<%BANNER%>

Management of Pest Mole Crickets Using the Insect Parasitic Nematode Steinernema scapterisci


PAGE 1

MANAGEMENT OF PEST MOLE CRICK ETS USING THE INSECT PARASITIC NEMATODE Steinernema scapterisci By KATHRYN ANN BARBARA A DISSERTATION PRESENTED TO THE GRADUATE SCHOOL OF THE UNIVERSITY OF FLOR IDA IN PARTIAL FULFILLMENT OF THE REQUIREMENTS FOR THE DEGREE OF DOCTOR OF PHILOSOPHY UNIVERSITY OF FLORIDA 2005

PAGE 2

Copyright 2005 by Kathryn Ann Barbara

PAGE 3

This dissertation is dedicated to my family, Kathleen and Jack Barbara, Anthony, Lara, and Elizabeth Barbara, Lisa, Eric and Ry an Malkowski, Anne Barbara, Ann and Raymond Callahan, John Michael, Kathlee n, Melinda, and Maureen Gowdey, Camilla Barbara, James Dunford, and Magnum, the delightful Doberman. Without their love, thoughtfulness and support this would not have been possible.

PAGE 4

ACKNOWLEDGMENTS My heartfelt thanks go to my major advisor, Dr. Eileen Buss. Her guidance, advice, and trust have made this research possible. Dr. Buss has been a mentor in not only my academic life but my personal life as well. I consider myself lucky to have been her first Ph. D. student. I would also like to thank my committee members, Drs. J. Howard Frank, Norman C. Leppla, Grady L. Miller, and especially Khuong B. Nguyen, who has given me the knowledge about nematology to make my degree truly a work of entomology and nematology. I could not have had a better committee. Each has contributed significantly to this work and it would not have been possible without each and every member. This research would not have been possible without the help of many people. I would like to thank the cooperators and their assistants who allowed me to use their sites and were very helpful during my research. Special thanks go to Buddy Keene II, Lloyd J. Brown and Diane Delzell from Gainesville Golf and Country Club; Dave Carson, Kevin Cooke, and Vern Easter from Ironwood Golf Course; Fred Santana from Sarasota County Extension; and Matt Burke from the City of Altamonte Springs. Nematodes and advice on their use were provided by Tom Hinks, Gabe Diaz-Saavedra, and Al Clarke from Becker Underwood and Martin Adjei of the Ona Range Cattle REC. I acknowledge Dr. Tom Walker for mole cricket advice and assistance. I acknowledge Dr. Albrecht Koppenhfer from Rutgers University for technical advice and methodology for the pesticide compatibility tests. Special thanks go to John iv

PAGE 5

Fredricks for help with soil sample analysis, Marinela Capanu for help with statistical design and analysis, and Paul Skelley (DPI) for help with SEM photographs. I acknowledge Bayer Environmental Science, Valent Professional Products, and FMC Corporation for donating insecticides used in this study. Special thanks go to Brian Owens at the G. C. Horn Memorial Turfgrass Field Laboratory for his assistance and patience with setting up nightly collections with mole cricket sound callers. Angela Vincent and Shubin Saha assisted in sorting linear pitfall trap samples. I would also like to acknowledge the electronic thesis and dissertation technical staff for their help in formatting and submission of my dissertation. Many thanks go to the Florida Department of Agriculture and the United States Navy for graduate funding. Very special thanks go to the numerous people who helped in my various projects; without them this project would not have been possible. First I would like to thank Paul Ruppert for his assistance in numerous aspects of this study and his patience with me. I also would like to thank Bob Hemenway for rearing supplies, guidance and advice. My gratitude goes to Matthew Stanton and Y. Mike Wang for help with bi-weekly pitfall trap collections and mole cricket rearing. I thank J. Cara Congdon, Jay Cee Turner, Lois Wood, Rebecca Baldwin, Erin Finn, Alejandro Arevalo, Justin Emerson and Brian McElroy for assistance in pitfall trap installation and removal. Special thanks go to Robert Mans and Rachel Davis for help with mole cricket rearing and the dreaded curfew field test. Many thanks go to the University of Florida Entomology and Nematology Department for their support and friendship through the years. Several people I have met along this journey have become more than just my academic and professional peers but have become very good friends. I want to give v

PAGE 6

special thanks and warm gratitude to J. Cara Congdon and Jay Cee Turner who, while helping me become a better scientist by looking towards me for guidance and advice, have become two of my best friends. I am especially grateful to James Dunford. He has inspired me in all aspects of my life and several ideas in this dissertation were inspired and motivated by him. Jim has given me the opportunity to discover other aspects of entomology outside of my field. My only regret is that I did not meet him sooner. vi

PAGE 7

TABLE OF CONTENTS page ACKNOWLEDGMENTS .................................................................................................iv LIST OF TABLES ...............................................................................................................x LIST OF FIGURES ..........................................................................................................xii ABSTRACT ....................................................................................................................xiii CHAPTER 1 INTRODUCTION AND REVIEW OF LITERATURE..............................................1 Mole Crickets................................................................................................................1 Management Practices..................................................................................................4 Chemical Control...................................................................................................4 Non-Chemical Control..........................................................................................5 Biological Control.................................................................................................5 Insect Parasitic Nematodes....................................................................................6 Objectives...................................................................................................................10 2 ESTABLISHMENT AND SPREAD OF Steinernema scapterisci ON FLORIDA GOLF COURSES.......................................................................................................12 Materials and Methods...............................................................................................13 Study Sites...........................................................................................................13 Mole Cricket Monitoring.....................................................................................13 Laboratory Assay.................................................................................................15 Statistical Analysis..............................................................................................15 Results and Discussion...............................................................................................15 3 SURVIVAL AND INFECTIVITY OF Steinernema scapterisci AFTER CONTACT WITH SOIL DRENCH SOLUTIONS.......................................................................25 Materials and Methods...............................................................................................27 Nematodes and Mole Crickets.............................................................................27 Bioassay...............................................................................................................27 Statistical Analysis..............................................................................................30 Results and Discussion...............................................................................................30 vii

PAGE 8

4 INTEGRATION OF INSECT PARASITIC NEMATODES WITH INSECTICIDES FOR CONTROL OF PEST MOLE CRICKETS........................................................36 Materials and Methods...............................................................................................37 Survival and Infectivity of S. scapterisci After Exposure to Pesticides..............38 Nematode Infectivity After Exposure to Pesticide Treated Mole Crickets.........39 Statistical Analysis..............................................................................................40 Results and Discussion...............................................................................................40 5 EFFECT OF Steinernema scapterisci NGUYEN AND SMART EXPOSURE ON MOLE CRICKET TUNNELING, OVIPOSITION, AND AVOIDANCE BEHAVIOR................................................................................................................46 Materials and Methods...............................................................................................47 Nematode Infection and Nematode Treated Areas Effect on Mole Cricket Tunneling Behavior.........................................................................................48 Oviposition Behavior of Mole Crickets Exposed to S. scapterisci.....................50 Y-Tube Tests.......................................................................................................50 Observation Chamber...................................................................................50 Response to S. scapterisci or Pesticides.......................................................51 Statistical Analysis..............................................................................................52 Results.........................................................................................................................52 Nematode Infection and Nematode Treated Areas Effect on Mole Cricket Tunneling Behavior98 ....................................................................................52 Oviposition Behavior of Mole Crickets Exposed to S. scapterisci.....................53 Y-Tube Tests.......................................................................................................53 Discussion...................................................................................................................54 6 SUMMARY AND CONCLUSIONS.........................................................................61 APPENDIX A AMBIENT DATA AND TURFGRASS QUALITY DATA COLLECTED AT GAINESVILLE GOLF AND COUNTRY CLUB AND IRONWOOD GOLF COURSE.....................................................................................................................63 B DATA FROM ATHLETIC FIELD DEMONSTRATION SITES.............................66 Materials and Methods...............................................................................................66 Objective..............................................................................................................66 Study Site.............................................................................................................66 Mole Cricket Monitoring.....................................................................................66 Results and Discussion...............................................................................................68 C PRELIMINARY CHECKLIST OF ARTHROPODS ASSOCIATED WITH GOLF COURSE TURFGRASS............................................................................................71 viii

PAGE 9

D SCANNING ELECTRON MICROGRAPH PICTURES OF MOLE CRICKET SENSORY AREAS....................................................................................................73 LIST OF REFERENCES...................................................................................................76 BIOGRAPHICAL SKETCH.............................................................................................87 ix

PAGE 10

LIST OF TABLES Table page 2-1. Percent infection of Scapteriscus spp. mole crickets collected from sites treated with Steinernema scapterisci............................................................................................24 3-1. Mean nematode mortality and percent of mole crickets infected with Steinernema scapterisci after exposure for 24 h to various drenching solutions..........................34 3-2. Mean nematode mortality and infectivity after exposure for 24 h to various drenching solutions..................................................................................................34 3-3. Mean number of mole crickets emerging from bermudagrass using various drenching solutions in May 2003.............................................................................35 3-4. Percent nematode infection from mole crickets exposed to treatment solutions 1, 5, 8, 12, or 24 h post infection......................................................................................35 4-1. Percent survival, infectivity, and days until death of S. scapterisci incubated in solutions of insecticides for 24 h..............................................................................44 4-2. Average days until death and percent infectivity by S. scapterisci nematodes to mole crickets exposed to insecticides................................................................................45 5-1. Oviposition of Scapteriscus mole crickets directly infected with different numbers of S. scapterisci........................................................................................................59 5-2. Oviposition of Scapteriscus mole crickets in sand treated with S. scapterisci..........59 5-3. Response of Scapteriscus vicinus, S. borellii, and S. abbreviatus to Steinernema scapterisci nematodes versus sterilized sand...........................................................59 5-4. Response of Scapteriscus vicinus to Steinernema scapterisci nematodes versus pesticides treated sand..............................................................................................60 A-1. Ambient data collected from Gainesville Golf and Country Club on dates of mole crickets collections...................................................................................................63 A-2. Ambient data collected from Ironwood Golf Course on dates of mole crickets collections.................................................................................................................64 x

PAGE 11

A-3. Average turfgrass density ratings for treated and untreated plots on Gainesville Golf and Country Club and Ironwood Golf Course.........................................................65 xi

PAGE 12

LIST OF FIGURES Figure page 2-1. Linear pitfall trap used to collect mole crickets........................................................21 2-2. 3.8 L catch bucket inside 19 L bucket of linear pitfall trap.......................................21 2-3. Mean monthly percent infection of mole crickets collected in pitfall traps at Ironwood Golf Course from areas treated with Steinernema scapterisci. .............22 2-4. Mean monthly percent infection of mole crickets collected in pitfall traps at Gainesville Golf and Country Club from areas treated with Steinernema scapterisci.................................................................................................................23 5-1. Distance tunneled through sand in Plexiglas arenas every 2 d by mole crickets exposed to varying amounts of nematodes..............................................................57 5-2. Comparison of total tunnel distances by male and female Scapteriscus spp. mole crickets after exposure to S. scapterisci...................................................................58 5-3. Distance tunneled by mole crickets through sand treated with nematodes and untreated sand in Plexiglas arenas over 48 h............................................................58 B-1. Average monthly infection rates at the Sarasota athletic field research site.............69 B-2. Average monthly infection rates at the Altamonte Springs athletic field research site............................................................................................................................66 D-1. SEM photograph of a female Scapteriscus vicinus antennal mid-section................73 D-2. SEM photograph of a male Scapteriscus vicinus antennal mid-section...................74 D-3. SEM photograph of a female Scapteriscus vicinus mid-tarsal claw.........................74 D-4. SEM photograph of a female Scapteriscus vicinus labial palpomere.......................75 xii

PAGE 13

Abstract of Dissertation Presented to the Graduate School of the University of Florida in Partial Fulfillment of the Requirements for the Degree of Doctor of Philosophy MANAGEMENT OF PEST MOLE CRICKETS USING THE INSECT PARASITIC NEMATODE Steinernema scapterisci By Kathryn Ann Barbara May 2005 Chair: Eileen A. Buss Major Department: Entomology and Nematology Steinernema scapterisci Nguyen and Smart nematodes became established on two golf courses in Gainesville, FL, when applied as an augmentative application, and moved to untreated areas. It took about 4-8 wk post application for infection of mole crickets (Scapteriscus spp.) to equal or exceed pretreatment levels. Infection levels in untreated areas at least 80 m away from treated areas reached infection levels similar to the treated areas in approximately 20 wk post application. After a nematode application, mole crickets are frequently assayed to confirm nematode establishment. However, the standard soap flush was suspected of providing false negatives under field conditions. Thus, we examined the effect of several potential flushing solutions on the survival and infectivity of S. scapterisci as well as flushing ability under field conditions. Seventy percent of S. scapterisci died in the lemon dish detergent solution, confirming that assays for nematode infection of soap-flushed mole crickets are likely to be inaccurate. When sampling for mole crickets in areas where S. xiii

PAGE 14

scapterisci has been applied, a potential alternative to the standard soap drench is a dilute permethrin drench. Aqueous solutions of pesticides (acephate, bifenthrin, deltamethrin, fipronil and imidacloprid) used in turfgrass to control mole crickets were tested for compatibility with S. scapterisci in the laboratory. Survival of S. scapterisci was >95% in solutions of acephate, bifenthrin and imidacloprid. Infectivity of S. scapterisci previously exposed to insecticides was >60% in acephate and bifenthrin; however, infectivity was <40% in imidacloprid. The entomopathogenic nematode was compatible with all insecticides tested. Both healthy and nematode-infected mole crickets had similar tunneling behavior. Although not significant, crickets treated with 500 or 10,000 nematodes tunneled less than uninfected crickets. Mole crickets did not appear to differentiate between untreated and nematode-treated sand. Female crickets infected with nematodes were able to lay eggs, and clutch size and egg chamber size were not significantly different than healthy crickets. Crickets also laid eggs in sand treated with nematodes, suggesting that the nematode treated sand was not a deterrent. Mole crickets in Y-tube tests did not significantly choose untreated sand over sand treated with 500 or 10,000 nematodes. When given a choice between field rates of S. scapterisci and acephate, bifenthrin, deltamethrin, fipronil, or imidacloprid, crickets significantly chose nematodes over insecticides. xiv

PAGE 15

CHAPTER 1 INTRODUCTION AND REVIEW OF LITERATURE Mole Crickets Exotic mole crickets (Scapteriscus spp.) are the most injurious insect pests of golf courses, lawns, sod farms and pastures in Florida and throughout the southeastern United States (Walker and Nickle 1981). Mole crickets damage turf by tunneling in the soil which exposes and dries out roots and by direct root feeding. As a result, the turfgrass thins and bare patches appear. Weeds may invade these patches, leading to increased herbicide use. The tunneling and mounds that mole crickets make also disrupt the playing surface on golf courses, especially the roll of the golf ball on greens. Superintendents and golf course members typically have zero tolerance for damage (Frank and Parkman 1999). Insecticide treatments are usually targeted against the most destructive nymphal stages (Parkman and Frank 1996, 1998). Mole cricket damage and cost of control in Florida in 1986 were estimated at $45 million with an additional $33 million in Alabama, Georgia, and South Carolina combined (Frank and Parkman 1999). Estimates of annual expenditure on chemical insecticides are over $18 million in Florida turf, and over $12 million in control costs (Hudson et al. 1997). Mole crickets are omnivorous, feeding on animal as well as plant material. Studies have indicated that the southern mole cricket, Scapteriscus borellii Giglio-Tos, is less damaging than the tawny mole cricket, Scapteriscus vicinus Scudder. Scapteriscus borellii is a predator and feeds mostly on other insects while S. vicinus is mainly herbivorous (Matheny 1981, Matheny et al. 1981, Walker and Dong 1982). Dissection of 1

PAGE 16

2 field trapped S. borellii showed that their gut contents contained 66% animal material and only 2% of plant material, and the rest a combination of plant and animal material. In contrast, 84% of the gut contents of the short-winged mole cricket, Scapteriscus abbreviatus Scudder, and 88% of S. vicinus contained plant material (Hudson 1985). Both S. vicinus and S. borellii are pests of tomato and strawberry fields in Florida (Schuster and Price 1992), as well as many vegetables, peanut, sugarcane, tobacco, and ornamentals such as coleus, chrysanthemum, and gypsophila. Among turfgrasses, S. vicinus often injures bahiagrass and bermudagrass, whereas S. abbreviatus favors St. Augustinegrass and bermudagrass. Mole crickets also feed on weeds such as pigweed and Amaranthus spp. (Capinera and Leppla 2001). There are ten mole cricket species in the continental United States, Hawaii, Puerto Rico, and the Virgin Islands (Frank et al. 1998). All mole cricket species are not pests. Most species are innocuous and some are rare. For example, in Britain Gryllotalpa gryllotalpa (L.) has become so rare that efforts to reintroduce it to mainland England have been proposed (Spinney 1995). In the United States the prairie mole cricket, Gryllotalpa major Saussure, feeds on prairie vegetation and is restricted to four central states due to habitat loss, though it once had a much wider distribution (Vaughn et al. 1993, Frank et al. 1998). Mole crickets primarily live underground in excavated tunnels. The forelegs of the mole cricket are flattened and expanded, enabling it to burrow quickly in sandy and extremely dry soils. Mole crickets form vertical tunnels as well as horizontal galleries just below the soil surface. Mole crickets usually occur in the top 20-25 cm of soil and have been recorded to tunnel as deep as 75 cm (Hudson 1985). The depth of mole cricket

PAGE 17

3 tunneling varies with temperature and moisture. Galleries made by the crickets can be used as an indication of their presence in the turf. Adults and large nymphs occasionally move about on the soil surface on warm nights with high humidity and are often attracted to lights. Male mole crickets produce loud songs (50 dB) after sunset by rasping a stridulatory file on the forewing. Males use a funnel-shaped opening at the mouth of a subterranean calling chamber, which amplifies the sound of their song. Calling chambers are constructed each evening 10-20 min. before calling. The male mole cricket then tunes these chambers to the frequency of his song (Forrest 1985), and calling lasts for approximately 1 h after sunset. Songs function to attract flying and walking females. The males song intensity varies depending on male size and soil moisture (Forrest 1985). The following information on the mole cricket life cycle was determined by Walker (1985) and Frank and Parkman (1999). Adult S. vicinus fly in large numbers in early spring, typically in March, but as early as February in Florida after warm winters. In early summer, mole crickets mate (although some mating occurs the previous autumn), and oviposit one or more clutches of 25-60 eggs. Females mate with males for < 24 h, lay one or more egg clutches within 10-14 d after mating and then die. Eggs mature within 3 wk in an incubation chamber in the soil. Nymphs hatch from eggs as early as April, but continue to hatch from later deposited eggs through June. Nymphs develop for about 5 mo and adults begin to appear in September. In some years there is a minor peak of flight activity in the autumn, as early as August in the far south and as late as December farther north if the weather remains warm. In most of the southern USA, the spring activities occur in S. borellii about 3 wk later than in S. vicinus, but the autumnal

