damage is usually first noticed in drought-stressed areas along sidewalks,

pavements, or in poorly irrigated areas.

At least 15 Blissus species occur in the United States and Canada; however, only

four are considered damaging to turfgrass, including the buffalograss chinch bug, (B.

occiduus Barber); common chinch bug, [B. leucopterus leucopterus (Say)]; hairy chinch

bug, (B. 1. hirtus Montandon); and southern chinch bug, (B. insularis) (Brandenburg and

Villani 1995, Sweet 2000). Blissus occiduus is a serious pest of buffalograss in Arizona,

California, Colorado, Kansas, Montana, Nebraska, New Mexico, Oklahoma, and parts of

Canada (Baxendale et al. 1999, Vittum et al. 1999, Carstens 2003). Blissus leucopterus

leucopterus has caused severe damage to wheat, spelt, corn, millet, pearl millet, rye,

barley, sorghum, grain sorghum, rice, and Sudan grass (Sweet 2000). Blissus leucopterus

leucopterus can be found in Colorado, the Great Plains, New Mexico, and Nebraska

eastward to the Atlantic Ocean (Webster 1907, Leonard 1966, Khuhro 1994). Blissus

leucopterus hirtus is a major pest of cool-season grasses throughout the Northeast, parts

of the Midwest, into the mid-Atlantic states as far south as Virginia, and west to

Minnesota (Vittum et al. 1999). Blissus insularis occurs in Florida, Alabama, Georgia,

Louisiana, Mississippi, North and South Carolina, Texas, California, and Mexico (Henry

and Froeschner 1988, Sweet 2000).

Adult B. insularis are small and somewhat elongate insects measuring about 3 mm

long and 1 mm wide (Leonard 1968). Adult females are slightly larger than males. The

head is usually narrower than the posterior margin of the pronotum (Miller 1971). Wings









are white with a distinctive triangular-shaped 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 long-winged forms (macropterous), short-winged

forms brachypterouss), or both (Webster 1907). In Florida, macroptery is greatest during

the summer and fall (Cherry 2001a). Young nymphs are as small 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. Eggs are laid singly or a few at a time in sheaths, soft soil, or in

other protected areas. The eggs are white when first laid but turn bright orange just

before hatching. Development from egg to adult requires about 13 wk at 210C and 5 wk

at 280C (Potter 1998). All life stages are present year-round 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).

Blissus insularis move between lawns mainly by walking and large numbers have

been observed crawling across sidewalks and driveways bordering heavily infested lawns

(Kerr 1966). All life stages are distributed vertically through the turf thatch and into the

upper organic layer of the soil. Densities of 500-1,000 B. wi\ni.11 i% \. 1 m2 are common

and infestations of more than 2,000/0.1 m2 have been reported (Reinert and Kerr 1973).

Light to moderate infestations are extremely aggregated in small areas in the lawn rather

than evenly distributed (Cherry 2001b). Blissus insularis also attack other lawn grasses

including bahiagrass, bermudagrass, centipedegrass, and zoysiagrass, but most of the

injury to these has occurred near heavily infested St. Augustinegrass (Kerr 1966). Other

hosts include crabgrass, torpedograss, and Pangolagrass (Slater and Baranowski 1990,

Brandenburg and Villani 1995).









Management Practices

Chemical Control

Control of B. insularis has historically been achieved using insecticides such as

tobacco dust, calcium cyanide, nicotine sulfate, DDT, parathion, dieldrin, aldrin,

chlordane, chlorpyrifos, and diazinon (Watson and Bratley 1929a and b; Kelsheimer

1952, Wolfenbarger 1953, Kerr 1956, Brogdon and Kerr 1961). Up to six insecticidal

applications per year may be needed on Florida lawns to control damaging populations of

B. insularis (Reinert 1978). With such reliance on chemical use, this insect has

developed resistance to organophosphates and organochlorines (Kerr 1958, 1961; Reinert

& Niemczyk 1982; Reinert 1982; Reinert and Portier 1983). Currently, pyrethroids are

most commonly available to homeowners and turfgrass professionals for controlling B.

insularis outbreaks. A lack of rotation with other chemical formulations increases the

risk of B. insularis developing resistance to pyrethroids as well. Some alternatives to

chemicals are biological control, host-plant resistance, and cultural control.

Biological Control

Parasites, predators, and pathogens are found in association with Blissus spp. The

parasitic wasp, Eumicrosoma benefica Gahan (Hymenoptera: Scelionidae), attacks the

eggs of B. leucopterus (Say) in Kansas, reportedly reducing 'broods' of chinch bugs by as

much as 50% (McColloch and Yuasa 1914). Eumicrosoma benefica also parasitized B.

insularis eggs in Florida and field tests showed an average abundance of 34.5 wasps/0. 1

m2 in lawns containing B. insularis populations of 90.0/0.1 m2 (Reinert 1972a). Along

with E. benefica, predatory insects [e.g., Pagasapallipes Stal (Heteroptera: Nabidae),

Xylocoris vicarius (Reuter) (Heteroptera: Anthocoridae), Lasiochilus pallidulus Reuter

(Heteroptera: Anthocoridae), Sinea spp. (Heteroptera: Reduviidae), Labidura riparia









Pallas, Geocoris bullatus (Say) and Geocoris uliginosus (Say) (Heteroptera: Geocoridae)]

were observed feeding on B. insularis in Florida (Reinert 1978). The native fire ant,

Solenopsis geminata (F.), and a spider, Lycosa sp., were also observed attacking B.

insularis (Reinert 1978). However, the red imported fire ant, Solenopsis invicta Buren,

was unable to suppress B. insularis populations (Cherry 200 1c). Beauveria globulifera

(Spegazzini) Picard and Metarhizium anisopliae (Metch.) Sorokin were unsuccessful in

controlling B. insularis in field tests (Kerr 1958). Beauveria bassiana (Balsamo)

Vuillemin was pathogenic on all life stages of B. insularis; however, success appeared

limited to high humidity, B. insularis populations, and moisture levels (Reinert 1978).

Big-eyed bugs

Big-eyed bugs, Geocoris spp., are very abundant and widely distributed (Readio

and Sweet 1982). These generalist predators attack a wide range of prey species,

including various insect and mite pests of agricultural crops such as cotton, soybean,

strawberry, peanut, turfgrass, vegetable, sugarbeet, alfalfa, and tobacco (McGregor and

McDonough 1917, Knowlton 1935, York 1944, Champlain and Sholdt 1967a and b,

Mead 1972, Crocker and Whitcomb 1980). They specifically have been observed

attacking aphids, plant bugs, eggs and larvae of the bollworm [Heliothis zea (Boddie)],

flea beetles (Phyllotreta spp.), false chinch bugs [Nysius ericae (Schilling)], southern

green stink bugs [Nezara viridula (L.)], and green peach aphids [Myzuspersicae (Sulzer)]

(Bell and Whitcomb 1964, Crocker and Whitcomb 1980). Geocoris spp. will feed on

eggs, immatures, or adults of relatively small or passive prey. They attack by inserting

their proboscis into the head, thorax, or abdomen and tend to walk about with the prey

suspended on the labium (Sweet 2000).









Although Geocoris spp. are predators, they also require water from plant sources

(York 1944). In laboratory colonies, Geocoris spp. can temporarily be maintained on

plant food alone, but prey are required for proper development and fecundity (Stoner

1970). Geocoris bullatus, once thought to be a turfgrass pest in Connecticut, feeds on

lawn insects such as B. 1. hirtus (Dunbar 1971). Geocoris uliginosus had densities as

high as 16.0/0.1 m2 when high B. insularis populations were present and fed on an

average of 9.6 3.3 (mean SEM) nymphs in 5 d under laboratory conditions (Reinert

1978). The most common species in Florida are G. uliginosus, G. bullatus, and G.

punctipes (Say) (Mead 1972).

Geocoris punctipes

Geocorispunctipes are small oblong Lygaeids having the head broader than long,

large prominent eyes that curve backward and overlap the front of the pronotum, and

tylus that has a longitudinal groove (Mead 1972). They have relatively short antennae

that are slightly enlarged at the tip. Geocorispunctipes adults are about 0.5 cm long and

are silver-gray in color. Nymphs resemble adults but lack fully developed wings. Cigar-

shaped eggs are white to tan in color with a distinctive red spot and are laid singly on

leaves and stems of many crops. Eggs hatch in approximately 1 wk. Geocoris spp. have

five instars, each of which lasts from 4 to 6 d. Adults live for about 1 mo and a female

can lay up to 300 eggs during her lifetime. Several generations may occur throughout the

season. Adult and immature G. punctipes feed by sucking juices from their prey through

needle-like mouthparts. They are primarily active in the morning (Dumas et al. 1962).