PAGE 18

4 activities are concurrent. In south Florida, S. borellii has two generations during the summer months, with a second peak in adult flight in July, following the first peak in April. More S. borellii than S. vicinus overwinter as large nymphs. In contrast, all developmental stages of S. abbreviatus occur throughout the year but with two peak ovipositional periods, one in late spring and one in winter. Scapteriscus abbreviatus has short, non-functional wings, cannot fly, and does not produce calling sounds. The peak periods of damage caused by feeding of any species are when nymphs are abundant, developing rapidly, and ingesting much food. Management Practices Chemical Control Turf damage mitigation is often the top priority of turfgrass managers and this is usually achieved by applying pesticides. Insecticide classes used to manage mole cricket populations include carbamates, organophosphates, phenyl pyrazoles and pyrethroids. Insecticides commonly applied to control mole crickets in lawns and golf courses are expensive and not always effective while in pastures there is no economically feasible control (Walker 1985). Several risks are associated with insecticide use for mole cricket control. For example, insecticides often have a short residual and treated areas are subject to reinfestation. Insecticides are usually non-specific and therefore non-target insects, including natural enemies, are killed by applications. Insecticide-treated mole crickets may die on the soil surface, which attracts birds and other insectivores and risks secondary mortality of these organisms. Improper applications may contaminate groundwater by runoff or seepage (Frank and Parkman 1999). Finally, insecticide use on golf courses and athletic fields requires that play be suspended for the legally required

PAGE 19

5 no-entry time after treatment to minimize human exposure. Scapteriscus mole crickets are therefore good targets for classical biological control (Frank 1990). Non-Chemical Control Other options for non-chemical control of mole crickets exist (e.g., cultural, physical or mechanical controls), but are not readily applicable to turfgrass and pastures of the southeastern USA. Examples include tillage at appropriate times of year to expose eggs and small nymphs to desiccation and flooding (Denisenko 1986, Sithole 1986). Reducing mowing height acts as a mechanical control by killing any mole crickets on the soil surface. Biological Control There are several natural enemies of precinctive and exotic pest mole crickets in the USA. Birds, mammals, amphibians, reptiles and arthropods attack mole crickets. They include birds, raccoons and armadillos, toads, snakes, carabid beetles and earwigs. Several fungal pathogens have been isolated from mole crickets in Florida, including Aspergillus, Beauveria, Isaria, Metarhizium, and Sorosporella (Boucias 1985, Pendland and Boucias 1987). Several arthropods are also used as biological control agents of Scapteriscus mole crickets, including the phonotactic tachinid fly Ormia depleta (Wiedemann), the neotropical digger wasp Larra bicolor F., Megacephala tiger beetles, Pasimachus carabid beetles, and wolf spiders (Hudson et al. 1988, Parkman et al. 1996). Of these, O. depleta and L. bicolor have been released in Florida. In 1988, a Brazilian strain of O. depleta was released in Florida to parasitize exotic mole crickets. The female fly locates Scapteriscus spp. by mating songs and deposits larvae on or near the host. The flys seasonality limits its spread so it has been more successful in central and south Florida. Golf course superintendents have reported that

PAGE 20

6 counties in Florida with O. depleta populations had significantly less mole cricket damage than did counties that lacked O. depleta (Frank et al. 1996). Ormia depleta seems to overwinter more successfully in central Florida than in northern Florida, perhaps because of milder winters in central locations (Walker et al. 1996). The neotropical digger wasp L. bicolor parasitizes large nymphs and adults of mole crickets (Frank and Parkman 1999). The wasp attacks and stings mole crickets on the soil surface, causing paralysis. The wasp deposits an egg on the mole cricket near the pronotum, and the neonate larva develops as an external parasitoid. The parasitized mole cricket resumes normal activities and dies after a few weeks. Scapteriscus mole crickets are the only known host of L. bicolor (Frank et al. 1995). However, this wasp also feeds on wildflower nectar which can be used to maintain or increase populations in an area. Insect Parasitic Nematodes Insect parasitic nematodes have recently been investigated for use against subterranean and soil inhabiting pests (Kaya and Gaugler 1993). Insect parasitic nematodes in the genera Steinernema and Heterorhabditis are potent biological control agents that generally infect their hosts by entering natural openings such as the mouth, spiracles, and anus (Shapiro and Lewis 1999). Insect parasitic nematodes have been used to control mole crickets and other insect pests such as lepidopteran larvae (Epsky and Capinera 1994, Shapiro and Lewis 1999), banded cucumber beetle (Creighton and Fassuliotis 1985), sweet potato weevil (Jansson et al. 1993), pecan weevil larvae (Shipiro-Ilan 2001), western cherry fruit fly (Patterson-Stark and Lacey 1999), Japanese beetle grubs (Klein and Georgis 1992, Schroeder et al. 1993), and various Homopterans (English-Loeb et al. 1999). The nematodes in these genera are mutualistically associated with bacteria (Xenorhabdus spp. for steinernematids and Photorhabdus spp. for

PAGE 21

7 heterorhabditids). Infective juvenile nematodes enter the host through an opening in the arthropod. Once in the hemocoel, they release their symbiotic bacteria, which kill the host and provide nematodes with nutrients and defense against secondary invaders (Poinar 1990). The nematodes complete two to three generations within the arthropod host, and then infective juveniles are released to search out new hosts (Kaya and Gaugler 1993). Several steinermatid and heterorhabditid nematodes are in current use, or are being considered for use, as commercial biopesticides against soil insects in many agricultural and horticultural systems (Gaugler and Kaya 1990, Kaya 1990, Georgis and Gaugler 1991, Kaya and Gaugler 1993). Disadvantages in using insect parasitic nematodes include sensitivity to ultraviolet light, desiccation, and insect/pathogen interactions (Kaya and Gaugler 1993, English-Loeb et al. 1999). Certain nematode species are highly effective against a particular pest, whereas others may be ineffective or moderately effective against the same pest (Shapiro-Ilan 2001). The effectiveness of entomopathogenic nematodes depends on matching the target pest species with the most effective nematode. The entomopathogenic nematode Steinernema scapterisci Nguyen and Smart was collected in Uruguay in pitfall-trapped Scapteriscus mole crickets in the 1980s. The nematode was cultured and then released in Florida pastures in 1985, established populations and was spread from the release site by infected Scapteriscus mole crickets (Hudson et al. 1988, Nguyen and Smart 1990a, Parkman and Frank 1992, Frank 2001). The nematode successfully kills adults and large nymphs of S. borellii and S. vicinus and to a lesser extent S. abbreviatus. Small to medium nymphs of S. borellii and S. vicinus are less frequently infected (Hudson and Nguyen 1989a).

PAGE 22

8 Steinernema scapterisci live in moist soil and can survive without a host for at least 10 wk (Nguyen and Smart 1990a). Third stage infective juveniles can migrate up and down 10 cm in the soil and invade mole cricket bodies through the mouth or spiracles (Nguyen and Smart 1991). Once inside the mole cricket, the nematodes release bacteria that feed on the hemolymph killing the mole cricket through septicemia; the nematodes then eat the bacteria and reproduce inside the mole cricket resulting in mole cricket mortality within 2-3 d. The nematodes then exit the body and infect other mole crickets in the soil. Mole crickets can fly up to 8 km before dying (Walker 1985), thus spreading nematodes to new sites. The use of S. scapterisci as a biopesticide would allow treatment to hot spots rather than doing a broadcast application of synthetic insecticides. Anecdotal evidence indicates that since the original release of the nematodes (ca. 1985), overall mole cricket populations in Florida have declined (J.H. Frank, personal communication). Augmentative nematode releases are likely to further reduce mole cricket populations. Over the past 20 years, researchers have become very interested in changes in behavior displayed by parasitized animals and whether or not they represent parasite or host adaptations (Poulin 1995). Changes in behavior displayed by parasitized organisms vary from slight shifts in the time spent performing a given activity to appearance of drastically new and strange behaviors (Poulin 1994, Benton and Pritchard 1990). Oftentimes these parasite induced behavioral modifications are simply pathological side effects of parasite infection (Poulin 1995, Vance 1996, Williams et al. 2001). Studies with the thrips Frankliniella occidentalis (Pergande) and the entomopathogenic nematode Thripinema nicklewoodi Siddiqi demonstrated behavioral change in the host insect. The behavioral changes displayed were reduced feeding and reduced fecundity of F.

PAGE 23

9 occidentalis when infected with T. nicklewoodi (Arthurs and Heinz 2003). The results from this study proved useful in the biological control of thrips because T. nicklewoodi reduce populations by parasitizing F. occidentalis, reducing direct feeding damage, and reducing the spread of tomato spotted wilt virus. Nematodes also modify their hosts behavior to increase their own fitness. Parasitism of the beach hopper Talorchestia quoyana Milne-Edwards by mermithid nematodes results in a greater burrowing depth of the host. Adult mermithids live in water or moist soil; however their beach hopper host prefers dry terrestrial microhabitats. This behavior modification of the host allows the nematode to mate and lay eggs in more humid environments deeper in the soil (Poulin and Latham 2002). Maeyama et al. (1994) observed that the ant Colobopsis sp. infected with Mermis nematodes displayed a suicidal behavior by jumping into water and dying. This behavior is advantageous to the nematode because they require water to reproduce. In order to increase contact with a mate, the nematodes must emerge from the ants in water. More than 30 species of nematodes are associated with insects and other invertebrates (Poinar 1979, 1990; Kaya and Stock 1997; Lacey et al. 2001; Koppenhfer and Fuzy 2003). The nematode families Allantonematidae, Heterorhabditidae, Mermithidae, Tetradonematidae, Sphaerulariidae, and Steinernematidae are the focus of much research because of their potential as biological control agents of insects (Lacey et al. 2001). For example, inoculative releases of Deladenus siricidicola Bedding in New Zealand and Australia has been a successful classical biological control agent of the woodwasp Sirex noctilio F. (Bedding 1993). The mermithid Romanomermis culicivorax

PAGE 24

10 Ross and Smith has been used as a successful, inundative biological control agent for mosquito larval suppression (Petersen 1985). Steinernematid and heterorhabditid nematodes are second only to Bacillus thuringiensis Berliner in commercial sales at $2-3 millon dollars annually (Georgis 1997). These nematodes infect a number of insect species yet pose no threat to plants, vertebrates, and many invertebrates (Akhurst 1990, Kaya and Gaugler 1993). They can be mass produced, formulated, and easily applied as biopesticides; they also have been exempt from registration in many countries, are compatible with many pesticides, and are amenable to genetic selection (Georgis and Kaya 1988, Kaya and Gaugler 1993, Georgis and Manweiler 1994). Nematodes in the families Steinernematidae and Heterorhabditidae are especially efficacious against insects in soil and cryptic habitats (Lacey et al. 2001). These nematodes have been used inundatively in many high value crop systems (Georgis and Manweiler 1994, Koppenhfer 2000). Successful uses of these nematodes against economically important pests include the citrus root weevil, Diaprepes abbreviatus (L.), in citrus, the black vine weevil, Otiorhynchus sulcatus (F.), in nurseries and cranberries, the peach borer moth, Carposina niponensis Walsingham, in apples, and the black cutworm, Agrotis ipsilon (Hufnagel), in turfgrass (Lacey et al. 2001). Objectives Greater understanding of alternative controls, such as using insect parasitic nematodes and how they fit into an integrated pest management program for mole crickets in turfgrass, is needed. This research specifically assessed the effectiveness of Steinernema scapterisci through a series of laboratory and field experiments. The research included evaluating 1) the establishment of S. scapterisci in soil growing high

PAGE 25

11 value golf course turfgrass, 2) effective drenching solutions in order to sample areas to determine levels of nematode infected mole crickets, 3) compatibility of S. scapterisci with insecticides, and 4) the effect of S. scapterisci infection on the reproductive and tunneling behavior of pest mole crickets.

PAGE 26

CHAPTER 2 ESTABLISHMENT AND SPREAD OF Steinernema scapterisci ON FLORIDA GOLF COURSES An integrated pest management program for pest mole crickets (Scapteriscus abbreviatus, S. borellii, and S. vicinus) is being developed throughout the southeastern United States. These insects damage turfgrass by tunneling and root feeding, that results in large, irregular patches of dead turf throughout the year. Chemical control on golf courses is still the primary means of preventive and curative control. Using resistant or tolerant varieties or species of turfgrass, such as the bermudagrass hybrids TifSport and Ormond, is possible but not common (Hudson 1986, Hanna and Hudson 1997, Braman et al. 2000, Hanna et al. 2001, Reinert and Busey 2001). Cultural controls (i.e., adjusting irrigation, fertilization, mowing heights, etc.) have not affected mole cricket populations (Denisenko 1986, Frank et al. 1998, Frank and Parkman 1999. However, the introduction and conservation of natural enemies that attack mole cricket adults is gaining momentum, especially on pastures and Audubon International affiliated golf courses The purpose of this study was to establish the efficacy of augmentative releases of S. scapterisci against Scapteriscus spp. mole crickets on highly maintained golf courses and athletic fields (Appendix B) as well as determining if subsequent applications of nematodes would augment the nematode populations in the test area and increase the percentage of mole crickets infected. 12

PAGE 27

13 Materials and Methods Study Sites The establishment and spread of S. scapterisci was monitored on two golf courses in Alachua Co., FL: Ironwood Golf Course and Gainesville Golf and Country Club. Ironwood Golf Course (IGC) was an 18-hole city-owned public golf course built in 1964. The roughs were bermudagrass (Cynodon dactylon Pers. C. transvaalensis Burtt-Davy) var. Tifway, mowed at 4.7 cm. Gainesville Golf and Country Club (GGCC) was an 18-hole private course located 10.3 miles from Ironwood Golf Course. Gainesville Golf and Country Club was built in 1962 and originally planted with bermudagrass var. Ormond and the roughs were mowed at 3.2 cm. Ironwood Golf Course and GGCC had been previously treated with S. scapterisci in the late-1980s and did not have any subsequent treatments. Weather data, including minimum and maximum daily temperatures, relative humidity, amount of monthly precipitation (from local weather stations located approximately 1 mile from IGC and 9 miles from GGCC) and soil temperatures at 7.6 cm below the soil surface, were documented on each collection date (Appendix A). Mole Cricket Monitoring Twenty hot spots of mole cricket activity were located in the roughs of ten fairways on each golf course (two hot spots per hole). Linear pitfall traps (modified from Lawrence 1982) were installed in the ground at least 80 m apart. Each golf course fairway contained a trap located in one treated area and one untreated area. Pitfall traps consisted of a 19 L plastic bucket placed in the ground and four PVC (3 m long, 7.6 cm diameter) perpendicular arms with a 2.5 cm wide slit lengthwise along the top. The arms were placed in the ground so the slits were flush with the soil surface (Figure 2-1). The distal end of each arm was capped, so insects falling into an arm

PAGE 28

14 eventually fell into a 3.8 L bucket containing approximately 3-5 cm of sand, located within a 19 L bucket (Figure 2-2). Holes were drilled into the bottom of both buckets to allow for water drainage. Traps on all sites were installed in September and October 2001. Nematodes were released in the afternoon (approximately 1600) at Ironwood Golf Course (31 October 2001) and in the morning (approximately 0700) at the Gainesville Golf and Country Club (5 November 2001). Nematodes were applied in an aqueous suspension of 1 billion nematodes/ 378.5 L of water applied using a boom sprayer calibrated at 0.5 L/m 2 The area treated was 20.1 20.1 m (0.04 ha) around one trap on each fairway for both golf courses. All sites were irrigated with 0.6 cm of water before and 0.6 cm after application. The pre-treatment dates for Gainesville Golf and Country Club and Ironwood Golf Course were 11, 18, 25 October 2001. Pitfall traps were used to monitor infection levels and mole cricket abundance using methods similar to Parkman et al. (1993a, b). At each 24 h sampling period, the buckets and arms were cleaned out and 3-5 cm of sand was placed into the inner bucket. Traps were left for 24 h and adult and juvenile mole crickets with pronotal lengths > 4 mm (Hudson and Nguyen 1989a) were collected from traps and returned to the laboratory. Crickets were tested for infection weekly for the initial 6 wk post-application and one to two times a month thereafter for 1 yr on Gainesville Golf and Country Club and 2 yr on Ironwood Golf Course. Turf quality (density, color) in the area immediately surrounding the pitfall traps was assessed (1 sparse or brown grass, 9 dense or dark green grass). Soil samples were taken from each golf course and soil texture was

PAGE 29

15 analyzed using the sodium metaphosphate/hydrometer procedure. Traps were removed from GGCC in October 2002 at the superintendents request. Laboratory Assay Percent of mole crickets infected with nematodes caught in the traps was monitored and tested in the laboratory. Mole crickets were placed individually in 20 ml plastic scintillation vials (Fisher Scientific) with 1-2 drops of deionized water, capped and labeled. Mole crickets were examined at 7 and 10 d after death under a dissecting scope (10X) for the presence of nematodes. Steinernema scapterisci were identified by Dr. Khuong Nguyen, Entomology and Nematology Department, University of Florida. Statistical Analysis Comparisons of infection rates between sites and years were subjected to analysis of variance and Tukeys studentized range test or Students t-test (SAS Institute 2001). All comparisons were made at a 0.05 significance level. Non-transformed means plus or minus one standard error of the monthly mean are presented. Results and Discussion Mole crickets infected with S. scapterisci were collected at both golf courses before and after our augmentative applications. Infected mole crickets were only found in the spring and fall of each year when the late instar and adults, the most susceptible life stages to S. scapterisci infection, were present. Mean cumulative percentages ( SE) of infection for mole cricket trap collections during the 2001-2003 field season from the sites GGCC (22.1% 10.5) and IGC (15.8% 4.6) did not differ statistically (t = 2.00 ; df = 2, 56; P > 0.0001). Monthly infection and baseline pretreatment infection levels for IGC and GGCC are presented in Figures 2-3 and 2-4, respectively.