These insects are often confused for Blissus spp. Geocoris spp. are commonly found

throughout the southern United States.









Host Plant Resistance

Host resistant grasses are used to suppress pest damage. There are three types of

resistance that can occur: antibiosis, antixenosis, or tolerance. Antibiosis affects the

biology of the pest insect and often results in increased mortality, or reduced oviposition

and longevity. With antixenosis, the pest shows non-preference for a resistant grass

compared to a susceptible one. When a host resistant grass shows tolerance, the grass is

able to withstand or recover from damage caused by an insect. Floratam was released as

a chinch bug resistant cultivar in 1973 by the University of Florida and Texas A&M.

Floratam is the most widely produced and used cultivar of St. Augustinegrass and

accounts for 80% of sod production in Florida (Haydu et al. 1998). It has a very coarse

leaf texture, poor cold and shade tolerance, is resistant to the St. Augustine decline virus,

and successfully minimized B. insularis problems for years (Busey 1979, Trenholm et al.

2000). The mode of resistance was antibiosis (Reinert and Dudeck 1974). However, B.

insularis can now survive and reproduce on this variety (Busey and Center 1987; Busey

1990a and b). Research is being conducted to develop new cultivars of St.

Augustinegrass that are resistant to B. insularis.

Cultural Control

Cultural practices may influence the susceptibility of St. Augustinegrass to B.

insularis. Over-fertilizing, over-watering, and improper mowing can cause lawn grasses

to develop a thick thatch layer (Trenholm et al. 2000). Thatch is a layer of accumulated

dead leaf blades, stolons, and roots between the live plant and soil. Accumulation of

thatch often makes the lawn more susceptible to pests (Trenholm et al. 2001), possibly by

protecting them from predation and environmental stress. The abundance of B. 1. hirtus

was found to be closely linked to thatch thickness in lawns (Davis and Smitley 1990).









Excessive thatch (exceeding 2.5 cm) may need to be professionally removed by vertical

mowing (Trenholm et al. 2000). Heavy fertilization may cause increased growth, thatch,

susceptibility to pests such as B. insularis, and reduced tolerance to environmental

stresses (Kerr 1966, Busey and Snyder 1993, Trenholm et al. 2001). Excess irrigation

may lead to problems such as a shallow root system, increased thatch, and increased pest

problems (Trenholm et al. 2003). Reduced fertilization and proper irrigation practices

may help to reduce B. insularis problems (Potter 1998, Sweet 2000). St. Augustinegrass

should be mowed to a height of 8 10 cm (Trenholm et al. 2000). Proper cultural

practices promote healthy grass, which may be able to better tolerate chinch bug damage.

Objectives

An integrated pest management program (IPM) is needed for the southern chinch

bug to reduce the turfgrass industry's sole reliance on pesticides. The purpose of this

study was to evaluate the effectiveness of different components of an IPM program for B.

insularis including: 1) sampling techniques, 2) the potential of 'FHSA-115' as a chinch

bug resistant variety of St. Augustinegrass, 3) B. insularis predation by G. punctipes, 4)

several homeowner, professional, and experimental insecticides labeled for chinch bug

control, and 5) St. Augustinegrass growth response to three levels of irrigation and B.

insularis density.














CHAPTER 2
SAMPLING METHODS FOR THE SOUTHERN CHINCH BUG, Blissus insularis
BARBER (HETEROPTERA: BLISSIDAE)

Introduction

A successful integrated pest management (IPM) program largely depends on the

development of an effective and efficient sampling technique (Hutchins 1994). The

proper use of such techniques provides a means by which to make informed decisions,

such as establishing more accurate economic and/or aesthetic thresholds (Southwood

1978), evaluating product trials, and designing innovative monitoring or management

strategies. However, the researcher must have an understanding of the biology of the

insect population and environmental factors (i.e. temperature, humidity, light), which

may affect activity periods of the insect and critical thresholds (Horn 1988). Effective

sampling also depends on insect distribution and behavior (Tormala 1982, Standon

2000).

Insects with clumped distributions that live in dense plant material, such as the

southern chinch bug, Blissus insularis, may be somewhat difficult to accurately sample

(Cherry 2001b). Blissus insularis is one of the most important pests of St.

Augustinegrass (Kerr 1966, Crocker 1993, Cherry and Nagata 1997). Infestations usually

occur in open, sunny, and drought-stressed areas near sidewalks and driveways. Nymphs

and adults live in the thatch and suck fluids from the grass with their needle-like

mouthparts, resulting in yellow to brown patches in the lawn. Newly hatched B. insularis

nymphs often wedge themselves in between the grass blades at the nodes to feed (Kerr









1966), making it difficult to recover them without physically pulling the blades back to

expose the insects. Techniques used to extract turfgrass insects include flotation, irritant

sampling, and vacuuming. Although some studies have examined the effectiveness of

various techniques used in sampling turfgrass pests, including chinch bugs (Short &

Koehler 1979, Majeau et al. 2000), vacuum extraction was not included. Our objective

for this study was to determine the most accurate and efficient method for specifically

extracting B. insularis from St. Augustinegrass.

Materials and Methods

Blissus insularis were mass-collected from St. Augustinegrass lawns in Gainesville,

FL (Alachua Co.), using a modified blower/vacuum (Electrolux Home Products,

Augusta, GA) and transported in a mesh-covered bucket to the laboratory. Insects were

provided fresh cuttings of Palmetto St. Augustinegrass as needed and maintained in the

laboratory <10 d until being aspirated into plastic vials (2.5 x 10.2 cm) containing a cone-

shaped, moistened, 70 mm Whatman filter paper to prevent being damaged from

aspiration. Vials were closed using a foam cap to prevent insect escape.

Twenty 15.2 cm diameter clay pots of Palmetto St. Augustinegrass were

established in Arrendondo fine sand (loamy, siliceous, hypothermic, Grossarenic

Paleudalt) and maintained under ambient conditions in a greenhouse at the University of

Florida (Gainesville, FL). Plants were fertilized every 2 wk and watered as needed. The

thatch layer was <0.6 cm. Plants were maintained at a 7.6 cm height. Twenty fourth-

fifth instars and five adult B. insularis were transferred from vials using a camel-hair

brush into the center of each pot and caged on 13 May 2003 (Fig. 2-1). Insects were

maintained in caged pots for 7 d. Plants were arranged in a complete randomized design









(CRD). After 7 d, four sampling methods (flotation; large, gas powered vacuum; small,

battery operated vacuum; or berlese funnel) were used to extract the insects (n = 5 pots

per treatment).

The first method involved floating B. insularis from the plant material. Blissus

insularis were forced from plant debris using a modified version of the flotation

technique used in extracting Solenopsis invicta colonies from soil (Banks et al. 1981).

Potted plants were placed into 19 liter buckets, water was added to each bucket with 1.3

cm diameter PVC tubing, brass connectors, and 7.6 cm long x 0.6 cm diameter rubber

tubing (Fig. 2-2 A). Warm water (~380C) slowly filled up to the edge of each pot, was

reduced to a fast drip, and was turned off after about half of the grass blades were

submerged. Insects were collected as they crawled up the grass blades, were placed into

vials of 80% EtOH, and counted. Pots were submerged for ~2 h.

The second method used was a large, gas-powered blower/vacuum (Electrolux

Home Products, Augusta, GA) equipped with a 0.8 m long hose attachment (Fig. 2-2 B).

Insects were collected into a 12.7 cm diameter knitting-ring with a chiffon mesh that

covered the vacuum attachment's intake hose. Each pot of grass was vacuumed for 1

min, samples were placed into plastic bags, and the number of chinch bugs collected were

counted in the laboratory.

A smaller, hand-held, modified Black & Decker Dust-BusterTM vacuum (Bioquip

Products, Rancho Dominguez, CA) equipped with a 12.7 cm long x 3.8 cm diameter hose

attached to a removable collecting chamber (12.7 cm long x 5.1 cm diameter) was used

as the third sampling method (Fig. 2-2 C). The vacuum was powered by a portable, 12-









volt DC battery pack (Bioquip Products, Rancho Dominguez, CA). Each pot was

vacuumed as described above.

The last method involved placing infested plants into berlese funnels using 40

wattage bulbs. Potted plants were placed sideways into the berlese funnels and were left

in the pots to minimize debris falling into the collection containers (Fig. 2-2 D). It was

assumed that any living chinch bugs would fall into the funnel and container of 80%

EtOH as the soil and plants dried within 48 h.

The total number of B. insularis collected using each method was counted. The

mean number of chinch bugs recovered was analyzed using an analysis of variance

(ANOVA, P < 0.05) and treatment means were compared using the Tukey-Kramer HSD

multiple comparison test (Jmp@, SAS Institute Inc. 2001).

Results and Discussion

Flotation was the most effective technique for extracting a known number of B.

insularis from pots of grass (Table 2-1). Different versions of this method are commonly

used in B. insularis experiments. Under field conditions, a large metal cylinder is placed

3-5 cm into the soil near damaged turfgrass, filled with water, and any chinch bugs

present float to the surface within 3-10 min (Kerr 1966; Reinert 1972b, 1982). For

laboratory or greenhouse studies, cores or pots of grass are submerged in buckets of

water, insects crawl up the grass blades, and are collected (Nagata and Cherry 1999,

Richmond and Shetlar 2000). In this experiment, B. insularis were forced to escape

slowly increasing water levels and could float on the water if they fell off the plant

material. Even if completely submerged for considerable periods (<4 h), chinch bugs

were expected to survive (Britcher 1903, Janes et al. 1935). However, this method did









not recover all of the insects. Some mortality may have occurred during the experiment

or some insects may have remained in the plant material. Because soil particles were

heavy and fell to the bottom of the bucket, very little debris interfered with observation of

B. insularis movement on the plants or water. Although flotation was somewhat labor-

intensive, it extracted the highest number of B. insularis and was the only method where

variability among treatments was low.

The number of B. insularis recovered by flotation compared to the large

blower/vacuum was not statistically significant (22.0 and 15.2 respectively, Table 2-1).

Vacuums similar to the one used in this experiment have successfully been used for

collecting chinch bugs for laboratory assays and from field insecticide plots (Crocker and

Simpson 1981, Crocker 1993, Nagata and Cherry 1999). Standen (2000) compared the

effectiveness of pitfall traps and a D-vac suction trap combined with a lightweight swish

net and reported the highest number of Hemipterans being collected in D-vac samples.

Several different vacuums (whereby insects are sucked into a layer of fabric that is

stretched across an intake hose and released into a different container) or D-vac machines

are used to collect insects (Crocker and Simpson 1981, Crocker 1993, Nagata and Cherry

1999, Cherry 200 Ib). Vacuum extraction of insects may be faster and at least as

effective as the flotation method (Crocker 1993).

The number of B. insularis recovered by the large blower/vacuum compared to the

smaller, hand-held vacuum was not significant (Table 2-1). However, the smaller hand-

held vacuum recovered significantly fewer B. insularis than the flotation method (10.0

and 3.4 respectively). Additional sampling time may be required for the small, hand-held

vacuum to be effective in recovering known numbers of B. insularis. However, for its









convenient size, the hand-held vacuum might work well for rapid detection of chinch

bugs in infested lawns. For the berlese funnel method, soil or grass samples may be

placed into funnels and as actively moving insects escape the light and heat of low-

wattage bulbs, they drop into containers of alcohol (Niemczyk et al. 1992, Heng-Moss et

al. 2002). The lowest recovery of B. insularis occurred with the berlese funnel (Table 2-

1). Additional testing and improved methods (i.e. different bulb wattage, plant

arrangement, funnel design) for the berlese funnel technique may improve its

effectiveness in recovering B. insularis.

As part of developing an IPM program, several of these techniques would be useful

under the proper circumstances. The flotation method and both vacuums tested in this

study could be used to detect the presence of B. insularis in the lawn. However, when

conducting experiments using a known number of B. insularis, the flotation method or

large blower/vacuum may be best to use since these techniques recovered the highest

number of insects from infested pots of St. Augustinegrass.



























Figure 2-1. St. Augustinegrass grown in 15.2 cm diameter clay pot enclosed in a chiffon
mesh cage.

























Figure 2-2. Equipment used to recover Blissus insularis from test plants. A. Flotation
method. B. Large blower/vacuum. C. Hand-held vacuum. D. Berlese
funnel.












Table 2-1. Mean ( SEM) number and percentage of Blissus insularis recovered from
15.2 cm pots of 'Palmetto' St. Augustinegrass. 1
Sampling method Mean ( SEM) number of B. Percentage of B. insularis
insularis recovered 2 recovered
Flotation 22.0 1.0 c 88.0

Large blower/vacuum 15.2 2.6 bc 61.0

Hand-held vacuum 10.0 2.6 ab 40.0

Berlese funnel 3.40 1.3 a 14.0

25 B. insularis were placed into each pot.
2 Means SEM followed by the same letter were not significantly different (P <: 0.05) by
the Tukey-Kramer HSD multiple comparison test. ANOVA statistics: n = 5 reps; F=
15.16; df = 3, 19; P < 0.0001.














CHAPTER 3
SOUTHERN CHINCH BUG, Blissus insularis (HETEROPTERA: BLISSIDAE),
INTEGRATED PEST MANAGEMENT (IPM)

Introduction

St. Augustinegrass is the most widely used turfgrass in the >6.3 million lawns in

Florida (U.S. Census Bureau 2000). Its primary insect pest is the southern chinch bug,

Blissus insularis, which sucks fluid from the crown and stems of the grass. Control of

this pest has historically been achieved by use of up to six insecticide applications per

year (Reinert 1978, Cherry 2001c). Organophosphates, such as chlorpyrifos and

diazinon, were routinely used to control and prevent outbreaks, but are no longer

available because of the Food Quality Protection Act. The primary insecticides

remaining for urban turfgrass use are pyrethroids. Given the history of B. insularis

resistance to organophosphates (Kerr 1958, 1961; Reinert and Niemczyk 1982; Reinert

1982; Reinert and Portier 1983), eventual resistance to pyrethroids is possible with

repeated use without rotation with another pesticide class. Other problems from overuse

or misuse of pesticides include drift, run-off, ground water contamination, and non-target

effects. An integrated pest management program (IPM) is needed for this pest.

Biological control of turfgrass pests has been underutilized in the United States,

but both parasitism and predation of B. insularis have been observed (Beyers 1924,

Wilson 1929, Kerr 1966). Eumicrosoma benefica are chinch bug egg parasitoids in

Florida (Reinert 1972a). Reinert (1978) also observed Geocoris spp. (Heteroptera:

Lygaeidae) feeding on B. insularis. Geocoris spp. are generalist predators that occur in









grass systems, and feed on other chinch bug species, such as the buffalograss chinch bug,

Blissus occiduus (Heng-Moss et al. 1998, Carstens 2003). However, little is known about

the effectiveness of these biological control agents in suppressing B. insularis

populations.

Host plant resistance has been an effective tool against B. insularis, but new

resistant cultivars are needed. Floratam, a cultivar of St. Augustinegrass, was released in

1973 by the University of Florida and Texas A&M (Busey 1979, Trenholm et al. 2000).

It successfully minimized B. insularis problems for years and is still the most widely

produced cultivar of St. Augustinegrass (Haydu et al. 1998). The mode of resistance was

antibiosis (Reinert and Dudeck 1974). However, B. insularis can now survive and

reproduce on this variety (Busey and Center 1987; Busey 1990a and b).

The objective of this study was to examine several components of an IPM

program including host plant resistance, predation by G. punctipes, and the efficacy of

several professional and non-professional (homeowner) insecticides.

Materials and Methods

Insect Collection and Colony Maintenance

Blissus insularis used in the following experiments were collected from St.

Augustinegrass lawns in Gainesville, Alachua Co., FL., using a modified Weed Eater

Barracuda blower/vacuum (Electrolux Home Products, Augusta, GA) and transported in

a mesh-covered bucket to the laboratory. Insects were provided fresh cuttings of

Palmetto St. Augustinegrass as needed and maintained in the laboratory 7-14 d, until

being aspirated into plastic vials (2.5 x 10.2 cm) containing a cone-shaped, moistened, 70

mm Whatman filter paper, and a foam cap.









Host Plant Resistance

Blissus insularis survival on an experimental variety (FHSA-115) and susceptible

cultivar of St. Augustinegrass (Floratine) was tested in a laboratory assay. Terminal

sprigs of FHSA-1 15 and Floratine St. Augustinegrass were cut from established plots at

the G.C. Horn Memorial Turfgrass Field Laboratory at the University of Florida

(Gainesville, FL), transported on ice to the laboratory, and refrigerated (<12 h). All

sprigs were between 5.0-6.4 cm in length, with three leaflets and one node.

Lab assay. The arena consisted of a cone-shaped, moistened, 70 mm Whatman

filter paper at the bottom of a plastic vial (2.5 x 10.2 cm), one grass sprig (either FHSA-

115 or Floratine), and a foam cap to prevent insects from escaping. Ten B. insularis

(either first-second instar, third-fourth instar, fifth instar, or adult) were transferred into

each vial using a camel-hair brush. Certain instars were grouped because of the difficulty

separating and distinguishing between life stages. One uninfested sprig of each variety

served as a control to ensure that grasses did not die from factors other than B. insularis

feeding. Sprigs were replaced every 7 d. There were 10 replicates in a complete

randomized design (CRD). All vials were kept at 800F ( 20) and a photoperiod of 11:13

(L:D). The number of dead B. insularis were counted daily for 3 wk.