PAGE 30

16 Pre-treatment infection rates at GGCC and IGC ranged from 10-15%. The percentage of infected mole crickets in treated areas at GGCC exceeded pre-application levels about 4-8 wk after application. The percentage of infected mole crickets in untreated areas at both courses (>80 m from treated areas) equaled the percent infection in treated areas after about 20 wk. Significantly fewer Scapteriscus spp. were collected in year 2 than in year 1 at IGC (t = 2.47; df = 1, 37; P > 0.01). The S. scapterisci population persisted throughout the entire study period, 1 yr for GGCC and 2 yr for IGC; however at IGC the level of infection for year 2 was significantly less than year 1 (F = 6.63; df = 1, 37; P > 0.01). During this study fipronil and acephate were used on greens of both courses; acephate was used in the roughs as a spot treatment on IGC. Data from this study concur with Parkman et al. (1994) that more Scapteriscus adults were infected with S. scapterisci than nymphs. Contrary to Parkman et al. (1994), overall infection of S. borellii was lower than that of S. vicinus in this study. More S. vicinus were collected in traps than S. borellii (Table 2-1). Prior applications of S. scapterisci were made in 1988 and 1989 on Ironwood Golf Course and Gainesville Golf and Country Club, respectively. Infection levels present in mole crickets during the pre-treatment collections demonstrate that the nematodes can survive in golf course soil where mole crickets are present and pesticides are used regularly for 12 years. If the nematodes can persist for at least 12 years, then the populations will probably last for many more years as effective biological control agents. Steinernema scapterisci can survive for 10 wk in soil lacking mole crickets (Nguyen and Smart 1990a), however if crickets are present the nematode recycles within the host in soil environments and could survive for 12 years as demonstrated by our results. The

PAGE 31

17 persistence of nematodes in the soil may reduce the populations of mole crickets leading to an overall reduction of pesticide applications needed. Establishment and persistence of S. scapterisci in pastures was evaluated by Parkman et al. (1993a) for five years. Nematodes were applied as an aqueous solution using a handheld watering can or by burying infected mole cricket cadavers at the test sites. Crickets were collected using linear pitfall traps (Lawrence 1982) with electronic callers located nearby to enhance local mole cricket populations. Based on their findings S. scapterisci became established and persisted for five years at these pasture field sites. Parkman et al. (1993b) also evaluated the efficacy of a single inoculative release of S. scapterisci against mole crickets in pastures. Nematodes were applied in an aqueous solution using a tractor-drawn chisel rig. Crickets were collected in pitfall traps placed at 50, 100, and 200 m from the center of the treated area. The nematode persisted at five of the six sites and dispersed at least 150 m away from the initial treated area at three of the sites (Parkman et al. 1993b). They found that inoculative releases of S. scapterisci were an alternative to inundative releases for mole cricket suppression. Further tests (Parkman et al. 1994) showed that S. scapterisci serves as an inoculative biological control agent for Scapteriscus spp. mole crickets on golf courses and acted as a biopesticide for relatively rapid suppression of pest populations (Parkman et al. 1993b). Establishment on golf courses was not as successful as pastures; however there was a >27% pest population reduction in areas where the nematode did persist (Parkman and Smart 1996). Since its initial release in June 1985 in Alachua Co., Florida, S. scapterisci has been non-commercially released and presumably established in at least 28 counties in Florida. The nematode population established readily even after a single inoculative

PAGE 32

18 release. The establishment of S. scapterisci has been demonstrated by strip and broadcast spray applications. Mole crickets infected with S. scapterisci can fly several kilometers before dying (Parkman and Frank 1992), thus potentially spreading nematodes to uninfected sites. Therefore the use of S. scapterisci as a biopesticide would allow treatment to hot spots rather than strict reliance on broadcast applications of synthetic insecticides to provide long-term mole cricket suppression. However, the effect of augmentative applications to hot spots of mole cricket activity has not been previously determined. From our results it is evident that a single augmentative application of S. scapterisci is sufficient to enhance nematode populations throughout a local area (i.e., a fairway). Augmentative applications of nematodes provided higher than baseline infection levels for 17 mo post treatment at IGC and 8 mo post treatment at GGCC. Infection levels fluctuated with the host population in our study. Mole cricket adults and large nymphs are present in the spring and fall months of any given year therefore, crickets infected with S. scapterisci were usually found in larger numbers during these months. The reduction in mole cricket population levels in year 2 versus year 1 could be attributed in part to the S. scapterisci applications suggesting that the augmentation of nematode populations does help reduce the numbers of mole crickets present on highly managed golf courses in north Florida. Levels of infection may also be influenced by mole cricket age, activity, environmental conditions and predation by other arthropods within a trap. Although a single application can establish nematode populations, an augmentative application may be necessary to keep population levels high enough for sufficient mole cricket control.

PAGE 33

19 Mole crickets collected from the pitfall traps were subject to mortality from organisms other than S. scapterisci. Organisms observed infecting mole crickets included other species of insect parasitic nematodes (i.e., Heterorhabditis spp. and Steinernema spp.), fungi (i.e., Beauvaria bassiana), predators (i.e., ground beetles, spiders, earwigs, etc.), and other mole cricket parasitoids (Larra bicolor and Ormia depleta). Previous research (Gaugler and Boush 1978, Molyneux 1985, Hudson and Nguyen 1989b, Smith 1999) has shown that there are several abiotic factors that contribute to the success (or failure) of insect parasitic nematodes when used as a biopesticide. Abiotic factors can significantly limit the nematodes effectiveness to move, locate and enter a host (Smith 1999). Some of these abiotic factors include ultraviolet light, desiccation, soil moisture, soil texture and type, soil temperature, soil pH, and agrichemical compatibility (see Chapter 4). In this study applications were done in the early morning and late afternoon and watered in immediately after application to avoid nematode damage from UV light and desiccation. Moisture is required for nematodes to move through the soil (Hudson and Nguyen 1989b). Ames (1990) observed that infective juveniles of S. scapterisci can survive up to 13 wk at wilting point (15 bars moisture tension) and survive better in sandy loam than pure sand. In this study both courses had sandy loam soil and were irrigated regularly. Several of the fairways and roughs on both courses where crickets were collected were subject to flooding or extremely wet conditions due to rain or excess irrigation. Molyneux and Bedding (1984) observed that very saturated soils can inhibit nematode mobility and decrease their survival by creating anaerobic conditions.

PAGE 34

20 Steinernema scapterisci killed between 15-20% (on average) of mole crickets collected in linear pitfall traps. However, because mole crickets may die as soon as 48 h post-infection, this value may be an underestimate of the true percentage of kill. Nematodes can kill generation after generation and this combined effect may be something similar to compound interest (Frank et al. 2002). Steinernema scapterisci can be an effective part of an integrated pest management system on managed turfgrass if applied in the proper manner and in suitable locations where nematodes can survive.

PAGE 35

21 Figure 2-1. Linear pitfall trap used to collect mole crickets. Figure 2-2. 3.8 L catch bucket inside 19 L bucket of linear pitfall trap.

PAGE 36

22 0102030405060OCT 01NOV 01DEC 01JAN 02FEB 02MAR 02APR 02MAY 02AUG 02OCT 02MAR 03 MonthInfection Level (% ) Untreated Treated 5 4 27 7 19 34 18 4 91 3 13 4 78 9 4 73 10 56 78 7 2 Figure 2-3. Mean monthly ( SEM) percent infection of mole crickets collected in pitfall traps at Ironwood Golf Course from areas treated with Steinernema scapterisci. Only months with infection levels are presented. Untreated areas received no S. scapterisci and were >80 m from treated areas. Dashed line represents baseline pretreatment infection level. Data presented are for Scapteriscus vicinus and Scapteriscus borellii combined. Total numbers of mole crickets are presented above SEM bars.

PAGE 37

23 020406080100120OCT 01NOV 01DEC 01MAR 02APR 02MAY 02MonthInfection Level (% ) Untreated Treated 5 7 3 14 16 13 61 42 59 9 6 Figure 2-4. Mean monthly ( SEM) percent infection of mole crickets collected in pitfall traps at Gainesville Golf and Country Club from areas treated with Steinernema scapterisci. Only months with infection levels < 0 are presented. Untreated areas received no S. scapterisci and were >80 m from treated areas. Dashed line represents baseline infection level. Data presented are for Scapteriscus vicinus and Scapteriscus borellii combined. Total numbers of mole crickets are presented above SEM bars.

PAGE 38

24 Table 2-1. Percent infection (mean SEM) of Scapteriscus spp. mole crickets collected from sites treated with Steinernema scapterisci. Pairs of means within columns followed by asterisks are significantly different, t-test (P > 0.05). Numbers in parentheses are the total mole crickets collected. Gainesville Golf and Country Club (n=10 traps) 1 Ironwood Golf Course (n=10 traps) 2 S. borellii Nymphs 0 (1) 0 (15) Adults 0 (1) 3.3 2.6 (20) Total 0 (2)* 1.6 1.3 (35)* S. vicinus Nymphs 7.7 4.0 (70) 9.7 3.4 (208)* Adults 13.9 4.5 (163) 33.2 7.0 (313) Total 10.8 3.0 (233) 11.9 2.5 (521) 1 GGCC: F= 8.57; df= 2,111; P= 0.0003 2 IGC: F= 14.05; df= 2,155; P< 0.0001

PAGE 39

CHAPTER 3 SURVIVAL AND INFECTIVITY OF Steinernema scapterisci AFTER CONTACT WITH SOIL DRENCH SOLUTIONS Mole crickets are subterranean pests of turfgrass in Florida and much of the southeastern United States (Walker and Nickle 1981, Walker 1985). Mole cricket damage and cost of control in Florida in 1986 were estimated at $45 million with an additional $33 million in Alabama, Georgia, and South Carolina combined (Frank and Parkman 1999). Estimates of annual expenditure in 1996 were over $18 million for insecticides in Florida turf, and over $12 million in control costs (Hudson et al. 1997). Mole crickets damage turf by their tunneling in the soil, which exposes and dries out roots and by direct root feeding. As a result, the turfgrass thins and bare patches appear. The tunneling and mounds that mole crickets make also disrupt the playing surface on golf courses, especially the roll of the golf ball on greens. Superintendents and golf course members have little tolerance for damage (Frank and Parkman 1999). Insecticides are usually targeted against the most destructive, nymphal stage. A more sustainable, environmentally friendly management approach for mole cricket control is needed. Several biological control agents have been investigated for control of Scapteriscus spp. mole crickets in Florida (Hudson et al. 1988). One of these biological control agents is an entomopathogenic nematode, Steinernema scapterisci. Steinernema scapterisci was originally isolated from pitfall-trapped Scapteriscus vicinus in Uruguay in the 1980s (Nguyen and Smart 1990b). The nematode was cultured and released in several Florida counties in 1985, where it established a population, and was spread from the release site 25

PAGE 40

26 by infected Scapteriscus mole crickets (Hudson et al. 1988, Parkman and Frank 1992). The nematode kills the adult and late instar nymphs of Scapteriscus borellii and S. vicinus, and to a lesser extent S. abbreviatus. Fewer small to medium-sized nymphs of S. borellii and S. vicinus become infected (Nguyen 1988). Several techniques have been used to sample mole crickets including counts of dead nymphs and adults after insecticide applications (Short and Koehler 1979), estimation of surface burrowing (Walker et al. 1982, Cobb and Mack 1989), pitfall trapping (Lawrence 1982, Adjei et al. 2003), removal with a tractor mounted soil corer (Williams and Shaw 1982), sound trapping (Walker 1985) and soil drenching (Short and Koehler 1979, Walker 1979, Hudson 1989). However, results from each of these techniques are often inconsistent (Short and Koehler 1979, Lawrence 1982, Hudson 1988). Comparisons of different methods have indicated that soil drenching with soap solutions are the most practical and consistent at obtaining direct counts of mole crickets (Short and Koehler 1979, Hudson 1988). Soil drenching with a solution of 15 ml of lemon dishwashing detergent in 3.8 L of water is inexpensive and commonly used by turfgrass managers to sample soil pests. Soil drenches with soap solutions irritate mole crickets and force them out of the soil. Soap flushes are often used for monitoring mole crickets to determine the size, age, and species present, the relative population density over time, and for control timing. However, it was suspected that soap flushes, when used to monitor mole crickets potentially infected with S. scapterisci, might be lethal to the nematodes because we rarely found nematodes in soap-flushed mole crickets (K.B. Nguyen and G.C. Smart, Entomology and Nematology Dept., University of Florida, pers. comm.). Solutions such as pyrethroids,

PAGE 41

27 ammonia, vinegar, Lysol, and other soap detergents have previously been tested as potential soil drench solutions (Short and Koehler 1979). This study was conducted to determine whether a standard soap detergent solution affects S. scapterisci survival and infectivity in pest mole crickets. Potential alternatives to the standard soap drench solution were also evaluated. Materials and Methods Nematodes and Mole Crickets Steinernema scapterisci (Nematac S Becker Underwood, Ames, IA) was stored in a 7C cold room until use (<3 mo). Nematode viability was tested before each application by dissolving a pinch (~10 mg) of Nematac S into water and observing nematode shape and mobility under a light microscope. Healthy nematodes were opaque in color and S-shaped with undulating movements. Dead or unhealthy nematodes were translucent, straight, and lacked movement. The product was used if viability was >50% and discarded if <50% viable. Scapteriscus vicinus were collected from pitfall traps or sound traps in Alachua Co., FL, and returned to the laboratory. Each mole cricket was placed in a 120-ml plastic vial (Thornton Plastics Salt Lake City, UT) with sterilized sand and held for > 14 d to ensure health. Surviving mole crickets were used in this study. Mole crickets were maintained at 23C with a photoperiod of 12:12 (L:D) and fed commercial cricket chow (Purina St. Louis, MO). Bioassay Nematode viability and infectivity were assessed after exposure to various drenching materials. Steinernema scapterisci nematodes were extracted from Nematac

PAGE 42

28 S using a modified Baermann technique (Nguyen and Smart 1990b). Steinernema scapterisci nematodes were kept at a density of 10,000 infective juveniles in solutions of water (control), lemon dishwashing detergent (Joy Proctor and Gamble, Cincinnati, OH), insecticidal soap (Safer Soap Woodstream Corporation, Litiz, PA), and permethrin (Spectracide Bug Stop Spectrum Brands, St. Louis, MO) for test 1. The mixtures were kept at room temperature in a 125-ml Erlenmeyer flask with 125-ml per flask on a shaker at 65 rpm. There were five replicates for each treatment. Concentrations were selected based on recommendations for flush extraction of mole crickets in the field (Short and Koehler 1979) and label rates for mole cricket control. After 24 h, 10-l samples were taken from each treatment and placed on a microscope slide. The numbers of living and dead nematodes were counted using a dissecting microscope (10 x), three 10-l counts were taken and averaged to determine percent mortality for each replicate. Immobile nematodes were touched with a probe to determine survival. A second test was initiated to further test potential drench materials. Treatments for test 2 included water (control), azadirachtin (Safer Brand BioNeem, Woodstream Corporation, Litiz, PA), citrus oil (Green Sense Garland, TX), garlic extract (Garlic Barrier Garlic Research Labs, Inc., Glendale, CA), lemon juice (ReaLemon Motts, St. Louis, MO), permethrin (Spectracide Bug Stop Spectrum Brands, St. Louis, MO) and cyfluthrin (Bayer Advanced Lawn and Garden Complete Insect Killer, Bayer Environmental Sciences, Montvale, NJ). Concentrations were selected based on label and half label rates for mole cricket control. Methods from test 1 were repeated.

PAGE 43

29 Nematode infectivity was assessed by filtering nematodes from above solutions and adding 50 living infective juveniles (parasitic stage) to 120 ml plastic cups (Fisher Scientific) containing 20 g sterilized sand, 4% deionized water, and one S. vicinus adult. Dead mole crickets were dissected and the presence or absence of nematodes was recorded. The above solutions were tested for their effectiveness at flushing mole crickets at the University of Florida G.C. Horn Turfgrass Research Unit in Gainesville, FL, on 20 and 28 May 2003. Each treatment from tests 1 and 2 (3.8 L of each solution) was applied to areas of bermudagrass (Cynodon dactylon Pers. C. transvaalensis Burtt-Davy) var. Tifway, that had mole cricket damage (75 75 cm 2 ). The numbers of adult and first instar mole crickets emerging from the soil within 3 min were counted. Five replicates for each solution were completed. Any turfgrass phytotoxicity was noted 1 h post application and 1 wk post application. The effect of nematode infected crickets exposed to soap solutions was also tested. Scapteriscus abbreviatus adults were obtained from a lab colony at the University of Florida Entomology and Nematology Department, Gainesville, FL, and were inoculated with about 10,000 nematodes by applying a predetermined amount (approximately 150 l) of concentrated nematode solution onto a piece of filter paper (Fisher #P8, 5.5 cm) inside a petri dish with one S. abbreviatus adult. The mole cricket was allowed to incubate in the petri dish for 1, 5, 8, 12 or 24 h (five mole crickets per treatment). Scapteriscus abbreviatus was used because S. vicinus adults were unavailable at the time of the test. All infected mole crickets were then dipped into a 118 ml Solo souffl cup (Gainesville Paper Co., Gainesville, FL) containing the soapy water or soapy water

PAGE 44

30 followed by a deionized water rinse for 5 sec. Untreated controls were healthy, uninfected mole crickets dipped in water. Mole crickets were placed into 20-dram plastic scintillation vials (Fisher Scientific, Pittsburgh, PA) and observed every 24 h for 10 d. On day 10, mole crickets were dissected and the presence of nematodes was noted. Statistical Analysis Nematode mortality and field test data were subjected to an analysis of variance (SAS Institute 2001). Treatments were compared to the control (water) using Dunnetts means comparison method ( = 0.05). Nematode infectivity data were subjected to Chi-square analysis (SAS Institute 2001). Treatments were compared to the control (water) and the standard soap flush solution (4 ml lemon dish detergent/L water) using Dunnetts means comparison method (alpha= 0.05). Nematode mortality data were transformed using arcsine-square root transformation before statistical analysis; nontransformed data are presented. Effects of nematode infected crickets exposed to soap solutions data were subjected to ANOVA (SAS Institute 2001). Results and Discussion Permethrin at the label rate for mole cricket control caused significantly more nematode mortality than resulted from water (Table 3-1). Nematodes exposed to all treatments showed similar infectivity in mole crickets. Nematode mortality was similar among all treatments in test 2 except citrus oil (Table 3-2). Nematodes surviving all treatments, except azadirachtin and lemon juice, demonstrated a low percentage infectivity of mole crickets, no significant treatment differences were. In the field, insecticidal soap and the higher rate of permethrin flushed significantly more mole crickets than water (Table 3-3). However, when all treatments were compared to the standard lemon dish detergent, insecticidal soap and permethrin brought

PAGE 45

31 a similar number of mole crickets to the surface (n = 55; F = 2.88; df = 10,54; P = 0.008). None of the mixtures tested produced any noticeable phytotoxicity to the turf. Soil drenches with a mixture of lemon dish detergent and water are commonly used to monitor turfgrass insects such as mole crickets, chinch bugs (Blissus spp.), big-eyed bugs (Geocoris spp.), and several species of caterpillars (Short and Koehler 1979, Hudson 1989). Soil drenches are inexpensive and are not labor intensive when compared with other methods of monitoring mole cricket populations. These other methods include large linear pitfall traps (Lawrence 1982, Adjei et al. 2003), an emitter producing a synthetic song of male mole crickets (Parkman and Frank 1993), and a soil-coring device (Williams and Shaw 1982). Each method requires more than one person, and is labor intensive or costly (Lawrence 1982, Williams and Shaw 1982). Seventy percent of S. scapterisci died in the lemon dish detergent solution. Assays for nematode infection of soap-flushed mole crickets, the method currently used by many turfgrass managers, are likely to be inaccurate. Krishnayya and Grewal (2002) reported a toxic effect of a common soap surfactant (Ajax ) on S. feltiae Bovien nematodes. They found a 24% mortality level of nematodes when incubated at 4, 24, 72, and 120 h (Krishnayya and Grewal 2002). Kaya et al. (1995) reported an insecticidal soap (M-Pede ) adversely affected S. carpocapsae (Weiser) and Heterorhabditis bacteriophora Poinar survival and infectivity. However, infectivity may not be affected if the nematodes are combined with an insecticidal soap and applied immediately (Kaya et al. 1995). Nematodes cannot be stored in an insecticidal soap solution because without aeration, nematode survival can be adversely affected (Kaya et al. 1995). The toxicity of

PAGE 46

32 metal ions present in soap may be responsible for the high mortality in soap solutions (Jaworska et al. 1994, Krishnayya and Grewal 2002). Tests of exposure of nematode infected mole crickets to soap solutions show that soap flush solutions do not greatly affect nematode infection at least 8 h post infection (Table 3-4). The soap flush solutions may potentially kill nematodes in certain areas of the body (i.e., mouth) and further testing should be done to determine this. Immediately rinsing flushed mole crickets with clean water may potentially increase the accuracy of determining nematode infection. The unavailability of S. vicinus at the time of experimentation may have also led to inconsistent, low levels of infection. It is known that S. scapterisci does not infect S. abbreviatus as successfully as S. vicinus or S. borellii (Nguyen 1988). Although permethrin solutions killed some nematodes in our experiments, S. scapterisci infectivity was not compromised and field flushes successfully extracted mole crickets from the soil. Short and Koehler (1979) reported that pyrethrins were the most effective material, flushing a mean of 19 mole crickets/m 2 Hudson (1988) compared three sampling techniques, soil flushing with lemon dish detergent or synergized pyrethrins and a tractor mounted soil corer. None of the methods gave significantly different results. Our results from the field test show drenching solutions of permethrin are useful in determining whether mole crickets collected in the field are infected with S. scapterisci nematodes. A soil drench containing permethrin may be the best monitoring tool to flush mole crickets to determine the presence of S. scapterisci. However, there are disadvantages to using pyrethroids as soil drenches for mole crickets. Pyrethroid drenches at the half or full label rate may cause more mole cricket

PAGE 47

33 mortality than using a soap solution. Subsurface mortality of mole crickets can be as high as 65% when using pyrethroids or similar insecticides (Ulagaraji 1974, 1975; Walker 1979; Hudson 1988). Applicator exposure to insecticides is increased when using a pyrethroid soil drench. Soil drenches are effective, non labor-intensive methods to sample soil insect populations. Soap detergent solutions, although inexpensive, may not accurately indicate mole crickets infected with S. scapterisci. Permethrin solutions are less cost effective, but are effective at flushing mole crickets potentially infected with nematodes.

PAGE 48

34 Table 3-1. Mean nematode mortality and percent of mole crickets infected with Steinernema scapterisci after exposure for 24 h to various drenching solutions. Treatment Rate Mean % nematode mortality ( SEM) 1 Number of mole crickets infected with S. scapterisci 2 (n=3) Water n/a 2.0 1.4 2 Lemon Joy 15 ml/ 3.79 L 32.4 1.6 0 Insecticidal Soap 15 ml/ 3.79 L 40.0 5.5 1 Permethrin 18 ml/ 3.79 L 11.6 1.0 2 *Statistically significant values using Dunnetts method comparing treatments to water 1 n = 20; F = 34.58; df = 19, 3; P = <0.0001 2 More than 10 replicates are needed for statistical analysis. Table 3-2. Mean nematode mortality and infectivity after exposure for 24 h to various drenching solutions. 1 n = 45; F = 3.80; df = 44, 8; P = 0.0025 Treatment Rate Mean % nematode mortality ( SEM) 1 Number of mole crickets infected with S. scapterisci 2 (n=3) Water n/a 3.6 2.2 1 Citrus Oil 15 ml/ 3.79 L 10.6 3.2 1 Cyfluthrin 7.5 ml/ 3.79 L 2.2 2.2 1 Cyfluthrin 15 ml/ 3.79 L 0.6 0.6 1 Garlic Extract 111 ml/ 3.79 L 0 1 Lemon Juice 15 ml/ 3.79 L 1.8 1.8 0 Azadirachtin 60 ml/ 3.79 L 0.8 0.8 0 Permethrin 9 ml/ 3.79 L 2.0 1.3 1 Permethrin 18 ml/ 3.79 L 0 1 2 More than 10 replicates are needed for statistical analysis.

PAGE 49

35 Table 3-3. Mean number of mole crickets emerging from bermudagrass using various drenching solutions in May 2003. Means statistically significant values using Dunnetts method comparing treatments to water n = 54; F = 2.88; df = 59, 10; P = 0.01 Treatment Rate Mean number of mole crickets flushed ( SEM) Water n/a 0 Citrus oil 15 ml/ 3.79 L 2.6 1.6 Cyfluthrin 7.5 ml/ 3.79 L 0.2 0.2 Cyfluthrin 15 ml/ 3.79 L 4.0 2.1 Garlic extract 111 ml/ 3.79 L 0.4 0.2 Lemon juice 15 ml/ 3.79 L 0.6 0.4 Azadirachtin 60 ml/ 3.79 L 3.2 1.2 Permethrin 9 ml/ 3.79 L 2.6 1.1 Permethrin 18 ml/ 3.79 L 5.8 1.4 Insecticidal soap 15 ml/ 3.79 L 5.4 1.3 Lemon joy 15 ml/ 3.79 L 4.6 2.1 Table 3-4. Percent nematode infection from mole crickets exposed to treatment solutions 1, 5, 8, 12, or 24 h post infection. n = 75; F = 6.77; df = 14, 2; P < 0.0001 Time Post Infection 1h 5h 8h 12h 24h Joy (15 ml/ 3.79 L) 0 40 60* 60* 100* Joy (15 ml/ 3.79 L) + H 2 O rinse 40 40 100* 80* 100* Control 1 0 0 0 0 0 Means within columns statistically significant values when compared to control. 1 Control = uninfected, healthy mole crickets immersed in water.

PAGE 50

CHAPTER 4 INTEGRATION OF INSECT PARASITIC NEMATODES WITH INSECTICIDES FOR CONTROL OF PEST MOLE CRICKETS Appreciation of high-quality turfgrass has recently led to rapid growth in the golf course and landscape management industries (Zimmerman and Cranshaw 1990). Associated with this growth have been an increased number of pesticide applications and environmental concerns (Zimmerman and Cranshaw 1990). Restrictive legislation has resulted in a greater need for alternative control methods. Insect parasitic nematodes provide acceptable control of several soil and thatch infesting pests, such as white grubs (Coleoptera: Scarabaeidae), billbugs (Coleoptera: Curculionidae), cutworms (Lepidoptera: Noctuidae), and mole crickets (Orthoptera: Gryllotalpidae) (Zimmerman and Cranshaw 1990). Insect parasitic nematodes could be effective in integrated pest management programs as long-term suppressive agents used in combination with quick knockdown products like commercially available insecticides. Effects of selected pesticides on entomopathogenic nematodes other than S. scapterisci have been reported in aqueous solutions (Prakasa et al. 1975, Hara and Kaya 1983a, Das and Divakar 1987, Rovesti and Dese 1990, Zimmerman and Cranshaw 1990, Ishibashi and Takii 1993, Gordon et al. 1996, Koppenhfer and Kaya 1998) and in insect hosts following pesticide exposure (Hara and Kaya 1982, 1983b). Several studies have demonstrated that pesticides can decrease survival and infectivity of several entomopathogenic nematode species in the families Steinernematidae and Heterorhabditidae (Hara and Kaya 1982, 1983a, 1983b; 36

PAGE 51

37 Forschler et al. 1990; Zimmerman and Cranshaw 1990; Head et al. 2000). However, combinations of insecticides and insect parasitic nematodes have a synergistic effect on nematode infection rates against white grubs (Koppenhfer and Kaya 1998; Koppenhfer et al. 2000a, 2000b; Koppenhfer et al. 2002). It is important to determine what interactions, if any, might occur when certain insecticides are used together with S. scapterisci. Knowing which insecticides might affect nematode performance is important in the development of integrated pest management programs. Our first objective was to assess the compatibility of chemical and biological management of pest mole crickets. A second objective was to determine if label rates or half-label rates of these insecticides could increase nematode infection rates. Materials and Methods Steinernema scapterisci (Nematac S Becker Underwood, Ames, IA) used in this study was stored in a 7C cold room until use (< 3 mo). Nematode viability was tested before each application by dissolving a pinch of Nematac S into water and observing nematode shape and mobility under a light microscope. Healthy nematodes were opaque in color and S-shaped with undulating movements. Dead or unhealthy nematodes were translucent, straight, and lacked movement. The product was used if viability was >50% and discarded if <50% viable. Scapteriscus vicinus adults were collected from pitfall and sound traps in Alachua County, FL, and returned to the laboratory. Each mole cricket was placed in a 120-ml plastic vial (Thornton Plastics, Salt Lake City, UT) with sterilized sand and held for 14d to ensure health; only mole crickets that appeared healthy were used in this study. Mole

PAGE 52

38 crickets were maintained at 24-27C, with a photoperiod of 12:12 (L:D), and fed commercial cricket chow (Purina St. Louis, MO). Five insecticides commonly used for mole cricket control were evaluated for their effects on the entomopathogenic nematode S. scapterisci in the laboratory. They included acephate (Orthene Turf, Tree and Ornamental Spray, Valent Professional Products, Walnut Creek, CA), bifenthrin (Talstar GC Flowable, FMC Corporation, Philadelphia, PA), imidacloprid (Merit 75 WP, Bayer Environmental Science, Montvale, NJ), fipronil (Chipco Choice granular, Bayer Environmental Science, Montvale, NJ) and deltamethrin (DeltaGard T&O granular, Bayer Environmental Science, Montvale, NJ). These commercially formulated insecticides were selected due to their widespread use for mole cricket control in the southeastern United States. However, only products or formulations that are typically mixed with water or watered in were chosen for this test. Products were tested at half and full label rates. Survival and Infectivity of S. scapterisci After Exposure to Pesticides Nematode viability and infectivity were assessed after exposure to the pesticides. To count nematodes, S. scapterisci were extracted from Nematac S using a modified Baermann funnel technique (Nguyen and Smart 1990b). Approximately 40-50 g Nematac S was placed on two layers of unscented, non-lotion, white facial tissue (Puffs Procter & Gamble, Cincinnati, OH). The tissue was then placed on top of a 1-mm screen filter in a square plastic container (13 13 cm 2 ) and moistened with deionized water. Approximately 100-110 ml of deionized water was added to the plastic container. The setup was then placed on a laboratory counter at 24-27C for 5-8 h, during which

PAGE 53

39 time the living nematodes moved through the tissue and filter into the water below. The nematodes were then counted by taking an average of three, 10-l samples. Steinernema scapterisci were kept at a density of 10,000 infective juveniles per 125 ml in aqueous solutions of water (control), half or full label rates of acephate (1 or 2 kg AI/ha), bifenthrin (112 or 224 g AI/ha), or imidacloprid (275 or 451 g AI/ha). The mixtures were kept at room temperature (24-27 C) in a 125-ml Erlenmeyer flask on a rotator at 65 rpm. There were five replicates for each treatment. After 24 h, 10-l samples were taken from each flask and placed on a microscope slide. The numbers of living and dead nematodes were counted using a dissecting microscope (10 x). Immobile nematodes were touched with a minuten probe to determine survival. Three 10-l counts were averaged to determine percent mortality for each replicate. Nematode infectivity was assessed by removing nematodes from above solutions and adding 50 living infective juveniles to 120 ml plastic cups (Fisher Scientific, Pittsburgh, PA) containing 50 g of sterilized sand moistened with deionized water and one S. vicinus adult. After death, mole crickets were dissected and the presence or absence of nematodes was recorded. Nematode Infectivity After Exposure to Pesticide Treated Mole Crickets Percent infectivity of nematodes when exposed to mole crickets treated with insecticides was also assessed. Mole crickets were placed in 120 ml plastic cups containing 50 g of sterilized sand that was treated with label or half label rates of acephate, bifenthrin, imidacloprid, fipronil (140 or 280 g AI/ha), or deltamethrin (73 or 146 g AI/ha). Pesticides were applied to the top of the sand and mixed into the sand by capping the cup and shaking by hand before adding the mole cricket. The mole crickets were allowed to move through the insecticide treated sand for 24 h. Mole crickets were then removed and placed into new cups containing 50 g of sterilized sand moistened with

PAGE 54

40 deionized water and nematodes. To ensure mole cricket infection the number of nematodes used was increased to 500 infective juvenile S. scapterisci. Mole crickets were fed cricket chow twice a week. Dead mole crickets were removed from the sand cups and placed individually in 20 ml plastic scintillation vials. Mole crickets were examined 5-7 d after death under a dissecting scope (10 X) for the presence or absence of nematodes. The test concluded at 47 d, by which time all of the mole crickets had died. There were five replicates (mole crickets) for each treatment. Statistical Analysis Nematode mortality data were subjected to an analysis of variance (SAS Institute 2001). Treatment means were compared by Tukey-Kramer Honestly Significant Difference means separation test (P= 0.05). Nematode infectivity data are presented as number of crickets infected with S. scapterisci out of five total crickets tested. Nematode mortality data were transformed using arcsine square root transformation before statistical analysis; nontransformed data are presented. Results and Discussion None of the solutions tested reduced S. scapterisci survival (Table 1). Survival ranged from 95.6% (bifenthrin, label rate) to 100% (imidacloprid, half label rate). The average number of days until mole cricket death was not significantly different from the control in any treatment except imidacloprid (Table 1; F=11.83; df=10, 54; P<0.001). None of the infectivity levels were significantly different from the control. Between 2 and 4 of the mole crickets tested became infected with the nematodes from the solutions tested. Mole crickets treated with imidacloprid survived significantly longer than those in the other treatments, but it was not significantly different from the control (Table 2). The percent nematode infection of mole crickets previously exposed to solutions of

PAGE 55

41 insecticides ranged from 40-100% (Table 2; F=14.78; df=10, 54; P<0.001). Infectivity did not significantly differ among any of the treatments tested. These results suggest that the pesticides tested can be successfully used with S. scapterisci applications. Organophosphate insecticides can kill S. carpocapsae (Weiser) infective juveniles (Dutky 1974), or as with carbamates, paralyze them (Hara and Kaya 1982, 1983a). The organophosphate used in our study, acephate, did not have a significant effect on survival and infectivity of S. scapterisci adults. Our results indicate that the pesticides tested in this study are compatible with S. scapterisci infective juveniles. Previous laboratory studies with other steinernematid nematodes (excluding S. scapterisci) have demonstrated compatibility with pesticides (Ishibashi et al. 1987, Zimmerman and Cranshaw 1990, Ishibashi and Takii 1993, Head et al. 2000). Results from field collected mole crickets have demonstrated that S. scapterisci applied to intensively managed bermudagrass (Cynodon spp.) in Gainesville, FL, in October 1989 persisted and were recovered in mole crickets collected in October 2001 (Frank et al. 2002). This evidence along with the results presented here support the compatibility of using S. scapterisci with current mole cricket management strategies. Frank and Parkman (1999) stated that the optimal use of S. scapterisci would be in the roughs of golf courses and areas where tolerance for damage is higher and turf is less intensively maintained. Our results show that an insecticide treated mole cricket has the potential of being infected by nematodes and nematode reproduction does occur within the treated host.

PAGE 56

42 Although our study did not directly test tank-mixing effects, we suggest the potential for tank mixing exists using the wettable or liquid/flowable insecticides tested in this study and Nematac S However, there are several possible limitations to tank-mixing with S. scapterisci. First, chemicals for mole cricket control are usually applied in early summer for control of young nymphs while S. scapterisci is applied in the spring or fall for adult control. Irrigation is recommended before and after Nematac S is applied, thus tank mixing is only possible with pesticides that require watering in. Because nematodes are sensitive to extreme heat and ultraviolet light (Gaugler and Boush 1978), they should be applied at dawn or dusk, which may conflict with applicator work hours. The pesticides tested may provide a quick knockdown for late instar and adult mole crickets, and S. scapterisci may provide a more sustainable, long-term control of mole crickets. An insecticide that could modify an insects behavior to stop or prevent feeding or tunneling damage, and also inhibit the insects ability to defend against parasitism or infection, would be a valuable component of integrated pest management. For example, imidacloprid is synergistic with S. glaseri (Steiner) or Heterorhabditis bacteriophora Poinar against white grubs (Koppenhfer et al. 2000a, 2000b). Imidacloprid disrupts a grubs normal nerve function, which drastically reduces its activity, affects grooming and evasive behaviors, and facilitates nematode attachment onto the cuticle (Koppenhfer et al. 2000a, 2000b). In our study, mole crickets treated with imidacloprid survived longer than those treated with the other insecticides, but still died from nematode infection. Steinernema scapterisci are ambush nematodes, unlike the cruiser nematodes tested by Koppenhfer et al. (2000a, b). Pesticides which increase mole cricket activity, rather

PAGE 57

43 than slow it down, may result in increased contact with ambush nematodes. However, if grooming is also increased, then nematodes may not be able to attach to the host. The half-label rates of acephate and deltamethrin had greater infectivity (Table 2). A more integrated and sustainable management plan for mole crickets is possible using biological, cultural and chemical control. Natural enemies have become established in Florida and actively target adult mole crickets and large nymphs. Insect parasitic nematodes (e.g., S. scapterisci) can be applied as biopesticides for large nymph and adult mole cricket control in the spring and fall. Cultural controls such as using non-attractive lights during adult flight periods, monitoring life stages using pitfall traps or soap flushes in order to time control methods (Hudson 1985), and planting nectar sources for parasitic insects (Frank et al. 1995) should also minimize infestations while selective use of preventive and curative insecticides can quickly control young nymphs during the summer.