An analysis of variance (ANOVA) was conducted to determine B. insularis

mortality differences between grasses. Treatment means were analyzed using Tukey's

Studentized Range (HSD) test (SAS Institute Inc. 2000).

Predation Assay

Geocorispunctipes were acquired from a laboratory colony maintained by

Entomos, LLC. (Gainesville, FL), and immediately separated to avoid cannibalism.









Individuals were transferred with a camel-hair brush to petri dishes containing four

Entomos food beads, a piece of moistened cotton ball, and one-fourth of an 11 cm

Whatman filter paper. After 24 h of feeding on the food beads, G. punctipes were starved

24 h before bioassay, but were provided moistened cotton balls. Blissus insularis were

collected as previously described.

Bioassay. A cone-shaped, moistened, 70 mm Whatman filter paper was placed at

the bottom of each plastic vial (2.5 x 10.2 cm) and one Palmetto St. Augustinegrass sprig

was added to the vial. Sprigs of Palmetto St. Augustinegrass were cut from established

pots maintained in the Landscape Entomology greenhouse at the University of Florida

(Gainesville, FL) and refrigerated until use (<12 h). All sprigs were between 5.0-6.4 cm

in length, with three leaflets and one node. Twenty B. insularis from three different age

groups (either first-third instar, fourth-fifth instar, or adult) were transferred into each vial

using a camel-hair brush. Certain instars were grouped because of the difficulty

separating and distinguishing between life stages. Insects were allowed to acclimate for

24 h before the bioassay.

Either one adult or one fifth-instar G. punctipes was transferred into each vial using

a camel-hair paintbrush. Control vials containing first-third B. insularis and no G.

punctipes were set up to ensure B. insularis survival. Vials were kept in the laboratory at

800F and 12:12 h (L:D). There were ten replicates in a CRD. After 24 h, G. punctipes

were removed from vials. The number of B. insularis live, injured, or dead were counted,

and percentage mortality were determined.

Percent mortality was square root arcsine transformed. An analysis of variance

(ANOVA) was conducted to determine if mean mortality of a particular age group was









greater when compared to others. Treatment means were analyzed using Tukey's

Studentized Range (HSD) test (SAS Institute, Inc. 2000).

Insecticidal Control

Lab assay. Different concentrations of an experimental biorational product

(Bioblitz, Jentree Canada, INC.) and an industry standard (diazinon, Spectracide Group,

Division of United Industries Corporation) were tested in the laboratory. All treatments

were applied on 27 Nov 2002 and replicated ten times in a CRD. Floratine St.

Augustinegrass sprigs were cut from established plots at the G.C. Horn Memorial

Turfgrass Field Laboratory in Gainesville, FL, and dipped in 1, 10, or 100 ppm Bioblitz

or diazinon. Control sprigs were dipped in water. For the bioassay, a cone-shaped,

moistened, 70 mm Whatman filter paper was placed at the bottom of each plastic vial

(2.5 x 10.2 cm), and one dry, treated sprig was added to the vial. All sprigs were

between 5.0-6.4 cm in length, with three leaflets and one node. Blissus insularis were

collected and maintained as previously described. Ten healthy B. insularis adults were

transferred into each vial using a camel-hair brush. All vials were kept at 800F ( 20) and

a photoperiod of 12:12 (L:D). The number of live B. insularis were counted 2 and 4 d

after treatment (DAT).

Percent mortality was square root arcsine transformed. An analysis of variance

(ANOVA) was conducted to determine B. insularis mortality differences between

treatments. Treatment means were analyzed using Tukey's Studentized Range (HSD)

test (SAS Institute, Inc. 2000).

Field tests. The efficacy of several professional and non-professional (homeowner)

insecticides were field-tested against B. insularis. Two schedule 80 PVC rings (15 cm









diameter x ca. 20 cm high) were placed 10 cm into the greenest sections of established 1

m2 Floralawn St. Augustinegrass plots at the G.C. Horn Memorial Turfgrass Field

Laboratory at the University of Florida, Gainesville, FL (Fig. 3-1 A). Grass was

maintained at a height of 7.6 cm and vacuumed to ensure that no B. insularis were

present before ring establishment. Twenty fourth-fifth instar and ten adult B. insularis

were placed in the center of each ring and allowed to acclimate for 24 h. Rings were

covered with chiffon mesh to prevent insect escape and predation during the experiment.

Homeowner products were applied in June 2003 [n = 4, CRD] and professional

products were applied in September 2003 (n = 5, CRD design). Homeowner products

included bifenthrin (Scotts MaxGuard Insect Protection with Turf Builder Fertilizer 24-

3-10, The Scotts Company, Marysville, OH), carbaryl (GardenTech Sevin Concentrate

Bug Killer, GardenTechTM, Lexington, KY), cyfluthrin (Bayer Advanced Lawn and

Garden Multi-Insect Killer Ready to Use Spray, Bayer Advanced LLC, Birmingham,

AL), deltamethrin (Southern Ag Mole Cricket and Chinch Bug Lawn Insect Control,

Southern Agricultural Insecticides, Inc., Palmetto, FL), X-cyhalothrin (Spectracide

TriazicideTM Soil & Turf Insect Killer Granules, Realex Corporation, Spectrum Brands,

St. Louis, MO), and permethrin (Real Kill Multi-Purpose Insect Killer Concentrate,

Realex Corporation, Spectrum Brands, St. Louis, MO). Professional products included

bifenthrin (Talstar F, FMC Corp., Philadelphia, PA), carbaryl (Sevin SL, Bayer

Environmental Science, Montvale, NJ), cyfluthrin (Tempo SC Ultra, Bayer

Environmental Science, Montvale, NJ), cypermethrin (Demon TC, Syngenta Crop

Protection, Inc., Greensboro, NC), deltamethrin (Deltagard T & 0, Bayer

Environmental Science, Montvale, NJ), X-cyhalothrin (Scimitar CS, Syngenta Crop









Protection, Inc., Greensboro, NC), and permethrin (Astro, FMC Corp., Philadelphia,

PA). Control plots were untreated. All treatments were applied at the label rate. An

80% active non-ionic surfactant (Amway APSA 80, Amway Phils., L.L.C.) was added

to Scimitar CS and Demon TC according to label recommendation. Liquid

formulations were sprayed uniformly onto the grass using a hand-held spray bottle and

granular insecticides were applied by hand and watered according to label

recommendations. During the course of the experiment, irrigation was applied every

morning in amounts corresponding to average monthly evapotranspiration (ET) rates for

Florida. June treatments received 0.42 cm and September treatments received 0.31 cm.

Soil cores were removed at 1 and 4 wk post-treatment and surviving insects were

removed by flotation (Fig. 3-2 B).

Percent mortality was square root arcsine transformed. An analysis of variance

(ANOVA) was conducted to determine B. insularis mortality differences between

treatments. Treatment means were analyzed using Tukey's Studentized Range (HSD)

test (SAS Institute, Inc. 2000).

Results and Discussion

More than 62% of the B. insularis successfully survived on both Floratine and

FHSA-115 during the 3 wk experiment with feeding, mating, and molting observed. The

different life stages of B. insularis had similar mortality after feeding on either variety

(Table 3-1), suggesting that neither variety was resistant to B. insularis.

Fifth instar and adult G. punctipes consumed a similar number of B. insularis (F=

0.15; df = 1, 78; P = 0.7). When pooled together, fifth instar and adult G. punctipes

consumed significantly more first-third instar B. insularis compared with the other chinch









bug life stages. These results are similar to reports of G. uliginosus feeding on B.

occiduus, where feeding was highest in first-fourth instars (Carstens 2003). After 24 h,

the mean percentage ( SEM) of first-third instar, fourth-fifth instar, or adult B. insularis

killed by G. punctipes were 57.0 0.4, 17.2 0.3, and 16.0 0.4, respectively (n = 20; F

= 37.89; df = 2, 57; P < 0.0001). A small percentage (5.3%) of B. insularis died in the

controls, possibly from handling.

In the laboratory insecticide assay, diazinon (100 ppm) killed significantly more B.

insularis adults on sprigs than any other diazinon or Bioblitz treatment 2 and 4 DAT

(Table 3-2). Diazinon at the 1 and 10 ppm rates and Bioblitz at the 100 ppm rate were

also statistically different from the untreated control at 4 DAT (Table 3-2). However, B.

insularis mortality was likely too low for industry standards. Some mortality occurred in

the untreated control, possibly from handling. No phytotoxicity was observed.