PAGE 58

Table 4-1. Percent survival, infectivity, and days until death of S. scapterisci incubated in solutions of insecticides for 24 h. Common Name Trade name Rate Nematode survival (%) SEM (n=5) Days until mole cricket death SEM (n=5) Number of mole crickets infected (n= 5) Acephate Orthene TT&O Spray 1 kg AI/ha 96.3 1.6a 1.4 0.2b 4 2 kg AI/ha 97.5 1.2a 1.4 0.2b 3 Bifenthrin Talstar GC Flowable 112 g AI/ha 95.9 1.7a 2.8 1.0b 3 224 g AI/ha 95.6 1.8a 1.6 0.2b 3 Imidacloprid Merit 75 WP 275 g AI/ha 100.0 0a 23.2 8.9a 2 451 g AI/ha 98.2 1.8a 22.2 4.0a 2 Control (water) n/a n/a 99.8 0.2a 17.0 3.9ab 4 Mean standard error of the mean (SEM), means within columns followed by different letters are significantly different at = 0.05 using Tukeys Honestly Significant Difference means separation test. 44

PAGE 59

Table 4-2. Average days until death and percent infectivity by S. scapterisci nematodes to mole crickets exposed to insecticides. Common Name Trade name Rate Days until mole cricket death SEM (n=5) Number of mole crickets infected (n= 5) Acephate Orthene TT&O Spray 1 kg AI/ha 1.0 0b 3 2 kg AI/ha 1.0 0b 2 Bifenthrin Talstar GC Flowable 112 g AI/ha 1.0 0b 2 224 g AI/ha 1.0 0b 5 Deltamethrin DeltaGard T&O G 73 g AI/ha 1.0 0b 5 146 g AI/ha 1.0 0b 2 Fipronil Chipco Choice G 140 g AI/ha 1.0 0b 2 280 g AI/ha 1.4 0.2b 3 Imidacloprid Merit 75 WP 275 g AI/ha 25.6 9.7a 2 451 g AI/ha 17.4 6.9a 3 Control (water) n/a n/a 39.6 5.0a 2 45

PAGE 60

CHAPTER 5 EFFECT OF Steinernema scapterisci NGUYEN AND SMART EXPOSURE ON MOLE CRICKET TUNNELING, OVIPOSITION, AND AVOIDANCE BEHAVIOR Avoidance of nematodes has been observed in soil inhabiting insects (Thurston 1994, Wang et al. 2002, Zhou et al. 2002). Laboratory evaluation of four entomopathogenic nematodes for control of subterranean termites revealed that Heterorhabditis indica Poinar repelled termites at high concentrations (Wang et al. 2002), but length of repellency varied with nematode concentration. Ants are also repelled by products produced by the bacteria within nematodes (Zhou et al. 2002). The symbiotic bacteria Xenorhabdus nematophila and Photorhabdus luminescens of Steinernema carpocapsae and Heterorhabditis bacteriophora respectively produce ant-deterrent factors (ADF). These ADFs are compounds that deter scavengers and protect the nematodes from being eaten during reproduction within host cadavers (Zhou et al. 2002). Nematodes have the ability to detect and avoid toxic compounds in their environment (Hilliard et al. 2004). The nematode Caenorhabditis elegans has the ability to avoid toxic compounds such as quinine (Hilliard et al. 2004). Soil insects have been observed to sense and avoid areas treated with insecticides, pathogens, or nematodes (Villani et al. 1994, Milner and Staples 1996, Villani et al. 1999, Thompson 2004). Soil insects are often highly susceptible to pathogenic organisms in the laboratory, however are rarely infected in the field, indicating a possible behavioral component of microbial defensive tactics (Villani et al. 1999). Metarhizium anisopliae (Metsch.) incorporated into soil is repellent to Japanese beetle grubs (Villani et al. 1994). 46

PAGE 61

47 Termites also detect and avoid Metarhizium conidia in soil (Milner and Staples 1996). Mole crickets are repelled by soil treated with Beauveria bassiana (Brandenburg 2002, Villani et al. 2002). Thompson (2004) observed mole crickets avoiding soil treated with the Beauvaria bassiana strain 10-22 and Talstar in the greenhouse. This study determined how mole cricket tunneling changes over time after exposure to varying numbers of S. scapterisci, as well as to determine whether healthy mole crickets avoid nematode treated soil and how the oviposition behavior of infected female mole crickets is affected. We wanted to ascertain if mole crickets can sense and avoid nematodes over long distances (ca. 15 cm) and, when given a choice between nematodes and pesticides, would the nematode treated areas be preferred over pesticide treated areas. This information will help in developing a successful IPM program using S. scapterisci and pesticides, as well as determine ways in which application of S. scapterisci could be improved or modified to increase infection rates and reduce pesticide rates. Materials and Methods Steinernema scapterisci (Nematac S Becker Underwood, Ames, IA) used in this study was stored in a 7C cold room until use (< 3 mo). The viability of the nematodes was tested before each application by dissolving a pinch of Nematac S into water and observing nematode shape and mobility under a light microscope. Healthy nematodes are opaque in color and S-shaped with undulating movements. Dead or unhealthy nematodes are translucent, straight, and lack movement. The product was used if viability was >50% and discarded if <50% viable.

PAGE 62

48 Tawny (Scapteriscus vicinus) and southern (Scapteriscus borellii) mole crickets were collected from pitfall traps or sound traps in Alachua Co., FL, during the fall and spring of 2003 and 2004 and returned to the laboratory. Each mole cricket was placed in a 120-ml plastic vial (Thorton Plastics Salt Lake City, UT) with sterilized sand and held for 14 d to ensure health. Mole crickets were maintained at 23C with a photoperiod of 12:12 (L: D) and fed commercial cricket chow (Purina). Scapteriscus abbreviatus were obtained from a laboratory colony at the University of Florida Entomology and Nematology Department and maintained under the same conditions described above. Nematode Infection and Nematode Treated Areas Effect on Mole Cricket Tunneling Behavior Two dimensional tunneling behavior assays were conducted on Scapteriscus vicinus adults that were either healthy or infected with Steinernema scapterisci. Plexiglas containers (30.5 cm wide 30.5 cm high 2.5 cm deep) were filled with autoclaved sand, moistened with deionized water, positioned vertically, kept in a dark area to simulate a subterranean environment, and observed under a red light bulb (40W). Ryegrass (Lolium multiflorum) was grown on top of the containers as a food source for S. vicinus. After the grass established a root system approximately 10-15 cm in length, one adult mole cricket previously exposed to nematodes (0, 500, or 10,000 infective juveniles) was placed in a randomly selected area at the top of the container. Five males and five females were tested for each treatment level. A small strip of Plexiglas was placed in the top opening of the arena to prevent mole cricket escape. Tunnel dimensions in the sand were measured at 1, 4, 8, 24, 48, 72, and 96 h then every 48 h after that for 10 d or until mole cricket death, whichever occurred first. Tunnel patterns were traced on acephate paper, and then lengths were measured with a cloth measuring tape. Time that

PAGE 63

49 tunnel construction slowed down or ceased was recorded. Daily tunnel distances were determined by subtracting total tunnel distance from the previous day's tunnel distance. Mole crickets were then removed from the arenas and assayed for nematode infection. There were 10 replicates (crickets) for each treatment. Additional tests were completed to determine whether healthy mole crickets could sense and/or avoid nematode-treated soil. Autoclaved builders sand was colored with blue-powdered chalk (American Tool Companies, Inc., Wilmington, OH) and placed on one half of a 30.5 cm wide 30.5 cm high 2.5 cm deep Plexiglas arena with the other half containing sand colored with orange powder (American Tool Companies, Inc., Wilmington, OH). Ryegrass was grown on top of the containers to provide food for the mole crickets. The blue colored sand was inoculated with a label rate (approximately 25,000 nematodes/cm 2 ) of S. scapterisci by injecting approximately 100 ml of nematodes in solution to one corner of the container using a graduated cylinder. To determine if there were any deterrent effects of the chalk used to color the sand, control arenas were setup similar to the treated arenas. Healthy mole crickets were placed in the control arena with colored sand, however both sides of the arenas lacked nematodes. The controls were treated exactly as the treatment arenas. The containers were kept in a dark area to simulate subterranean conditions and observed under red light (40 W). One mole cricket was placed on the untreated side of the Plexiglas arena. Half of the replicates contained orange sand treated with nematodes with untreated blue sand and the other half contained blue sand treated with nematodes and untreated orange sand. Tunnels and movement of nematode infected sand was compared to movement of untreated sand by looking at the tunnel patterns and movement

PAGE 64

50 of the treated sand. Mole cricket behavior was observed at 1, 4, 8, 24, 48, 72, and 96 h then every 48 h after that for 10 d or until mole cricket death, whichever occurred first. Data were summarized into 48 h increments. The length of the tunnel system was determined as previously described. After 10 d mole crickets were removed from the arenas and assayed for nematode infection. There were ten replicates (mole crickets). Mole crickets used in this study were dissected after 10 d (or after death, whichever occurred first). Oviposition Behavior of Mole Crickets Exposed to S. scapterisci The oviposition behavior including number of eggs, clutch size, depth and width of egg chambers, and time until oviposition was also observed in the previous tests. After 10 d the arenas were dismantled and egg clutches were removed. An indentation was made in 55 mm Petri dishes filled with moistened sterilized sand by pressing a gloved finger into the moistened sand. Eggs were carefully removed using feather-weight forceps and placed into the depression. Petri dishes were covered and wrapped with parafilm (Fisher Scientific, Pittsburgh, PA) to keep humidity levels high. Eggs were monitored every 48 h until egg hatch. Y-Tube Tests Observation Chamber Response of mole crickets to potentially lethal compounds was tested in a dual choice observation chamber intended to simulate below ground tunnel conditions. Y-tube arenas were constructed to evaluate the response of Scapteriscus borellii, S. vicinus, and S. abbreviatus to Steinernema scapterisci or untreated sand, and the choice between nematodes or pesticides. Arenas were constructed of 1.27 cm diameter plastic tubing (Fisher Scientific, Pittsburgh, PA), the basal section was 15 cm long, and each arm was

PAGE 65

51 10 cm long. The tubing was connected to form a Y using a 1.27 cm plastic y-connector (Fisher Scientific, Pittsburgh, PA). Autoclaved builders sand was loosely packed in either arm of the Y-tube. Comparisons consisted of 500 against 0 nematodes. Nematodes were added to the sand using a pipet. Each arm was capped with a 20 ml plastic scintillation vial (Fisher Scientific, Pittsburgh, PA). The 15 cm entry arm was left void of sand to facilitate mole cricket movement. Ten replicates of S. borellii and S. vicinus were completed for each level of nematodes; each replicate was completed using a different cricket. Experiments were carried out at 23 C in dark conditions using red light (40W) to observe the mole crickets. One adult S. borellii or S. vicinus was placed into the basal section of the Y-tube. The cricket was observed as it moved through the tubing at the base and then chose between the arms containing treatments. Choice was determined when a cricket traveled completely through one arm and entered the scintillation vial at the end. Y-tubes were cleaned with a 1% bleach solution between runs and re-used for nematode studies only. Response to S. scapterisci or Pesticides Five insecticides commonly used for pest mole cricket control were evaluated in the laboratory against nematodes for their attractive or repellent characteristics to mole crickets. The pesticides tested were acephate (0.91 kg AI/ha) (Orthene Turf, Tree and Ornamental Spray, Valent Professional Products, Walnut Creek, CA), bifenthrin (224 g AI/ha) (Talstar GC Flowable, FMC Corporation, Philadelphia, PA), imidacloprid (451 g AI/ha) (Merit 75 WP, Bayer Environmental Science, Montvale, NJ), fipronil (280 g AI/ha) (Chipco Choice, Bayer Environmental Science, Montvale, NJ) and deltamethrin (146 g AI/ha) (DeltaGard T&O, Bayer Environmental Science, Montvale, NJ). Autoclaved builders sand was loosely packed in one arm of the Y-tube and 10,000

PAGE 66

52 nematodes were added using a pipet. The second arm contained autoclaved builders sand mixed with 1 ml of the label rate of the above pesticides. Each arm was capped with a 20 ml plastic scintillation vial (Fisher Scientific). The 15 cm basal entry arm was empty. Seven replicates were completed for each insecticide; each replicate was conducted using a naive cricket. Experiments were conducted using the same methods described in the previous Y-tube tests. Y-tube arenas used in pesticide studies were discarded after use to prevent contamination. Statistical Analysis Tunneling and oviposition data were subjected to an ANOVA (SAS Institute 2001) and means were separated using the Tukey-Kramer Honestly Significant Difference means separation test. Within each Y-tube test, the number of crickets responding to each choice was analyzed by a chi-square test against the null hypothesis of a 1:1 ratio (Steel and Torrie 1981). Results Effect of Nematode Infection and Nematode Treated Areas on Mole Cricket Tunneling Behavior Tunneling rate of mole crickets treated with 0, 500, 10,000 nematodes did not significantly differ over time among the treatments (Figure 5-1). Total distance tunneled over the entire 10 d period (29-44 cm) was not statistically significant among the three treatments (F= 0.87; df= 2, 35; P= 0.43). Mole crickets treated with 500 or 10,000 S. scapterisci tended to tunnel less (lower average daily distance) when compared to healthy mole crickets (Figure 5-2). There were no significant differences between male and female mole cricket tunneling when exposed to different levels of nematodes (F=1.073; df= 5, 23; P=0.408). During the 10 d test, the eight mole crickets treated with 0, 500, or

PAGE 67

53 10,000 nematodes had 0, 1, and 3 crickets respectively infected with nematodes. Tunnel distances began to decrease after 6 d in crickets treated with 500 or 10,000 nematodes. Mole crickets did not appear to differentiate between untreated and nematode-treated sand (F= 1.93; df=1, 79; P= 0.1684). Tunnel length did not differ in either side of the arena at anytime during the test (Figure 5-3). There were no significant effects on tunneling or viability of crickets in arenas with sand lacking nematodes and tinted with orange or blue colored chalk (F=2.94; df=1, 49; P=0.5899). Several mole crickets did not venture far from the top 10 cm, and these crickets were the first to die. All mole crickets were infected with S. scapterisci. Oviposition Behavior of Mole Crickets Exposed to S. scapterisci Mole crickets infected with 500 or 10,000 nematodes were able to lay eggs, to about the same soil depth, in the same chamber dimensions, and in a similar quantity to uninfected females (Table 5-1). These behaviors also did not differ in treated vs. untreated sand (500 or 10,000 nematodes) (Table 5-2). Healthy mole crickets and those treated with 10,000 nematodes laid eggs between 1-6 d. Crickets treated with 500 nematodes laid eggs between 1-10 d. One female cricket treated with 500 nematodes either buried or consumed her clutch of eggs during the study period. None of the eggs survived due to mold growth. Y-Tube Tests None of the three mole cricket species tested demonstrated a significant choice between arms treated with 500 or 10,000 S. scapterisci (S. borellii, S vicinus, or S. abbreviatus; 2 = 2.4, 2.1, and 1.6 respectively for 500 nematodes; S. abbreviatus and S. vicinus; 2 = 1.6, and 2.4, respectively for 10,000 nematodes). Most mole crickets tested chose the arm that contained S. scapterisci when given a choice between S. scapterisci or

PAGE 68

54 insecticides (Table 5-4). Crickets, when given a choice between S. scapterisci and fipronil chose the arm containing fipronil. Fipronil granules did not readily dissolve in water and the solution tended to stay at the very distal end of the arm reducing contact with the mole cricket. Discussion Although mole crickets have been observed avoiding entomopathogens such as Beauveria bassiana and Metarhizium anisopliae (Villani et al. 1999, Brandenburg 2002, Villani et al. 2002, Thompson 2004), they do not appear to avoid the insect parasitic nematode S. scapterisci. In this study we sought to determine if infection of Scapteriscus spp. with S. scapterisci caused any modification of tunneling behavior. Although the results were not statistically significant, the crickets infected with 10,000 nematodes tended to tunnel less than crickets treated with 500 nematodes and healthy crickets. The shape and length of the tunnels formed by S. borellii concur with Brandenburg et al. (2002). Both healthy and nematode-infected crickets typically formed an inverted y-shaped tunnel with branching occurring lower than the surface of the arenas. Our results show that crickets infected for >6 d may not tunnel as much or as deep as uninfected mole crickets. This shallow tunneling behavior may make the mole crickets more vulnerable to insecticides, management practices such as aeration, or increase vulnerability to predators such as birds and armadillos. Because insect parasitic nematodes are slower-acting than chemical pesticides the mole crickets may continue to oviposit during the period of infection before death. Our results indicate no significant differences in the ovipositional behavior of female crickets. Females infected with nematodes may not have the energy or resources available for egg laying because these resources are required for immune system defense and the female

PAGE 69

55 may die before completing oviposition. Research on Beauveria bassiana has involved the effects of infection on oviposition behavior of Lygus hesperus (Hemiptera: Miridae) an important crop pest in the western United States (Noma and Strickler 2000). Lygus hesperus oviposition is significantly reduced after inoculation with B. bassiana in the laboratory. Nematodes in the genus Steinernema have also resulted in reduced overall host fecundity (Belair and Boivin 1995, Kim et al. 2004). Scapteriscus borellii adults laid a similar number of eggs in nematode treated and untreated sand in the Plexiglas arenas, indicating a lack of repellency by the nematodes. Thus, a female ovipositing in soil treated with Nematac S may then become infected, although her offspring are not likely to be immediately affected. We were unable to assess egg viability in these tests due to excessive mold development (none of the eggs hatched). Mole crickets have been observed avoiding insecticides (Brandenburg 2002; Thompson 2004). If mole crickets can sense and avoid insecticides applied to tees, greens, and fairways this may potentially force the cricket into the roughs where nematode applications could be made. In our tests, when given a choice, mole crickets chose sand treated with S. scapterisci over sand treated with the insecticides tested. The sensory system of mole crickets is not well understood (Villani et al. 1999; Brandenburg 2002; Villani et al. 2002; Thompson 2004). Initial scanning electron microscope photographs (Appendix D) indicate many hairs covering the antennae, tarsi, and palps. These hairs may be due to the insects subterranean habitat and not for sensory detection. Sensory detection in mole crickets should be studied further. Knowledge of mole cricket sensory detection would be useful in development of a mole cricket IPM program.

PAGE 70

56 The fact that Scapteriscus borellii oviposited in a laboratory arena is also new information. The common rearing method for S. abbreviatus in the University of Florida colony in Gainesville, FL is in cylindrical 120-ml plastic vials with autoclaved sand. This method was unsuccessful for the author when rearing S. borellii or S. vicinus; however oviposition was observed in females placed in the Plexiglas sandwich arenas used for this study. The larger arenas may provide more room for the crickets to move around; also females could tunnel down 30.5 cm versus only several centimeters in the plastic vials. The moisture level also remained relatively high in the Plexiglas arenas. High humidity is necessary for successful mole cricket oviposition (Hertl et al. 2001).

PAGE 71

57 0102030405060246810DayDistance Tunneled (cm) 0 nematodes 500 nematodes 10,000 nematodes Figure 5-1. Distance (cm) tunneled ( SEM) through sand in Plexiglas arenas every 2 d by mole crickets exposed to varying amounts of nematodes. ANOVA for 2,4,6,8, and 10 d are (F=0.45; df=2,34; P=0.64); (F=1.45; df=2,32; P=0.25); (F=0.32; df=2,31; P=0.73); (F=0.61; df=2,31; P=0.55); (F=1.49; df=2,29; P=0.24); respectively.

PAGE 72

58 010203040506070050010000Number of Nematodes Total Distance Tuneled (cm) Female Male Figure 5-2. Comparison of total tunnel distances ( SEM) by male and female Scapteriscus spp. mole crickets after exposure to S. scapterisci. 0510152025303515152448Time (h)Total Distance Tunneled (cm) Treated Untreated Figure 5-3. Distance tunneled ( SEM) by mole crickets through sand treated with nematodes and untreated sand in Plexiglas arenas over 48 h.

PAGE 73

59 Table 5-1. Oviposition of Scapteriscus mole crickets directly infected with different numbers of S. scapterisci. Treatment (# nematodes) # Mole crickets that laid eggs (n=8) Number of eggs ( SEM) 1 Width of egg chamber (cm) ( SEM) 2 Depth of egg chamber (cm) ( SEM) 3 0 5 17.25 5.36 2.81 1.07 12.93 4.14 500 6 19.25 5.01 2.74 0.91 11.34 2.71 10,000 4 12.63 4.85 1.70 0.66 7.99 3.26 1 F = 0.14; df = 2,14; P = 0.87 2 F = 0.35; df = 2,14; P = 0.67 3 F = 1.82; df = 2,14; P = 0.20 Table 5-2. Oviposition of Scapteriscus mole crickets in sand treated with S. scapterisci. Treatment # Mole crickets that laid eggs (n=5) Number of eggs ( SEM) 1 Width of egg chamber (cm) ( SEM) 2 Depth of egg chamber (cm) ( SEM) 3 Treated sand 3 33.5 17.99 4.5 0.42 15.5 1.89 Untreated sand 5 24.0 25.45 5.0 0.60 19.5 2.68 1 F = 0.970; df = 2, 4; P = 0.506 2 F = 52.600; df = 2,4; P = 0.019 3 F = 77.933; df = 2,4; P = 0.013 Table 5-3. Response of Scapteriscus vicinus, S. borellii, and S. abbreviatus to Steinernema scapterisci nematodes versus sterilized sand. Species tested # crickets selecting the y-tube arm 1 S. scapterisci Sterilized sand 2 S. vicinus 3 7 2.1* S. borellii 6 4 2.4* S. abbreviatus 2 8 1.6** All species combined 11 19 6.3 1 Response of 10 individuals of each mole cricket species when given a choice between two arms. *P<0.01, **P<0.005

PAGE 74

60 Table 5-4. Response of Scapteriscus vicinus to Steinernema scapterisci nematodes versus pesticides treated sand. S. scapterisci Insecticide No choice Acephate 7 0 0 Bifenthrin 3 4 0 Deltamethrin 3 1 2 Fipronil 3 4 0 Imidacloprid 5 1 1 Pesticide tested Y-tube arm selected 1 1 Response of 7 mole crickets when given a choice between two arms. More than 10 replicates are needed for statistical analysis.

PAGE 75

CHAPTER 6 SUMMARY AND CONCLUSIONS Steinernema scapterisci is an insect parasitic nematode that has been formulated by Becker Underwood as the biopesticide Nematac S. It differs from other commercially-available nematodes used for mole cricket control by being able to reproduce in the hosts body and persisting in the soil after application. Thus, after being sprayed like a pesticide to infested turfgrass, the nematodes may continually help to suppress mole crickets as a natural enemy. Nematac S was originally marketed for use on pastures because insecticides were not a cost effective control option. To expand the marketing and use of Nematac S, golf courses, athletic fields, parks, lawns and other turfgrass areas were also considered. The research conducted in this dissertation is relevant primarily to the golf course industry, but results may be extrapolated into the other turfgrass sites. The intensity of management on Florida golf courses ranges from high on private courses to somewhat lower on municipal or public courses. Golfers, golf pros, and superintendents tend to have little tolerance for damage on their greens, tees, and fairways. Roughs and driving ranges, however, may receive less attention and may therefore be more ideal areas for introducing or conserving natural enemies. This research has demonstrated that S. scapterisci can survive on intensively managed golf courses for more than ten years. Most of the insecticides (e.g., acephate, bifenthrin, deltamethrin, fipronil, imidacloprid) used against mole cricket nymphs and adults do not reduce nematode survival or infectivity. But, products used to control plant parasitic 61

PAGE 76

62 nematodes would undoubtedly also kill beneficial nematodes. Perhaps this particular problem, which many golf courses deal with, is why insect parasitic nematodes have not been heavily used on golf courses in the past. The beneficial effects of S. scapterisci increase in an additive fashion over time. Steinernema scapterisci can survive and reproduce within the soil and can be transported to new areas via mole cricket movement; therefore its benefits are seen year after year, unlike pesticides which need to be reapplied. Golf courses unknowingly benefit from nematode populations by an overall reduction in mole cricket populations over time. The reduction in mole cricket populations would then reduce the pesticide applications required for mole crickets possibly to the level of applying hot spot applications only. Steinernema scapterisci is an important part of any mole cricket integrated pest management program.

PAGE 77

APPENDIX A AMBIENT DATA AND TURFGRASS QUALITY DATA COLLECTED AT GAINESVILLE GOLF AND COUNTRY CLUB AND IRONWOOD GOLF COURSE Table A-1. Ambient data collected from Gainesville Golf and Country Club on dates of mole crickets collections. Date Air Temp. 1 (C) Rel. Humid. 1 (%) Soil Temp. 1 (C) Rainfall 2 (cm) October 2001 20.2 68.0 21.1 0.3 November 2001 15.3 60.0 14.6 2.7 December 2001 13.6 75.7 16.1 3.8 January 2002 12.8 64.5 13.3 13.3 February 2002 19.6 72.0 16.4 2.8 March 2002 24.1 80.0 15.3 8.3 April 2002 21.1 74.5 21.1 1.0 May 2002 21.0 79.3 20.4 4.0 July 2002 23.9 68.0 23.3 13.8 August 2002 23.3 82.2 23.9 26.5 September 2002 25.0 68.0 25.6 12.8 October 2002 22.6 74.5 22.5 4.2 1 Average of monthly data taken on each collection date. Soil temp taken at 7.6 cm below surface. 2 Average monthly rainfall value from data collected at the Gainesville Regional Airport. 63

PAGE 78

64 Table A-2. Ambient data collected from Ironwood Golf Course on dates of mole crickets collections. n/a indicates data not collected for that date. Date Air Temp. 1 (C) Rel. Humid. 1 (%) Soil Temp 1 (C) Rainfall 2 (cm) October 2001 24.9 61.3 23.7 0.3 November 2001 18.9 64.8 16.1 2.7 December 2001 15.0 80.4 15.0 3.8 January 2002 n/a n/a 13.1 13.3 February 2002 17.9 84.0 16.7 2.8 March 2002 16.8 66.5 16.1 8.3 April 2002 23.9 72.6 22.2 1.0 May 2002 18.5 94.3 20.9 4.0 July 2002 23.3 93.0 23.3 13.8 August 2002 24.2 74.0 24.6 26.5 September 2002 25.1 56.0 23.3 12.8 October 2002 22.6 60.5 24.0 4.2 November 2002 11.7 78.5 11.1 13.3 December 2002 13.3 93.0 10.0 18.3 January 2003 3.1 68.0 7.22 0.5 February 2003 10.6 60.0 5.0 16.8 March 2003 21.1 68.5 17.2 19.5 April 2003 16.2 76.0 17.2 4.1 May 2003 19.3 74.0 20.6 5.1 1 Average of monthly data taken on each collection date. Soil temp taken at 7.6 cm below surface. 2 Average monthly rainfall value from data collected at the Gainesville Regional Airport.

PAGE 79

65 Table A-3. Average turfgrass density ratings for treated and untreated plots on Gainesville Golf and Country Club and Ironwood Golf Course. 1=poor; 5=acceptable; 9=excellent. n/a indicates data not taken. Gainesville Golf and Country Club Ironwood Golf Course Collection Date treated plots untreated plots treated plots untreated plots October 2001 5.7 5.2 5.0 3.7 November 2001 7.7 7.6 6.2 4.4 December 2001 6.7 6.3 6.0 4.9 January 2002 5.7 5.7 6.0 4.8 February 2002 6.1 6.2 5.9 4.8 March 2002 6.8 5.9 5.0 4.3 April 2002 6.3 5.6 4.8 3.7 May 2002 6.5 5.4 5.4 3.8 July 2002 8.7 6.8 6.4 5.5 August 2002 5.3 5.5 5.9 5.1 September 2002 5.8 6.3 5.7 5.0 October 2002 4.9 4.6 5.6 5.0 November 2002 n/a n/a 5.2 5.1 December 2002 n/a n/a 6.2 4.8 January 2003 n/a n/a 5.3 3.4 February 2003 n/a n/a 5.5 4.6 March 2003 n/a n/a 4.6 3.6 April 2003 n/a n/a 4.6 3.6 May 2003 n/a n/a 4.5 3.4

PAGE 80

APPENDIX B DATA FROM ATHLETIC FIELD DEMONSTRATION SITES Materials and Methods Objective A demonstration project was conducted on athletic field turfgrass. Methods were similar to those in Chapter 2 of this dissertation; however fewer replicates were used due to limited space. Nematodes are an important part of an athletic field IPM program because they are a non-chemical way of reducing mole cricket populations. This in turn reduces the exposure to children who use these fields. Study Site The establishment and spread of S. scapterisci was monitored on two baseball fields located in Sarasota, FL (Twin Lakes Park) and Altamonte Springs, FL (Eastmonte Park). Both baseball fields were open to the public. The Sarasota field was built around 1975 and the Altamonte Springs field was built around 1970. The Sarasota field was common bermudagrass (Cynodon dactylon [L.] Pers.) and the Altamonte Springs field was bermudagrass (Cynodon dactylon C. transvaalensis) var. Tifway. Both parks were mowed at 1.91 cm. Neither park had previously received Steinernema scapterisci treatments. Mole Cricket Monitoring Three hot spots of mole cricket activity were located in each baseball outfield. Linear pitfall traps (modified from Lawrence 1982, described in Chapter 2) were installed 66

PAGE 81

in the ground at least 100 m apart. Each field contained two treated areas and one control area. Traps on all sites were installed in September 2001. Nematodes were released in the morning (approximately 0700). Nematodes were applied in an aqueous suspension of 1 billion nematodes/ 378.5 L of water applied using a boom sprayer calibrated at 40 L/1000 m 2 The area treated was 20.1 20.1 m (404.7 m 2 ) around each trap. Treated areas were irrigated with 0.6 cm of water before and 0.6 cm after application. The pre-treatment dates for Sarasota were 11,18 October 2001 and 11 October 2001 for Altamonte Springs. Nematodes were applied on 11 November 2001 at Sarasota and 25 October 2001 at Altamonte Springs. A second application of nematodes was made on the Sarasota field 11 October 2002. Pitfall traps were sampled to monitor infection levels and mole cricket abundance using methods similar to Parkman et al. (1993a,b). At each 24 h sampling period the buckets and arms were cleaned out and 3 to 5 cm of sand was placed into the inner bucket. Traps were left for 24 h and all live mole crickets were collected. Mole crickets were placed individually in 20 ml plastic scintillation vials (Fisher Scientific) with 1-2 drops of deionized water, capped and labeled and mailed via overnight courier delivery to Gainesville, FL. The nematode doesnt move much in the soil and cant be identified through soil samples, so adult and juvenile mole crickets with pronotal lengths > 4 mm (Hudson and Nguyen 1989a) were collected from traps, using the same methods as in the pretreatment sampling, and tested for infection weekly for the first 6 weeks post-application and one to two times a month thereafter for 2 yr. Turf quality (density, color) in the area immediately surrounding the pitfall traps was assessed (1 to 9 scale with 1 sparse or brown grass, 9 dense or dark green grass). A second application of S.

PAGE 82

scapterisci was made on 11 October 2002 at Sarasota due to low levels of infection in collected mole crickets. The percentage of infected mole crickets was determined at 7 and 10 d after death under a dissecting scope (10 X). Steinernema scapterisci were identified by Dr. Khuong Nguyen, Entomology and Nematology Department, University of Florida. Results and Discussion It took approximately 8 wk for infection levels to reach 25% in the mole crickets collected at the Sarasota site. Levels remained at or near this for about 1 yr post-initial application. After the second application in October 2002 levels reached 25% (Figure B-1) and remained at this level in the following spring mole cricket population (March and April 2003). The baseball field had heavy rains and flooding as well as a non-functional irrigation system in 2002. Infection level may have been higher if these factors had not occurred. Very low levels of mole cricket damage/activity were observed in spring through fall 2003 when compared to previous years. Levels of infection were also low in the summer months due to the small size of the crickets present during these months (see Chapter 2). A second application of nematodes may be necessary to reestablish nematode populations high enough in the soil to result in infection levels of 15-30% in collected mole crickets. Data from the Altamonte Springs site was terminated early due to the very low numbers of crickets (< 20) collected during the research period. The study on this site was canceled in May 2002. Statistics presented in both graphs are comparisons of monthly means (SAS Institute 2001).

PAGE 83

69 Sarasota Athletic Field051015202530354045Oct-01Nov-01Dec-01Jan-02Feb-02Mar-02Apr-02May-02Jun-02Jul-02Aug-02Sep-02Oct-02Nov-02Dec-02Jan-03Feb-03Mar-03Apr-03May-03Jun-03Jul-03Aug-03Sep-03Oct-03Nov-03Month% Infected Mole Crickets Figure B-1. Average monthly infection rates at the Sarasota athletic field research site. Arrows indicate Steinernema scapterisci applications. F=3.1212; df=24,92; P=0.0001

PAGE 84

Altamonte Springs020406080100120OCT 01NOV 01DEC 01JAN 02FEB 02MAR 02APR 02MAY 02Month% Infected Mole Crickets 70 Figure B-2. Average monthly infection rates at the Altamonte Springs athletic field research site. Arrow indicates Steinernema scapterisci applications. F=9.35; df=6,15; P=0.002

PAGE 85

APPENDIX C PRELIMINARY CHECKLIST OF ARTHROPODS ASSOCIATED WITH GOLF COURSE TURFGRASS Below is a preliminary checklist of arthropods found in linear pitfall traps located on Gainesville Golf and Country Club and Ironwood Golf Course in Gainesville Florida from November 2001 through October 2003. List is considered preliminary and incomplete due to limited time in which the identifications could be completed. Class Order Family: subfamily Genus: species Insecta Coleoptera Carabidae Omophron labiatum Pasimachus spp. Curculionidae Sphenophorus spp. Elateridae Meristhus squamiger Geotrupidae Peltotrupes profundus Histeridae Lampyridae Ptiliidae Scarabaeidae Ataenius spp. Cotinis nitida Cyclocephala spp. Phylophaga spp. Tomarus subtropicus Silvanidae Staphylinidae Tenebrionidae Tenebrionidae:Diaperinae Tenebrionidae:Alleculinae Dermaptera Labiduridae Labidura riparia Forficula auricularia Lepidoptera Geometridae Mocis spp. Noctuidae Herpetogramma phaeopteralis Pyralidae Spodoptera frugiperda Saturnidae Arachnida Agelenidae Anyphaenidae Corinnidae Dictynidae 71

PAGE 86

72 Class Order Family: subfamily Genus: species Arachnida Gnaphosidae Hahniidae Lycosidae Mimetidae Mysmenidae Pisauidae Tetrognathidae Theridiidae Zorocratidae

PAGE 87

APPENDIX D SCANNING ELECTRON MICROGRAPH PICTURES OF MOLE CRICKET SENSORY AREAS Scanning electron micrograph photographs were taken using a tungsten low vacuum scanning electron microscope model JSM-5510LV (JEOL-USA, Peabody, MA) to determine if any significant sensory hairs, pores, or other morphological aspects were apparent on mole crickets. Pictures were taken of dried, curated mole crickets focusing on the antenna of an adult male and one adult female, a female mid-tarsomere, and a female labial palpomere. These photographs are only an initial look at mole cricket sensory organs and more in-depth photographs should be taken for future study. Figure D-1. SEM photograph of a female Scapteriscus vicinus antennal mid-section. 73

PAGE 88

74 Figure D-2. SEM photograph of a male Scapteriscus vicinus antennal mid-section. Figure D-3. SEM photograph of a female Scapteriscus vicinus mid-tarsal claw.