Homeowner products containing bifenthrin, carbaryl, deltamethrin, and X-

cyhalothrin achieved over 80% control 1 wk post-treatment in the field test (Table 3-3).

Carbaryl killed significantly more B. insularis than any other treatment 4 wk after

treatment. Although plots treated with bifenthrin, deltamethrin, and permethrin had less

than 10% B. insularis survival, these treatments did not differ from the control (Table 3-

2). All of the professional products were statistically different from the control 1 wk after

treatment, although carbaryl was the only treatment to kill all B. insularis at 1 wk (Table

3-4). Though most product labels recommended watering in after application, the label

for carbaryl did not. The effectiveness of carbaryl may be reduced by irrigation and/or

rainfall within 24 h of application. Plots treated with bifenthrin had few live B. insularis

at 1 wk and complete mortality at 4 wk post-treatment.






26


The results from this research provide groundwork for the development of an IPM

program for B. insularis. An IPM program utilizing biological, host plant resistance, and

cultural control could help reduce the amount of insecticides used to control B. insularis.






27


Table 3-1. Mean percent mortality of Blissus insularis ( SEM) after feeding on
Floratine or FHSA 115 St. Augustinegrass.
% Mortality SEM

Life Stage Floratine FHSA 115

1st-2nd instar 15.4 7.3* 23.4 4.7*

3rd-4th instar 23.0 4.7* 37.7 8.1*

5th instar 21.0 4.1* 32.0 4.2*

Adult 23.0 5.8* 32.3 5.1*
1 Percent SEM followed by are not significantly different (P : 0.05) by ANOVA (n =
20 B. insularis of each respective age per vial; F = 2.02; df = 7, 72; P = 0.06).









Table 3-2. Mean percent mortality of Blissus insularis ( SEM) killed by different rates
of Bioblitz and diazinon at 2 and 4 days after treatment.
% Mortality
Treatment Rate 2 DAT 4 DAT

Untreated control ---- 12.0 0.7 23.0 0.6
Bioblitz 1 ppm 23.0 0.7 33.0 0.3
10 ppm 19.0 0.9 34.0 0.4
100 ppm 18.0 0.7 38.0 0.2 *
Diazinon 1 ppm 14.0 0.8 40.0 0.3 *
10 ppm 16.0 0.8 54.0 0.3 *
100 ppm 82.0 0.5 97.0 0.08 *

'Percent SEM followed by are significantly different (P <: 0.05) using Tukey's
Studentized Range (HSD) test (n = 10 B. insularis of each respective age per vial; 2
DAT: F= 8.28; df = 6, 63; P < 0.0001; 4DAT: F= 23.0; df = 6, 63; P < 0.0001).









Table 3-3. Mean percent survival of Blissus insularis ( SEM) at 1 and 4 wk post
treatment.

Treatment Rate 1 wk 4 wk

Control ---- 76.7 3.6 18.0 8.3

Bifenthrin 152.5 kg/ha 18.0 8.3*1 6.7 4.1

Carbaryl 38.2 L/ha 3.3 1.4* 0*

Cyfluthrin 19.1 L/ha 55.9 12.6 20.8 5.7

Deltamethrin 146.4 kg/ha 2.5 1.6* 5.0 1.7

k-cyhalothrin 146.4 kg/ha 9.0 2.1* 21.6 4.4
Permethrin 19.1 L/ha 39.0 10.4 8.3 5.0
1 Percent SEM followed by are significantly different (P <: 0.05) using Tukey's
Studentized Range (HSD) test (n = 4 reps. 1 wk: F= 13.23; df= 6, 21; P < 0.0001; 4
wk: F= 4.30; df = 6, 21; P = 0.006).









Table 3-4. Mean percent survival of Blissus insularis ( SEM) at 1 and 4 wk post
treatment.
Treatment Rate 1 wk 4 wk

Control --- 76.0 5.5 26.6 11.4

Bifenthrin 1.6 L/ha 1.0 0.7 1 *

Carbaryl 18.78 L/ha 0 0 *

Cypermethrin 2.1 L/ha 1.0 + 0.7 0.7 1.0 *

Cyfluthrin 0.86 L/ha 34.0 11.2* 9.3 4.0
Deltamethrin 146 kg/ha 3.0 + 2.0 1.3 1.0 *

k-cyhalothrin 1.5 L/ha 14.0 + 3.3 4.5 3.2 *
Permethrin 2.5 L/ha 7.3 4.4 2.7 2.7 *

1 Percent SEM followed by are significantly different (P <: 0.05) using Tukey's
Studentized Range (HSD) test (n = 5 reps. 1 wk: F = 26.28; df = 7, 32; P < 0.0001; 4
wk: F = 6.37; df = 7, 32; P < 0.001).






31















Figure 3-1. Methods used in pesticide field trials. A. PVC rings used in pesticide field
trials. B. Flotation of St. Augustinegrass post-treatment.














CHAPTER 4
ST. AUGUSTINEGRASS GROWTH RESPONSE TO THREE LEVELS OF
IRRIGATION AND Blissus insularis DENSITY

Introduction

Blissus insularis, an important pest of St. Augustinegrass, has been reported to

cause more damage and be more abundant in sunny, open, drought-stressed areas of

lawns (Kuitert and Nutter 1952, Reinert and Kerr 1973). Particularly susceptible areas

include turf near sidewalks, pavements, or in poorly irrigated areas. The nymphs and

adults live in the thatch and suck fluids from the crown and stem of grasses. This feeding

results in brown, dead patches of turf that are aesthetically displeasing and allow weed

encroachment. Because B. insularis populations are aggregated with up to 2,000/0.1 m2

(Reinert and Kerr 1973), turfgrass managers attempt to prevent outbreaks with frequent

insecticide applications. Currently, 20-25 B. ii\nli11 i% 0.1 m2 is considered enough to

warrant a treatment (Short et al. 1982).

Moisture has been reported to have a "marked but paradoxical" effect on B.

insularis (Kerr 1966). Adequately irrigated turf may be attractive to B. insularis or easier

to feed on. But rapidly growing grass may withstand the effects of feeding, and excess

moisture may actually suppress populations (Braman 1995). However, little is known

about the response of St. Augustinegrass to B. insularis feeding and the interaction with

irrigation. The objective of this research was to quantify St. Augustinegrass growth

response to three levels each of irrigation and B. insularis densities.









Materials and Methods

Insect Collection and Maintenance

Blissus insularis were collected from St. Augustinegrass lawns in Ocala, FL

(Marion Co.), using a modified Weed Eater Barracuda blower/vacuum (Electrolux Home

Products, Augusta, GA) and transported in mesh-covered buckets containing fresh

cuttings of Palmetto St. Augustinegrass to the laboratory. Grass was replaced as needed

and buckets were maintained at 800F and 13:11 (L:D) for <2 wk. Just before bioassay, B.

insularis were aspirated into plastic vials (2.5 x 10.2 cm) containing a cone-shaped,

moistened, 70 mm Whatman filter paper and a foam cap.

Bioassay. Forty-five sewer polyvinyl chloride (PVC) pipes (15.2 cm diameter x

43.2 cm long) fitted with sewer caps were filled with 7.6 cm of river rock, a piece of #4

Whatman filter paper, and Arrendondo fine sand up to 1.3 cm from the top (Fig. 4-1).

Holes were drilled into the sewer caps to allow drainage and the filter paper layer

prevented root and soil migration into the rock layer. Soil was allowed to settle for 24 h.

Forty-five 15.2 cm diameter plugs of Palmetto St. Augustinegrass were transplanted from

pots onto the top of the lysimeters on 14 Aug 2003 and lysimeters were placed on

reinforced metal platforms in a climate-controlled greenhouse at the G. C. Horn

Memorial Turfgrass Field Laboratory at the University of Florida in Gainesville, FL.

Daytime and nighttime temperatures were 270C and 240C, respectively. Plants were

fertilized with 16-4-8 water soluble complete N source NH4NO3 at 0.5 lb N/1000 ft2 and

allowed to establish for 1 mo before bioassay.

Treatments included irrigation [low (30%), medium (60%), or high (100%)

saturation] and either 0, 30, or 200 fourth-fifth instar B. insularis. There were five









replicates in a CRD. A mesh cage was placed 5.0 cm into the soil and around the plants,

such that the grass blades could be maintained at 7.6 cm height. Lysimeters were

completely saturated with 1,500 ml of water and allowed to drain for 24 h. After 24 h,

insects were transferred from vials using a camel-hair brush and cages were closed with

nylon. Each lysimeter was weighed to determine its initial saturation.