PAGE 89

75 Figure D-4. SEM photograph of a female Scapteriscus vicinus labial palpomere.

PAGE 90

LIST OF REFERENCES Adeji, M. B., J. H. Frank, and C. S. Gardner. 2003. Survey of pest mole cricket (Orthoptera: Gryllotalpidae) activity in South-Central Florida. Florida Entomol. 86: 199-205. Akhurst, R. J. 1990. Safety to nontarget invertebrates of nematodes of economically important pests, pp. 233-240. In M. Laird, L. A. Lacey, and E. W. Davidson (eds.), Safety of microbial insecticides. CRC Press, Boca Raton, FL. Ames, L. M. 1990. The role of some abiotic factors in the survival and motility of Steinernema scapterisci. M. S. Thesis. University of Florida, Gainesville, FL. Arthurs, S. and K. M. Heinz. 2003. Thrips parasitic nematode Thripinema nicklewoodi (Tylenchida: Allantonematidae) reduces feeding, reproductive fitness, and tospsvirus transmission by its host, Frankliniella occidentalis (Thysanoptera: Thripidae). Environ. Entomol. 32: 853-858. Bedding, R. A. 1993. Biological control of Sirex noctilio using the nematode Deladenus siricidicola. pp. 11-20. In R. Bedding, R. Akhurst, and H. Kaya (eds.), Nematodes and the biological control of insect pests. CSIRO Publications, East Melbourne. Belair, G. and G. Boivin. 1995. Evaluation of Steinernema carpocapsae Weiser for control of carrot weevil adults, Listronotus oregonensis (LeConte) (Coleoptera: Curculionidae), in organically grown carrots. Biocon. Sci. Tech. 5: 225-231. Benton, M. J. and G. Pritchard. 1990. Mayfly locomotory responses to endoparasitic infection and predator presence: the effects on predator encounter rate. Freshwater Biol. 23: 363-371. Boucias, D. G. 1985. Diseases. pp 3-10. In T.J. Walker (ed.), Mole crickets in Florida. Florida Agric. Exp. Sta. Bull. 846. Braman, S. K. 1993. Azadirachtin affects growth and survival of immature tawny mole crickets (Orthoptera: Gryllotalpidae). Florida Entomol. 76: 526-530. Braman, S. K., R. R. Duncan, W. W. Hanna, and W. G. Hudson. 2000. Evaluation of turfgrasses for resistance to mole crickets (Orthoptera: Gryllotalpidae). HortScience 35: 665-668. Brandenburg, R. L. 2002. Improving mole cricket management by targeting their weaknesses. USGA Turf. and Environ. Res. 1: 1-8. 76

PAGE 91

77 Brandenburg, R. L., Y. Xia, and A. S. Schoeman. 2002. Tunnel architectures of three species of mole crickets (Orthoptera: Gryllotalpidae). Florida Entomol. 85: 383-385. Capinera, J. L. and N. C. Leppla. 2001. Pest Mole Crickets. http://creatures.ifas.ufl.edu/ Accessed 12 February 2002. Cobb, P. P. and T. P. Mack. 1989. A rating system for evaluating tawny mole cricket, Scapteriscus vicinus Scudder, damage (Orthoptera: Gryllotalpidae). J. Entomol. Sci. 24: 142-144. Creighton, C. S. and G. Fassuliotis. 1985. Heterorhabditis sp. (Nematoda: Heterorhabditidae): a nematode parasite isolated from the banded cucumber beetle Diabrotica balteata. J. of Nematol. 17: 150-153. Das, J. N. and B. J. Divakar. 1987. Compatibility of certain pesticides with DD-136 nematode. Plant Prot. Bull. 39: 20-21. Denisenko, M. K. 1986. An effective method of controlling mole crickets. Zaschchita Rastenii 10: 51. Dutky, S. R. 1974. Nematode parasites. pp. 576-590. In F. G. Maxwell and F. A. Harris (eds.), Proceedings of the Summer Institute on Biological Control of Plant Insects and Diseases. Univ. Press of Mississippi, Jackson. English-Loeb, G., M. Villani, T. Martinson, A. Forsline, and N. Consolie. 1999. Use of entomopathogenic nematodes for control of grape phylloxera (Homoptera: Phylloxeridae): a laboratory evaluation. Environ. Entomol 28: 890-894. Epsky, N. D. and J. L. Capinera. 1994. Invasion efficiency as a measure of the efficacy of the entomogenous nematode Steinernema carpocapsae (Rhabditida: Steinernematidae). J. Econ. Entomol. 87: 366-370. Forrest, T. G. 1985. Reproductive behavior. pp 3-10. In T. J. Walker (ed.), Mole crickets in Florida. Florida Agric. Exp. Sta. Bull. 846. Forschler, B. T., J. N. All, and W. A. Gardner. 1990. Steinernema feltiae activity and infectivity in response to herbicide exposure in aqueous and soil environments. J. Inv. Path. 55: 375-379. Frank, J. H. 1990. Mole crickets and other arthropod pests of turf and pasture. pp. 131-139. In D. H. Habeck, F. D. Bennett, and J. H. Frank (eds.), Classical biological control of insects and weeds in the southern United States. South. Coop. Ser. Bull. 355. Frank, J. H. 2001. Statewide controls for mole crickets? Florida Turf Digest. July/August 2001.

PAGE 92

78 Frank, J. H., E. A. Buss, and K. A. Barbara. 2002. Compound interest beneficial nematodes in turf: good for how many years against pest mole crickets? Florida Turf Digest 19: 48-50. Frank, J. H., T. R. Fasulo and D.E. Short. 1998. MCricket Knowledgebase. University of Florida, Institute of Food and Agricultural Sciences. Available: http://molecrickets.ifas.ufl.edu/index.htm Accessed: 6 December 2001. Frank, J. H. and J. P. Parkman. 1999. Integrated pest management of pest mole crickets with emphasis on the southeastern USA. Integrated Pest Management Reviews 4: 39-52. Frank, J. H., J. P. Parkman and F. D. Bennett. 1995. Larra bicolor (Hymenoptera: Sphecidae), a biological control agent of Scapteriscus mole crickets (Orthoptera: Gryllotalpidae) established in northern Florida. Fla. Entomol. 78: 619-623. Frank, J.H., T.J. Walker and J.P. Parkman. 1996. The introduction, establishment, and spread of Ormia depleta in Florida. Biol. Control 6: 368-377. Gaugler, R. and G. M. Boush. 1978. Effects of ultraviolet radiation and sunlight on the entomogenous nematode, Neoplectana carpocapsae. J. Inv. Pathol. 32:291-296. Gaugler, R. and H. Kaya (eds.). 1990. Entomopathogenic nematodes in biological control. CRC, Boca Raton, FL. Georgis, R. 1997. Commercial prospects of microbial insecticides in agriculture. Proc. Br. Crop. Prot. Council Symp. 68: 243-252. Georgis, R. and R. Gaugler. 1991. Predictability in biological control using entomopathogenic nematodes. J. Econ. Entomol. 84: 713-720. Georgis, R., and H. K. Kaya. 1998. Formulation of entomopathogenic nematodes. pp. 289-308. In H. D. Burges (ed.), Formulation of microbial biopesticides. Kluwer Academic, Dordrecht. Georgis, R. and S. A. Manweiler. 1994. Entomopathogenic nematodes: a developing biological control technology. Vol. 6, pp. 63-94. In Agricultural Zoology Reviews, K. Evans, (ed.) Intercept, Andover. Gordon, R., J. Chippett, and J. Tilley. 1996. Effects of two carbamates on infective juveniles of Steinernema carpocapsae All strain and Steinernema feltiae Ume strain. J. Nematol. 28: 310-317. Hanna, W. W., S. K. Braman, and W. G. Hudson. 2001. Bermudagrass hybrids just say no to mole crickets. Golf Course Mgmt. 69: 49-51. Hanna, W. W., and W. G. Hudson. 1997. Genetic resistance to mole crickets in turf bermudagrass. Turfgrass Trends 6: 17-18.

PAGE 93

79 Hara, A. H. and H. K. Kaya. 1982. Effect of selected insecticides and nematicides on the in vitro development of the entomogenous nematode, Neoplectana carpocapsae. J. Nematol. 14: 486-491. Hara, A. H. and H. K. Kaya. 1983a. Toxicity of selected organophosphate and carbamate pesticides to infective juveniles of the entomogenous nematode, Neoplectana carpocapsae (Rhabditida: Steinernematidae). Environ. Entomol. 12: 496-501. Hara, A. H. and H. K. Kaya. 1983b. Development of the entomogenous nematode, Neoplectana carpocapsae (Rhabditida: Steinernematidae), on insecticide-killed beet armyworm (Lepidoptera: Noctuidae). J. Econ. Entomol. 76: 423-426. Head, J., K. F. A. Walters, and S. Langton. 2000. The compatibility of the entomopathogenic nematode, Steinernema feltiae, and chemical insecticides for the control of the South American leafminer, Liriomyza huidobrensis. BioControl 45: 345-353. Hertl, P. T., R. L. Brandenburg, M. E. Barbercheck. 2001. Effect of soil moisture on ovipositional behavior in the southern mole cricket (Orthoptera: Gryllotalpidae). Environ. Entomol. 30: 466-473. Hilliard, M. A., C. Bergamasco, S. Arbucci, R. H. A. Plasterk, and P. Bazzicalupo. 2004. Worms taste bitter: ASH neurons, QUI-1, GPA-3 and ODR-3 mediate quinine avoidance in Caenorhabditis elegans. The EMBO Journal 23: 1101-1111. Hudson, W. G. 1985. Other behavior, damage, and sampling. pp. 16-21. In T. J. Walker (ed.), Mole crickets in Florida. Florida Agric. Exp. Sta. Bull. 846. Hudson, W. G. 1986. Mole crickets (Orthoptera: Gryllotalpidae) damage to Hemarthria altissima: resistance or nonpreference? J. Econ. Entomol. 79: 961-963. Hudson, W. G. 1988. Field sampling for mole crickets (Orthoptera: Gryllotalpidae: Scapteriscus): a comparison of techniques. Florida Entomol. 71: 214-216. Hudson, W. G. 1989. Field sampling and population estimation of the tawny mole cricket (Orthoptera: Gryllotalpidae). Florida Entomol. 72: 337-343. Hudson, W. G., S. K. Braman, R. D. Oetting and B. L. Sparks. 1997. Ornamental, lawn and turf insects. pp. 21-23. In D.G. Riley, G.K. Douce and R.M. McPherson (eds.). Summary of losses from insect damage and costs of control in Georgia 1996, Georgia Agric. Exp. Sta. Spec. Publ. 91. Hudson, W. G., J. H. Frank and J. L. Castner. 1988. Biological control of Scapteriscus spp. mole crickets (Orthoptera: Gryllotalpidae) in Florida. Bull. Entomol. Soc. Am. 34: 193-198.

PAGE 94

80 Hudson, W. G. and K. B. Nguyen. 1989a. Infection of Scapteriscus vicinus (Orthoptera: Gryllotalpidae) nymphs by Neoplectana Spp. (Rhabdita: Steinernematidae). Florida Entomol. 72: 383-384. Hudson, W. G. and K. B. Nguyen. 1989b. Effects of soil moisture, exposure time, nematode age, and nematode density on laboratory infection of Scapteriscus vicinus and S. acletus (Orthoptera: Gryllotalpidae) by Neoplectana sp. (Rhabditida: Steinernematidae). Environ. Entomol. 18: 719-722. Ishibashi, N., D. R. Choi, and E. Kondo. 1987. Integrated control of insects/nematodes by mixing applications of steinernematid nematodes and chemicals. J. Nematol. 19: 531. Ishibashi, N. and S. Takii. 1993. Effects of insecticides on movement, nictation, and infectivity of Steinernema carpocapsae. J. Nematol. 25: 204-213. Jansson, R. K., S. H. Lecrone and R. Gaugler. 1993. Field efficacy and persistence of entomopathenogenic nematodes (Rhabditida: Steinernematidae, Heterorhabditidae) for control of sweet potato weevil (Coleoptera: Apionidae) in southern Florida. J. Econ. Entomol. 86: 1055-1063. Jaworska, M., J. Sepiol, and P. A. D. Tomasik. 1994. Effect of metal ions on Steinernema carpocapsae nematodes. pp. 227. In A. M., R. U. Ehlers, and J. P. Masson (eds.),Genetics of entomopathogenic nematodesBacterium complexes. Burnell, European Commission, Directorate-General XII, Science Research Programme, Luxembourg, Belgium,. Kaya, H. K. 1990. Soil ecology, pp. 189-198. In R. Gaugler and H.K. Kaya (eds.), Entomopathenogenic nematodes in biological control. CRC, Boca Raton, FL. Kaya, H. K., T. M. Burlando, H. Y. Choo, and G. S. Thurston. 1995. Integration of entomopathogenic nematodes with Bacillus thuringiensis or pesticidal soap for control of insect pests. Biol. Contr. 5: 432-441. Kaya, H. K. and R. Gaugler. 1993. Entomopathenogenic nematodes. Annu. Rev. Entomol. 38: 181-206. Kaya, H. K. and S. P. Stock. 1997. Techniques in insect nematology. pp 281-324. In: Lacey, L. A, (ed.) Techniques in insect pathology. London, Academic Press. Kim, H. H., H. Y. Choo, H. K. Kaya, D. W. Lee, S. M. Lee, H. Y. Jeon. 2004. Steinernema carpocapsae (Rhabditida: Steinernematidae) as a biological control agent against the fungus gnat Bradysia agrestis (Diptera: Sciaridae) in propagation houses. Biocon. Sci. Tech. 14: 171-183. Klein, M. G. and R. Georgis. 1992. Persistence of control of Japanese beetle (Coleoptera: Scarabaeidae) larvae with steinernematid and heterorhabditid nematodes. J. Econ. Entomol. 85: 727-730.

PAGE 95

81 Koppenhfer, A. M. 2000. Nematodes. pp. 283-301. In L. A. Lacey and H. K. Kaya (eds.), Field manual of techniques in invertebrate pathology: application and evaluation of pathogens for control of insects and other invertebrate pests. Kluwer Academic, Dordrecht. Koppenhfer, A. M., I. M. Brown, R. Gaugler, P. S. Grewal, H. K. Kaya, and M. G. Klein. 2000a. Synergism of entomopathogenic nematodes and imidacloprid against white grubs: greenhouse and field evaluation. Biol. Control 19: 245-251. Koppenhfer, A. M., R. S. Cowles, E. A. Cowles, E. M. Fuzy, and L. Baumgartner. 2002. Comparison of neonicotinoid insecticides as synergists for entomopathogenic nematodes. Biol. Control 24: 90-97. Koppenhfer, A. M. and E. M. Fuzy. 2003. Effects of turfgrass endophytes (Clavicipitaceae: Ascomycetes) on white grub (Coleoptera: Scarabaeidae) control by the entomopathogenic nematode Heterorhabditis bacteriophora (Rhabditida: Heterorhabditidae). Environ. Entomol. 32: 392-396. Koppenhfer, A. M., P. A. Grewal, and H. K. Kaya. 2000b. Synergism of imidacloprid and entomopathogenic nematodes against white grubs: the mechanism. Entomol. Exp. App. 94: 283-293. Koppenhfer, A. M. and H. K. Kaya. 1998. Synergism of imidacloprid and an entomopathogenic nematode: a novel approach to white grub (Coleoptera: Scarabaeidae) control in turfgrass. J. Econ. Entomol. 91: 618-623. Krishnayya, P. V. and P. S. Grewal. 2002. Effect of neem and selected fungicides on viability and virulence of the entomopathogenic nematode Steinernema feltiae. Biocon. Sci. Tech. 12: 259-266. Lacey, L. A., R. Frutos, H. K. Kaya and P. Vail. 2001. Insect pathogens as biological control agents: do they have a future? Biol. Control 21: 230-248. Lawrence, K. O. 1982. A linear pitfall trap for mole crickets and other soil arthropods. Florida Entomol. 65: 376-377. Maeyama, T., M. Terayama and T. Matsumoto. 1994. The abnormal behavior of Colobopsis sp. (Hymenoptera: Formicidae) parasitized by Mermis (Nematoda) in Papua New Guinea. Sociobiol. 24: 115-119. Matheny, Jr., E. L. 1981. Contrasting feeding habits of pest mole cricket species. J. Econ. Entomol. 74: 444-445. Matheny, Jr., E. L., A. Tsedeke and B. J. Smittle. 1981. Feeding response of mole cricket nymphs (Orthoptera: Gryllotalpidae: Scapteriscus) to radiolabeled grasses with, and without, alternative foods available. J. Georgia Entomol. Soc. 16: 492-495.

PAGE 96

82 Milner, R. J. and J. A. Staples. 1996. Biological control of termites: results and experiences within a CSIRO project in Australia. Biocontrol. Sci. Tech. 6: 3-9. Molyneux, A. S. 1985. Survival of infective juveniles of Heterorhabditis spp., and Steinernema spp. (Nematoda: Rhabditida) at various temperatures and their subsequent infectivity for insects. Revue Nematol. 8: 165-170. Molyneux, A. S. and R. A. Bedding. 1984. Influence of soil texture and moisture on the infectivity of Heterorhabditis sp. D1 and Steinernema glaseri for larvae of the sheep blowfly, Lucilia cuprina. Nematologica 30: 359-365. Nguyen, K. B. 1988. A new nematode parasite of mole crickets: its taxonomy, biology and potential for biological control. University of Florida Ph.D. Dissertation, Gainesville, FL. Nguyen, K. B. and G. C. Smart, Jr. 1990a. Preliminary studies on survival of Steinernema scapterisci in soil. Soil and Crop Sci. Soc. Florida, Proc. 49: 230-233. Nguyen, K. B. and G. C. Smart, Jr. 1990b. Steinernema scapterisci n. sp. (Rhabditida: Steinernematidae) J. Nematol. 22:187-199. Nguyen, K. B. and G. C. Smart, Jr. 1991. Mode of entry and sites of development of Steinernema scaptersici in mole crickets. J. Nematol. 23: 267-268. Noma, T. and K. Strickler. 2000. Effects of Beauveria bassiana on Lygus hesperus (Hemiptera: Miridae) feeding and oviposition. Environ. Entomol. 29: 394-402. Parkman, J. P. and J. H. Frank. 1992. Infection of sound-trapped mole crickets, Scapteriscus spp. by Steinernema scapterisci. Florida Entomol. 75: 163-165. Parkman, J. P. and J. H. Frank. 1993. Use of a sound trap to inoculate Steinernema scapterisci (Rhabditida: Steinernematidae) into pest mole cricket populations (Orthoptera: Gryllotalpidae). Florida Entomol. 76:75-82. Parkman, J. P. and J. H. Frank. 1996. Effects of fenxoycarb on reproduction of the shortwinged mole cricket, Scapteriscus abbreviatus (Orthoptera: Gryllotalpidae). Florida Entomol. 79: 87-91. Parkman, J. P. and J. H. Frank. 1998. Development and reproduction of mole crickets (Orthoptera: Gryllotalpidae) after treatments with hydroprene and pyriproxyfen. J. Econ. Entomol. 91: 392-397. Parkman, J. P., J. H. Frank, K. B. Nguyen, and G. C. Smart. 1993a. Dispersal of Steinernema scapterisci (Rhabdita: Steinernematidae) after inoculative applications for mole cricket (Orthoptera: Gryllotalpidae) control in pastures. Biol. Control 3: 226-232.