Amounts of water to apply at respective irrigation levels were determined by

replacing some fraction of the water used in evapotranspiration (ET). This was

determined gravimetrically by the following:

ET =Wmax Win

Needed = deficit irrigation level x ETcontrol

Wmax = Wmin + Wneeded

Wmin and Wmax were the lysimeter weights before and after water was applied. Wneeded

represented the water amount applied to lysimeters. ET,,ontro represented the mean of

100% irrigation levels of ET. ET rates were measured and plants were irrigated weekly.

After 2 mo, B. insularis were removed from lysimeters using a small, hand-held

Black & Decker Dust-BusterTM vacuum (Bioquip Products, Rancho Dominguez, CA)

equipped with a 12.7 cm long x 3.8 cm diameter hose attached to a removable collecting

chamber (12.7 cm long x 5.1 cm diameter). The vacuum was powered by a portable 12-

volt DC battery pack (Bioquip Products, Rancho Dominguez, CA). Each lysimeter was

vacuumed for 2 min, samples were placed into plastic bags, and the number of B.

insularis collected were counted in the laboratory.

To collect root data, lysimeters were emptied onto a screen table and the roots

were removed just below the crown. Crown contents were placed into buckets, covered









with mesh, and brought to the laboratory. Remaining B. insularis in the crown material

were removed by flotation and counted. Soil was washed off roots, and roots were

transported to the laboratory in paper bags. Roots were rinsed again in the laboratory

using a #20 standard testing sieve (Fisher Scientific Co.) and non-root debris were

removed. Wet and dry weights of roots were recorded. For initial dry weights, roots

were dried for 48 h in a Blue M Stabil-Therm mechanical convection oven with Pro-Set

II control at 550C. The dry ash procedure was done to obtain the percent organic matter

of the roots. Dry roots were placed in 150 ml Erlenmeyer beakers and baked at 4500C in

a Fisher Scientific Isotemp Muffle furnace (model #550-58). Dry ash weights were

subtracted from initial dry weights to obtain the percent organic matter of roots.

Grass blades were cut weekly to a 7.6 cm height, and clippings were collected and

transported to the laboratory in paper bags in a cooler to obtain fresh and dry weights.

For dry weights, grass blades were dried for 24 h in a Blue M Stabil-Therm mechanical

convection oven with Pro-Set II control at 550C.

An analysis of variance (ANOVA) was conducted to evaluate differences between

treatment means in root weights and weekly grass clippings. Data were analyzed as a

two-way factorial complete randomized design with irrigation and B. insularis as main

factors using the Student-Newman-Keuls test (SAS Institute, Inc. 2000).

Results and Discussion

Stunted growth from B. insularis feeding was observed from grass clippings

collected after 1 wk (Table 4-1). Dry weights of grass clippings collected from the

different B. insularis densities were different from one another except during weeks two,

six, and seven (Table 4-1). During weeks two, six, and seven, the dry weight of grass









clippings recovered from treatments containing 200 B. insularis were lower than dry

weights of grass clippings collected from treatments containing 0 or 30 B. insularis.

Blissus insularis feeding from densities of 30 and 200 lowered root weight as shown in

Table 4-2. Research on other Blissus spp. reports their feeding can cause wilting,

chlorosis, stunting, and death through clogging of vascular transport systems (Painter

1928, Negron and Riley 1990, Spike et al. 1991). Beyer (1924) observed that large

numbers of B. insularis created a 'dwarfed condition' of St. Augustinegrass, eventually

leading to plant death.

The dry weights of grass clippings collected from treatments with different

irrigation levels were different during weeks five and seven. At week five, the dry

weight of grass clippings collected from treatments with low irrigation were lower than

the dry weights of grass clippings collected from treatments containing medium or high

irrigation (Table 4-1). At week seven, dry weights of grass clippings collected from

treatments with low irrigation were lower than clippings from treatments with high

irrigation levels. Irrigation did not affect root weight (Table 4-2). The interaction

between irrigation and B. insularis was not significant for grass clipping (F= 0.75; df=

4, 39; P = 0.57) or root weight data (F= 0.54; df = 4, 39; P = 0.71). It is possible that B.

insularis populations are found first in drought-stressed areas in the lawn because of an

increased chance of survival. Heavy rainfall and irrigation have been observed to drown

early instar Blissus spp. (Luginbill 1922, Beyer 1924, Wilson 1929, Kuitert and Nutter

1952). Also, environmental factors such as temperature and light may play a role in

where B. insularis outbreaks occur first in the lawn.














St. Augustinegrass
enclosed in mesh cage





Arrendondo soil





River rock filled up to
sewer cap and filter
paper placed on top of
rock


Figure 4-1. Lysimeter used in experiment.














Table 4-1. Mean dry weight (mg) ( SEM) of grass clippings by week. 1
Main


Efc Level 1 wk 2 wk 3 wk 4 wk 5 wk 6 wk 7 wk 8 wk
Effect
B. 0 400.0 20.0a 320.0 20.0a 270.0 60.0a 180.0 30.0a 170.0 30.0a 130.0 20.0a 110.0 20.0a 110.0 20.0a
insularis
30 330.0 20.0b 280.0 20.0a 190.0 60.0b 140.0 40.0b 140.0 50.0b 110.0 30.0a 100.0 20.0a 80.0 20.0b

200 230.0 + 20.0c 140.0 10.0b 60.0 1 10.0 40.0 + 3.00c 20.0 4.00c 30.0 + 6.00b 20.0 + 3.00b 20.0 4.00c

F-value 17.98 34.78 59.14 37.89 72.86 38.25 30.61 26.03

P-value < 0.0001 < 0.0001 < 0.0001 < 0.0001 <0.0001 <0.0001 <0.0001 <0.0001

Irrigation low 330.0 30.0" 240.0 20.0a 160.0 10.0a 110.0 0 90.0 1.00b 70.0 0 60.0 0.30b 60.0 2.00a

med 310.0 + 20.0a 250.0 + 30.0a 180.0 10.0a 120.0 5.00a 120.0 10.0a 100.0 3.00a 80.0 + 3.00ba 70.0 10.0a

high 320.0 + 20.0a 250.0 + 20.0a 190.0 10.0a 130.0 4.00a 130.0 4.00a 100.0 6.00a 90.0 3.00a 70.0 4.00a

F-value 0.39 0.12 2.12 1.55 6.88 2.90 3.18 26.0

P-value 0.68 0.88 0.13 0.23 0.003 < 0.07 < 0.05 0.10

1Means SEM within a column followed by the same letter are not significantly different (P <: 0.05) by the Student-Newman-Kuels
test.









Table 4-2. Mean root weight ( SEM) of St. Augustinegrass after 8 wk.
Treatment Level Mean root wt (mg)
B. insularis 0 2100.0 140.0
30 1600.0 170.0 *1
200 1300.0 130.0 *
Irrigation low 1700.0 200.0
med 1400.0 130.0
high 1800.0 150.0


1Mean root weight SEM followed by are significantly different (P <: 0.05) using
Student-Newman-Kuels test. (B. insularis: F = 7.33; df = 2, 42; P = 0.002. Irrigation: F
= 1.81; df = 2, 42; P = 0.18).
















LIST OF REFERENCES


Backus, E. A., W. B. Hunter, and C. N. Arne. 1988. Technique for staining
leafhopper salivary sheaths and eggs within unsectioned plant tissue. J. Econ.
Entomol. 81: 1819-1823.

Banks, W. A., C. S. Lofgren, D. P. Jouvenaz, C. E. Stringer, P. M. Bishop, D. F.
Williams, D. P. Wojcik, and B. M. Glancey. 1981. Techniques for collecting,
rearing, and handling imported fire ants. U.S. Dept. Agric. Tech. AAT-S-21.

Baxendale, F. P., T. M. Heng-Moss, and T. P. Riordan. 1999. Blissus occiduus
(Hemiptera: Lygaeidae): A chinch bug pest new to buffalograss turf. J. Econ.
Entomol. 92: 1172-1176.

Bell, K. 0., and W. H. Whitcomb. 1964. Field studies on egg predators of the
bollworm, Heliothis zea (Boddie). Fla. Entomol. 47: 171-180.

Beyers, A. H. 1924. Chinch bug control on St. Augustinegrass. Proc. Fla. State Hort.
Soc. 37: 216-219.

Braman, S. K. 1995. Insect/plant stress interactions, pp. 112-114. In R. L.
Brandenburg and M. G. Villani (eds.), Handbook of turfgrass insect pests.
Entomological Society of America, Lanham, MD.

Brandenburg, R. L. and M. G. Villani. 1995. Handbook of turfgrass insect pests.
Entomological Society of America, Lanham, MD.

Britcher, W. H. 1903. The chinch bug in Maine. Maine Agr. Exp. Sta. Bull. No. 91.

Brogdon, J. E. and S. H. Kerr. 1961. Home gardner's lawn insect control guide. Fla.
Univ. Agric. Ext. Serv. Cir. No. 213.

Bruton, B. D., R. W. Toler & J. A. Reinert. 1983. Combined resistance in St.
Augustinegrass to the southern chinch bug and the St. Augustinegrass decline strain
of panicum mosaic virus. Plant Disease. 67: 171-172.

Busey, P. 1979. What is Floratam? Proc. Fla. State Hort. Soc. 92: 228-232.

Busey, P. 1990a. Inheritance of host adaptation in the southern chinch bug (Hemiptera:
Lygaeidae). Ann. Entomol. Soc. Am. 83: 563-567.









Busey, P. 1990b. Polyploid Stenotaphrum D. L. germplasm resistance to the polyploid
damaging population southern chinch bug (Hemiptera: Lygaeidae). Crop Sci. 30:
588-593.

Busey, P. and B. Center. 1987. Southern chinch bug (Hemiptera: Heteroptera:
Lygaeidae) overcomes resistance in St. Augustinegrass. J. Econ. Entomol. 80: 608-
611.

Busey, P. and G. H. Snyder. 1993. Population outbreak of the southern chinch bug is
regulated by fertilization. Int. Turf. Soc. Res. J. 7: 353-357.

Buss, E. A. 2001. Insect pest management on golf courses. Florida Cooperative
Extension Service, Institute of Food and Agricultural Sciences, University of
Florida, Gainesville. ENY351.

Buss, E. A. and D. L. Caldwell. 2001. Biology and management of tropical sod
webworms. Florida Cooperative Extension Service, Institute of Food and
Agricultural Sciences, University of Florida, Gainesville. ENY318.

Carstens, J. D. 2003. Influence of buffalograss management practices on the chinch
bug, Blissus occiduus Barber, and its natural enemies. Master's Thesis. University
of Nebraska, Lincoln.

Champlain, R. A. and L. L. Sholdt. 1967a. Life history of Geocorispunctipes
(Hemiptera: Lygaeidae) in the laboratory. Ann. Entomol. Soc. Am. 60 (5): 881-
883.

Champlain, R. A. and L. L. Sholdt. 1967b. Temperature range for development of
immature stages of Geocorispunctipes (Hemiptera: Lygaeidae). Ann. Entomol.
Soc. Am. 60 (60): 883-885.

Cherry, R. H. 2001a. Seasonal wing polymorphism in southern chinch bugs
(Hemiptera: Lygaeidae). Fla. Entomol. 84: 737-739.

Cherry, R. H. 2001b. Spatial distribution of southern chinch bugs (Hemiptera:
Lygaeidae) in St. Augustinegrass. Fla. Entomol. 84: 151-153.

Cherry, R. H. 2001c. Interrelationship of ants (Hymenoptera: Formicidae) and southern
chinch bugs (Hemiptera: Lygaeidae) in Florida lawns. J. Entomol. Sci. 36 (4): 411-
415.

Cherry, R. H. and R. T. Nagata. 1997. Ovipositional preference and survival of
southern chinch bugs (Blissus insularis Barber) on different grasses. Int. Turf. Soc.
J. 8: 981-986.

Crocker, R. L. 1993. Chemical control of southern chinch bug in St. Augustinegrass.
Int. Turf. Soc. Res. J. 7: 358-363.









Crocker R. L. and W. H. Whitcomb. 1980. Feeding niches of the big-eyed bugs
Geocoris bullatus, G. punctipes, and G. uliginosus (Hemiptera: Lygaeidae:
Geocorinae). Entomol. Soc. Am. 9 (5): 508-513.

Crocker, R. L. and C. L. Simpson. 1981. Pesticide screening test for the southern
chinch bug. J. Econ. Entomol. 74: 730-731.

Davis, M. G. K, and D. R. Smitley. 1990. Association of thatch with populations of
hairy chinch bug (Hemiptera: Lygaeidae) in turf J. Econ. Entomol. 83: 2370-2374.

Dumas, B. A., W. P. Boyer, and W. H. Whitcomb. 1962. Effect of time of day on
surveys of predaceous insects in field crops. Fla. Entomol. 45: 121-128.

Dunbar, D. M. 1971. Big-eyed bugs in Connecticut lawns. Conn. Agric. Exp. Stn. Cir.
No. 244.

Haydu, J. J., L. N. Satterthwaite, and J. L. Cisar. 1998. An economic and agronomic
profile of Florida's sod industry in 1996. Food and Resource Economics
Department, Agricultural Experiment Stations and Cooperative Extension Service,
Institute of Food and Agricultural Sciences, University of Florida, Gainesville.

Heng-Moss, T. M., F. P. Baxendale, and T. P. Riordan. 1998. Beneficial arthropods
associated with buffalograss. J. Econ. Entomol. 91(5): 1167-1172.

Heng-Moss, T. M., F. P. Baxendale, T. P. Riordan, and J. E. Foster. 2002.
Evaluation of buffalograss germplasm for resistance to Blissus occiduus
(Hemiptera: Lygaeidae). J. Econ. Entomol. 95: 1054-1058.

Henry, T. J. and R. C. Froeschner [eds.]. 1988. Catalog of the Heteroptera or True
Bugs of Canada and the continental United States. E. J. Brill, Leiden.

Hodges, A. W., J. J. Haydu, P. J. van Blokland, and A. P. Bell. 1994. Contribution of
the turfgrass industry to Florida's economy, 1991-92: A value-added approach.
Economics Report ER 94-1, Food and Resource Economics Department Institute of
Food and Agricultural Sciences, University of Florida, Gainesville.

Horn, D. J. 1988. Ecological approach to pest management. Guilford Press, New York.

Hutchins, S. 1994. Techniques for sampling arthropods in integrated pest management,
pp. 73-98. In Larry P. Pedigo and G. David Buntin (eds.), Handbook of sampling
methods for arthropods in agriculture. CRC Press, Inc., Boca Raton, Florida.

Janes, M. J., A. Hager, and G. E. Carman. 1935. Preliminary studies on starvation
and drowning of the chinch bug, Blissus leucopterus (Say). J. Econ. Entomol. 28:
638-646.

Kelsheimer, E. G. 1952. Insects and other pests of lawns and turf Fla. Agric. Exp. Sta.
Cir. S-42.






43


Kerr, S. H. 1956. Chinch bug control on lawns in Florida. J. Econ. Entomol. 49: 83-85.

Kerr, S. H. 1958. Tests on chinch bugs and the current status of controls. Proc. Fla.
Hort. Soc. 71: 400-403.

Kerr, S. H. 1961. Lawn chinch bug research. Proc. Univ. Fla. Turf. Man. Conf. 9: 211-
221.

Kerr, S. H. 1966. Biology of the lawn chinch bug, Blissus insularis. Fla. Entomol.
49(1): 9-18.

Khuhro, R. D. 1994. Biological studies on the chinch bug, Blissus leucopterus
leucopterus (Say) in northeast Mississippi. Ph.D. dissertation. Mississippi State
University, Starkville.

Knowlton, G. F. 1935. Beet leafhopper predator studies. Proc. Utah Acad. Sci. Arts.
Letters 12: 255-60.

Kuitert, L. C., and G. C. Nutter. 1952. Chinch bug control and subsequent renovation
of St. Augustinegrass lawns. Univ. Fla. Agric. Exp. Sta. Cir. S-50.

Leonard, D. E. 1966. Biosystematics of the leucopteruss complex" of the genus Blissus
(Heteroptera: Lygaeidae). Conn. Agric. Exp. Stn. Bull. No. 677.

Leonard, D. E. 1968. A revision of the genus Blissus (Heteroptera: Lygaeidae). Ann.
Entomol. Soc. Am. 61: 239-250.

Luginbill, P. 1922. Bionomics of the chinch bug. U. S. Dept. Agric. Bull. No. 1016.

Majeau, G., J. Brodeur, and Y. Carriere. 2000. Sequential sampling plans for the
hairy chinch bug (Hemiptera: Lygaeidae). J. Econ. Entomol. 93: 834-839.

McCarty, L. B. and J. L. Cisar. 1997. St. Augustinegrass for Florida lawns, pp. 12-13.
In Kathleen C. Ruppert and Robert J. Black (eds.), Florida lawn handbook, 2nd ed.
Department of Environmental Horticulture, Institute of Food and Agricultural
Sciences, University of Florida, Gainesville.

McColloch, J. W., and N. Yuasa. 1914. A parasite of the chinch bug egg. J. Econ.
Entomol. 14: 219-227.

McGregor, E. A., and F. L. McDonough. 1917. The red spider on cotton. U. S. Dept.
Agric. Bull. No. 416.

Mead, F. W. 1972. Key to the species of big-eyed bugs, Geocoris spp., in Florida
(Hemiptera: Lygaeidae). Fla. Dept. Agric. Cons. Serv., Division of Plant Industry.
ENY Cir. No.121.






44


Miller, N. C. E. 1971. The biology of the Heteroptera, 2nd ed. Classey Ltd, Hampton,
UK.

Nagata, R. T. and R. H. Cherry. 1999. Survival of different life stages of the southern
chinch bug (Hemiptera: Lygaeidae) following insecticidal applications. J.
Entomol. Sci. 34: 126-131.

Negr6n, J. F., and T. J. Riley. 1990. Long-term effects of chinch bug (Hemiptera:
Lygaeidae) feeding on corn. J. Econ. Entomol. 83: 618-620.

Niemczyk, H.D., R. A. J. Taylor, M. P. Tolley, and K T. Power. 1992. Physiological
time-driven model for predicting first generation of the hairy chinch bug
(Hemiptera: Lygaeidae) on turfgrass in Ohio. J. Econ. Entomol. 85: 821-829.

Painter, R. H. 1928. Notes on the injury to plant cells in chinch bug feeding. Ann.
Entomol. Soc. Am. 21: 232-241.

Potter, D. 1998. Destructive Turfgrass Insects: Biology, Diagnosis, and Control. Ann
Arbor Press. Chelsea, MI.

Readio, J. and M. H. Sweet. 1982. A review of the Geocorinae of the United States
East of the 100th Meridian (Hemiptera: Lygaeidae). Misc. Publ. Entomol. Soc. Am.
12: 1-91.

Reinert, J. A. 1972a. New distribution and host record for the parasitoid Eumicrosoma
benefica. Fla. Entomol. 55(3): 143-144.

Reinert, J. A. 1972b. Control of the southern chinch bug, Blissus insularis, in south
Florida. Fla. Entomol. 55: 231-235.

Reinert, J. A. 1978. Natural enemy complex of the southern chinch bug in Florida.
Ann. Entomol. Soc. Am. 71: 728-731.

Reinert, J. A. 1982. Carbamate and synthetic pyrethroid insecticides for control of
organophosphate-resistant southern chinch bugs (Heteroptera: Lygaeidae). J. Econ.
Entomol. 75: 716-718.

Reinert, J. A. and S. H. Kerr. 1973. Bionomics and control of lawn chinch bug. Bull.
Entomol. Soc. Am. 19: 91-92.

Reinert, J. A. and A. E. Dudeck. 1974. Southern chinch bug resistance in St.
Augustinegrass. J. Econ. Entomol. 67: 275-277.

Reinert, J. A. and H. D. Niemczyk. 1982. Insecticide resistance in epigeal insect pests
of turfgrass. II. Southern chinch bug resistance to organophosphates in Florida,
pp. 77-80. In H. D. Niemczyk and B. G. Joyner (eds.), Advances in turfgrass
entomology. Hammer Graphics, Piqua, OH.









Reinert, J. A. and K. Portier. 1983. Distribution and characterization of
organophosphate-resistant southern chinch bugs (Heteroptera: Lygaeidae) in
Florida. J. Econ. Entomol. 76: 1187-1190.

Richmond, D. S. and D. J. Shetlar. 2000. Hairy chinch bug (Hemiptera: Lygaeidae)
damage, population density, and movement in relation to the incidence of perennial
ryegrass infected by Neotyphodium endophytes. J. Econ. Entomol. 93: 1167-1172.

SAS Institute, Inc. 2000. SAS user's guide statistics, version 8.2. SAS Institute Inc.,
Cary, NC .

SAS Institute, Inc. 2001. Jmp In: Statistical discovery software. SAS Institute Inc.,
Cary, N.C.

Short, D. E. and P. G. Koehler. 1979. A sampling technique for mole crickets and
other pests in turfgrass and pasture. Fla. Entomol. 62: 282-283.

Short, D. E., J. A. Reinert and R. A. Atilano. 1982. Integrated pest management for
urban turfgrass culture Florida. pp. 25-30. In H.D. Niemcyk and B.G. Joyner
(eds.) Advances in turfgrass entomology. Hammer Graphics, Inc., OH.

Slater, J. A., and R. M. Baranowski. 1990. Lygaeidae of Florida (Hemiptera:
Heteroptera). Vol 14. Arthropods of Florida and neighboring land areas. Fla.
Dept. Agric. Cons. Serv., Gainesville, Florida.

Southwood, T. R. E. 1978. Ecological methods. Halsted, New York.

Spike, B. P., R. J. Wright, S. Danielson, and D. W. Stanley-Samuelson. 1991. The
fatty acid compositions of phospholipids and triacyglycerols from two chinch bug
species Blissus leucopterus leucopterus and Blissus iowensis (Hemiptera:
Lygaeidae) are similar in the characteristic dipteran pattern. Comp. Biochem.
Physiol. 998: 799-802.

Sprenkel, R. K. 2003. Insect management in pasture. Florida Cooperative Extension
Service, Institute of Food and Agricultural Sciences, University of Florida.
ENY402.

Standen, V. 2000. The adequacy of collecting techniques for estimating species
richness of grassland invertebrates. J. Appl. Ecol. 37: 884-893.

Stoner, A. 1970. Plant feeding by a predaceous insect, Geocorispunctipes. J. Econ.
Entomol. 63: 289-298.

Sweet, M. H. 2000. Seed and chinch bug (Lygaeoidea), pp. 143-264. In Carl W.
Schaefer and Antonio Ricardo Paninzzi (eds.), Heteroptera of economic
importance. CRC Press LLC, Boca Raton, Florida.









T6rmiila, T. 1982. Evaluation of five methods of sampling field layer arthropods,
particularly the leafhopper community in grasslands. Annales Entomologicae
Fennici, 48: 1-16.

Trenholm, L. E., J. L. Cisar & J. B. Unruh. 2000. St. Augustinegrass for Florida
lawns. Florida Cooperative Extension Service, Institute of Food and Agricultural
Sciences, University of Florida, Gainesville. ENH5.

Trenholm, L. E., J. L. Cisar & J. B. Unruh. 2001. Thatch and its control in Florida
lawns. Florida Cooperative Extension Service, Institute of Food and Agricultural
Sciences, University of Florida, Gainesville. ENH12.

Trenholm, L. E., J. L. Cisar & J. B. Unruh. 2003. Let your lawn tell you when to
water. Florida Cooperative Extension Service, Institute of Food and Agricultural
Sciences, University of Florida, Gainesville. ENH63.

Turgeon, A. J. 1996. Turfgrass management, 4th ed. Prentice Hall, Upper Saddle River,
NJ.

U.S. Census Bureau. 2000. General Housing Characteristics: 2000. Found in Census
2000 Summary File 1, Matrices H3, H4, H5, H6, H7, and H16.

U.S. Census Bureau. 2003. State rankings: Statistical abstract of the United States.
http://www.census.gov/statab/ranks/rank01.html, March 2004.

Vittum, P. J., M. G. Villani, and H. Tashiro. 1999. Turfgrass insects of the United
States and Canada, 2nd ed. Comstock Publishing Assoc., Ithaca.

Watson, J. R. and H. E. Bratley. 1929a. The chinch bug on St. Augustinegrass lawns.
Fla. Agric. Exp. Sta. Press Bull. No. 409.

Watson, J. R.,and H. E. Bratley. 1929b. The chinch bug on St. Augustinegrass. Fla.
Agric. Exp. Sta. Bull. No. 209: 18-20.

Webster, F. M. 1907. The chinch bug. U. S. Dept. Agric. Bull. No. 69.

White, W. W. and P. Busey. 1987. History of turfgrass production in Florida. Proc.
Fla. State Hort. Soc. 100: 167-174.

Wilson, R. N. 1929. The chinch bug in relation to St. Augustinegrass. U. S. Dept.
Agric. Cir. No. 51.

Wolfenbarger, D. 0. 1953. Insect and mite control problems on lawn and golf grasses.
Fla. Entomol. 36: 9-12.

York, G. T. 1944. Food studies of Geocoris spp., predators of the beet leafhopper. J.
Econ. Entomol. 37: 25-29.















BIOGRAPHICAL SKETCH

Julie Cara Congdon was born on October 25, 1970, in St. Petersburg, Florida.

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 College and

after taking several honors courses developed an interest in entomology. In 1998, Cara

entered the University of Florida as an undergraduate entomology major. While at the

University 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

Florida's Entomology and Nematology Department (urban entomology laboratory),

United States Department of Agriculture (USDA), and FMC Corporation. She is a

research associate for the Division of Plant Industry and is a member of the

Entomological Society of America, Florida Entomological Society, Florida Turfgrass

Association, Certified Pest Control Operators of Florida, Entomology and Nematology

Student Organization (ENSO) and the Urban Entomological Society (UES). Cara plans

to pursue a Ph.D. in entomology at the University of Florida.