PAGE 97

83 Parkman, J. P., J. H. Frank, K. B. Nguyen, and G. C. Smart, Jr. 1994. Innoculative release of Steinernema scapterisci (Rhabdita: Steinernematidae) to suppress pest mole crickets (Orthoptera: Gryllotalpidae) on golf courses. Environ. Entomol. 23: 1331-1337. Parkman, J. P., J. H. Frank, T. J. Walker and D. J. Schuster. 1996. Classical biological control of Scapteriscus spp. (Orthoptera: Gryllotalpidae) in Florida. Environ. Entomol. 25: 1415-1420. Parkman, J. P., W. G. Hudson, J. H. Frank, K. B. Nguyen, and G. C. Smart, Jr. 1993b. Establishment and persistence of Steinernema scapterisci (Rhabdita: Steinernematidae) in field populations of Scapteriscus spp. mole crickets (Orthoptera: Gryllotalpidae). J. Entomol. Sci. 28:182-190. Parkman, J. P. and G. C. Smart. 1996. Entomopathogenic nematodes, a case study: introduction of Steinernema scapterisci in Florida. Biocontrl. Sci. and Tech. 6: 413-419. Patterson-Stark, J. E. and L. A. Lacey. 1999. Susceptibility of western cherry fruit fly (Diptera: Tachinidae) to five species of entomopathogenic nematodes in laboratory studies. J. Invertebr. Pathol. 74: 206-208. Pendland, J. C. and D. G. Boucias. 1987. The hyphomycete Sorosporella-Syngliocladium from mole cricket, Scapteriscus vicinus. Mycopathologia 99: 25-30. Petersen, J. J. 1985. Nematode parasites. pp. 110-122. In H. C. Chapman (ed.), Biological control of mosquitoes. Am. Mosq. Control Assoc. Bull. 6. Poinar, G. O. 1979. Nematodes for biological control of insects. CRC Press, Boca Raton, FL. Poinar, G. O. 1990. Biology and taxonomy of Steinernematidae and Heterorhabditidae, pp. 23-62. In R. Gaugler and H. K. Kaya (eds.), Entomopathogenic nematodes in biological control. CRC, Boca Raton, FL. Poulin, R. 1994. Meta-analysis of parasite-induced behavioural changes. Animal Behav. 48: 137-146. Poulin, R. 1995. Adaptive changes in the behaviour of parasitized animals: a critical review. Int. J. for Parasitol. 25: 1371-1383. Poulin, R. and D. M. Latham. 2002. Parasitism and the burrowing depth of the beach hopper Talorchestia quoyana (Amphipoda: Talitridae). Animal Behav. 63: 269-275.

PAGE 98

84 Prakasa, R., P. K. Das, and G. Padhi. 1975. Note on compatibility of DD-136 (Neoplectana dutkyi), an insect parasitic nematode with some insecticides and fertilizers. Indian J. Agric. Sci. 45: 275-277. Reinert, J. A. and P. Busey. 2001. Host resistance to tawny mole cricket, Scapteriscus vicinus, in bermudagrass, Cynodon spp. Inter. Turfgrass Soc. Res. Journal 9: 793-797. Rovesti, L. and K. V. Dese. 1990. Compatibility of chemical pesticides with the entomopathogenic nematodes, Steinernema carpocapsae Weiser and S. feltiae Filipjev (Nematoda: Steinernematidae). Nematologica 36: 237-245. SAS Institute. 2001. JMP Start Statisitics Users Guide. Second ed., SAS Inst. Inc., Cary, NC. Schroeder, P. C., M. G. Villani, C. S. Ferguson, J. P. Nyrop and E. J. Shields. 1993. Behavioral interactions between Japanese beetle (Coleoptera: Scarabaeidae) grubs and an entomopathogenic nematode (Nematoda: Heterorhabditidae) within turf microcosms. Environ. Entomol. 22: 595-600. Schuster, D. J. and J. F. Price. 1992. Seedling feeding damage and preference of Scapteriscus spp. mole crickets (Orthoptera: Gryllotalpidae) associated with horticultural crops in west-central Florida. Florida Entomol. 75: 115-119. Shapiro, D. I. and E. E. Lewis. 1999. Comparison of entomopathogenic nematodes infectivity from infected hosts versus aqueous suspension. Environ. Entomol. 28: 907-911. Shapiro-Ilan, D.I., 2001. Virulence of entomopathogenic nematodes to pecan weevil larvae, Curculio caryae (Coleoptera: Curculionidae), in the laboratory. J. Econ. Entomol. 94: 7-13. Short, D. E., and P. G. Koehler. 1979. A sampling technique for mole crickets and other pests in turfgrass and pasture. Florida Entomol. 62: 282-283. Sithole, C. E. 1986. Mole cricket (Gryllotalpa africana). Zimbabwe Agric. J. 83: 21-22. Smith, K. 1999. Factors affecting efficacy, pp. 37-46. In S. Polavarapu (ed.), Proceedings, Optimal Use of Insecticidal Nematodes in Pest Management, 28-30 August 1999, New Brunswick, NJ. Spinney, L. 1995. High-scoring crickets return for a second inning. New Scientist 146: 6. Steel, R. G. D. and J. H. Torrie. 1980. Principles and procedures of statistics: a biometrical approach. 2 nd ed. McGraw-Hill, New York.

PAGE 99

85 Thompson, S. R. 2004. Tunneling responses of mole crickets (Orthoptera: Gryllotalpidae) to the entomopathogenic fungus, Beauveria bassiana (Balsamo) Vuillemin. Masters Thesis, North Carolina State University. Raleigh, NC. Thurston, G. S., W. N. Yule, and G. B. Dunphy. 1994. Explanations for the low susceptibility of Leptinotarsa decemlineata to Steinernema carpocapsae. Biol. Control 4: 53-58. Ulagaraj, S. M. 1974. Mole crickets: ecology, behavior, and acoustical communication (Orthoptera: Gryllotalpidae: Scapteriscus) Ph.D. Dissertation, Univ. Florida, Gainesville. 72 pp. Ulagaraj, S. M. 1975. Mole crickets: ecology, behavior, and dispersal flight (Orthoptera: Gryllotalpidae: Scapteriscus). J. Econ. Entomol. 4: 265-273. Vance, S. A. 1996. The effect of the mermithid parasite Gasteromermis sp. (Nematoda: Mermithidae) on the drift behavior of its mayfly host, Baetis bicaudatus (Ephemeroptera: Baetidae): a trade-off between avoiding predators and locating food. Can. J. Zool. 74: 1907-1913. Vaughn, C.C., S.M. Glenn and I. Butler. 1993. Characterization of prairie mole cricket chorusing sites in Oklahoma. Am. Midl. Nat. 130: 364-71. Villani, M. G., L. L. Allee, A. Diaz, and P. S. Robbins. 1999. Adaptive strategies of edaphic arthropods. Annu. Rev. Entomol. 44: 233-256. Villani, M. G., L. L. Allee, L. Preston-Wilsey, N. Consolie, Y. Xia, and R. L. Brandenburg. 2002. Use of radiography and tunnel castings for observing mole cricket (Orthoptera: Gryllotalpidae) behavior in soil. Amer. Entomol. 48: 42-50. Villani, M. G., S. R. Krueger, P. C. Schroeder, F. Consolie, and N. H. Consolie. 1994. Soil application effects of Metarhizium anisopliae on Japanese beetle (Coleoptera: Scarabaeidae) behavior and survival in turfgrass microcosms. Environ. Entomol. 23: 502-513. Walker, S. L. 1979. Population estimation, damage evaluation and behavioral studies on mole crickets Scapteriscus vicinus and S. acletus (Orthoptera: Gryllotalpidae). Masters Thesis, University of Florida, Gainesville. Walker, T. J. 1985. Systematics and life cycles. pp 3-10. In T.J. Walker (ed.) Mole Crickets in Florida. Florida Agric. Exp. Sta. Bull. 846. Walker, T. J. and N. Dong. 1982. Mole crickets and pasture grasses: Damage by Scapteriscus vicinus, but not S. acletus (Orthoptera: Gryllotalpidae). Fla. Entomol. 65: 300-306. Walker, T. J., R. C. Littell, and N. Dong. 1982. Which mole crickets damage bahiagrass pastures? Florida Entomol. 65: 110-116.

PAGE 100

86 Walker, T. J. and D. A. Nickle. 1981. Introduction and spread of pest mole crickets: Scapteriscus vinicus and S. acletus reexamined. Ann. Entomol. Soc. Am. 74: 158-163. Walker, T. J., J. P. Parkman, J. H. Frank and D. J. Schuster. 1996. Seasonality of Ormia depleta and limits to its spread. Biol. Control 6: 378-383. Wang, C., J. E. Powell, and K. B. Nguyen. 2002. Laboratory evaluations of four entomopathogenic nematodes for control of subterranean termites (Isoptera: Rhinotermitidae). Environ. Entomol. 31: 381-387. Williams, J. J. and L. N. Shaw. 1982. A soil corer for sampling mole crickets. Florida Entomol. 65: 192-194. Williams, J. K., C. R. Townsend and R. Poulin. 2001. Mermithid nematode infections and drift in the mayfly Deleatidium spp. (Ephemeroptera). J. Parasitol. 87: 1225-1227. Zhou, X., H. K. Kaya, K. Heungens, and H. Goodrich-Blair. 2002. Response of ants to a deterrent factor(s) produced by the symbiotic bacteria of entomopathogenic nematodes. Appl. Environ. Microbiol. 68: 6202-6209. Zimmerman, R. J. and W. S. Cranshaw. 1990. Compatibility of three entomogenous nematodes (Rhabditida) in aqueous solutions of pesticides used in turfgrass maintenance. J. Econ. Entomol. 83: 97-100.

PAGE 101

BIOGRAPHICAL SKETCH Kathryn Ann Barbara was born on 28 April 1975 in Pittsfield, Massachusetts. Upon graduation from St. Josephs Central High School in Pittsfield, Massachusetts, she entered the University of Vermont in Burlington, Vermont, graduating in 1997 with a Bachelor of Science degree in biology. In the two years following her graduation, Kathryn worked as a laboratory technician at Chiron Diagnostic Corporation in Walpole, Massachusetts, and at the prestigious Brigham and Women's Hospital. In the fall of 1999, she moved from Boston, Massachusetts, to begin her graduate studies in entomology at the University of Florida under the guidance of Dr. John Capinera. Kathryn received her Master of Science degree from the University of Florida in December 2001. In January 2002 she began her doctoral research under the tutelage of Dr. Eileen Buss. Kathryn joined the United States Navy under the Health Services Collegiate Scholarship Program in October 2002. Upon graduation in April 2005 Kathryn will be commissioned as a lieutenant in the U.S. Navy and serve in the medical service corps as a naval entomologist. 87


Permanent Link: http://ufdc.ufl.edu/UFE0009590/00001

Material Information

Title: Management of Pest Mole Crickets Using the Insect Parasitic Nematode Steinernema scapterisci
Physical Description: Mixed Material
Copyright Date: 2008

Record Information

Source Institution: University of Florida
Holding Location: University of Florida
Rights Management: All rights reserved by the source institution and holding location.
System ID: UFE0009590:00001

Permanent Link: http://ufdc.ufl.edu/UFE0009590/00001

Material Information

Title: Management of Pest Mole Crickets Using the Insect Parasitic Nematode Steinernema scapterisci
Physical Description: Mixed Material
Copyright Date: 2008

Record Information

Source Institution: University of Florida
Holding Location: University of Florida
Rights Management: All rights reserved by the source institution and holding location.
System ID: UFE0009590:00001


This item has the following downloads:


Full Text












MANAGEMENT OF PEST MOLE CRICKETS USING THE INSECT PARASITIC
NEMATODE Steinernema scapterisci














By

KATHRYN ANN BARBARA


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

UNIVERSITY OF FLORIDA


2005

































Copyright 2005

by

Kathryn Ann Barbara

































This dissertation is dedicated to my family, Kathleen and Jack Barbara, Anthony, Lara,
and Elizabeth Barbara, Lisa, Eric and Ryan Malkowski, Anne Barbara, Ann and
Raymond Callahan, John Michael, Kathleen, Melinda, and Maureen Gowdey, Camilla
Barbara, James Dunford, and Magnum, the delightful Doberman. Without their love,
thoughtfulness and support this would not have been possible.















ACKNOWLEDGMENTS

My heartfelt thanks go to my major advisor, Dr. Eileen Buss. Her guidance,

advice, and trust have made this research possible. Dr. Buss has been a mentor in not

only my academic life but my personal life as well. I consider myself lucky to have been

her first Ph. D. student. I would also like to thank my committee members, Drs. J.

Howard Frank, Norman C. Leppla, Grady L. Miller, and especially Khuong B. Nguyen,

who has given me the knowledge about nematology to make my degree truly a work of

entomology and nematology. I could not have had a better committee. Each has

contributed significantly to this work and it would not have been possible without each

and every member.

This research would not have been possible without the help of many people. I

would like to thank the cooperators and their assistants who allowed me to use their sites

and were very helpful during my research. Special thanks go to Buddy Keene II, Lloyd J.

Brown and Diane Delzell from Gainesville Golf and Country Club; Dave Carson, Kevin

Cooke, and Vern Easter from Ironwood Golf Course; Fred Santana from Sarasota County

Extension; and Matt Burke from the City of Altamonte Springs. Nematodes and advice

on their use were provided by Tom Hinks, Gabe Diaz-Saavedra, and Al Clarke from

Becker Underwood and Martin Adjei of the Ona Range Cattle REC. I acknowledge Dr.

Tom Walker for mole cricket advice and assistance.

I acknowledge Dr. Albrecht Koppenhofer from Rutgers University for technical

advice and methodology for the pesticide compatibility tests. Special thanks go to John









Fredricks for help with soil sample analysis, Marinela Capanu for help with statistical

design and analysis, and Paul Skelley (DPI) for help with SEM photographs. I

acknowledge Bayer Environmental Science, Valent Professional Products, and FMC

Corporation for donating insecticides used in this study. Special thanks go to Brian

Owens at the G. C. Horn Memorial Turfgrass Field Laboratory for his assistance and

patience with setting up nightly collections with mole cricket sound callers. Angela

Vincent and Shubin Saha assisted in sorting linear pitfall trap samples. I would also like

to acknowledge the electronic thesis and dissertation technical staff for their help in

formatting and submission of my dissertation. Many thanks go to the Florida Department

of Agriculture and the United States Navy for graduate funding.

Very special thanks go to the numerous people who helped in my various projects;

without them this project would not have been possible. First I would like to thank Paul

Ruppert for his assistance in numerous aspects of this study and his patience with me. I

also would like to thank Bob Hemenway for rearing supplies, guidance and advice. My

gratitude goes to Matthew Stanton and Y. Mike Wang for help with bi-weekly pitfall trap

collections and mole cricket rearing. I thank J. Cara Congdon, Jay Cee Turner, Lois

Wood, Rebecca Baldwin, Erin Finn, Alejandro Arevalo, Justin Emerson and Brian

McElroy for assistance in pitfall trap installation and removal. Special thanks go to

Robert Mans and Rachel Davis for help with mole cricket rearing and the dreaded curfew

field test. Many thanks go to the University of Florida Entomology and Nematology

Department for their support and friendship through the years.

Several people I have met along this journey have become more than just my

academic and professional peers but have become very good friends. I want to give









special thanks and warm gratitude to J. Cara Congdon and Jay Cee Turner who, while

helping me become a better scientist by looking towards me for guidance and advice,

have become two of my best friends. I am especially grateful to James Dunford. He has

inspired me in all aspects of my life and several ideas in this dissertation were inspired

and motivated by him. Jim has given me the opportunity to discover other aspects of

entomology outside of my field. My only regret is that I did not meet him sooner.
















TABLE OF CONTENTS



A C K N O W L E D G M E N T S ................................................................................................. iv

L IST O F T A B L E S .............. ................................................... ............... x.. ... ......x

LIST OF FIGURES .. ................... ............ ........ .............. xii

A B S T R A C T .................................................................................................................... x iii

CHAPTER

1 INTRODUCTION AND REVIEW OF LITERATURE ................... ..................... 1

M o le C ric k e ts ................................................................................................................ 1
M anagem ent Practices .................................................. .............. ...............4...
C h em ical C control ... .. ........................................... ........................ .... ......... ... 4
N on-C hem ical C control ................. .............................................................5......
B biological C control ........................................................................................ .. .5
Insect Parasitic N em atodes................................... ...................... ...............6......
O b je ctiv e s ............................................................................................................... .. 1 0

2 ESTABLISHMENT AND SPREAD OF Steinernema scapterisci ON FLORIDA
G O L F C O U R S E S ....................................................................................................... 12

M materials and M ethods .. ..................................................................... ............... 13
S tu d y S ite s ........................................................................................................... 1 3
M ole C ricket M monitoring ....................................... ...................... ............... 13
L ab oratory A ssay ... ... ......................................... ....................... .......... 15
Statistical A analysis .............. .................. .............................................. 15
R results and D discussion ............................ .......................................... 15

3 SURVIVAL AND INFECTIVITY OF Steinernema scapterisci AFTER CONTACT
WITH SOIL DRENCH SOLUTIONS ..................................................................25

M materials and M ethods ............................................ ........................... ................ 27
N em atodes and M ole C rickets........................................................ ................ 27
B io a ssa y .......................................................................................................... 2 7
Statistical A naly sis .............. ...... ............ .............................................. 30
R results and D discussion ............................ .......................................... 30









4 INTEGRATION OF INSECT PARASITIC NEMATODES WITH INSECTICIDES
FOR CONTROL OF PEST MOLE CRICKETS ............................. ..................... 36

M materials an d M eth o d s ............................. ......... ....... .......... ........................................3 7
Survival and Infectivity of S. scapterisci After Exposure to Pesticides..............38
Nematode Infectivity After Exposure to Pesticide Treated Mole Crickets .........39
Statistical A analysis .............. .... .............. ................................................ 40
R results and D discussion ................ .............. ............................................ 40

5 EFFECT OF Steinernema scapterisci NGUYEN AND SMART EXPOSURE ON
MOLE CRICKET TUNNELING, OVIPOSITION, AND AVOIDANCE
B E H A V IO R ................................................................................................................ 4 6

M materials an d M eth o d s ......................................................................... .... ................ 4 7
Nematode Infection and Nematode Treated Areas Effect on Mole Cricket
Tunneling B behavior ... ...... ............. ...... ................ ... ............... 48
Oviposition Behavior of Mole Crickets Exposed to S. scapterisci ..................50
Y -T u b e T e sts ....................................................................................................... 5 0
O b servation C ham b er................................................................. ............... 50
Response to S. scapterisci or Pesticides.................................. ................ 51
Statistical A analysis .............. .................. .............................................. 52
R e su lts ............................ ................................................................ ...... ..................... 5 2
Nematode Infection and Nematode Treated Areas Effect on Mole Cricket
T unneling B ehavior98 ........................................................... ... ................ 52
Oviposition Behavior of Mole Crickets Exposed to S. scapterisci.................. 53
Y -T u b e T e sts ....................................................................................................... 5 3
D isc u ssio n ............................................................................................................... ... 5 4

6 SUM M ARY AND CONCLU SION S.................................................... ................ 61

APPENDIX

A AMBIENT DATA AND TURFGRASS QUALITY DATA COLLECTED AT
GAINESVILLE GOLF AND COUNTRY CLUB AND IRONWOOD GOLF
C O U R S E ..................................................................................................................... 6 3

B DATA FROM ATHLETIC FIELD DEMONSTRATION SITES.............................66

M materials an d M eth o d s ...............................................................................................6 6
O objective ............................................................................. .......... ............... 66
S tu d y S ite ............................................................................................................. 6 6
M ole C ricket M monitoring ....................................... ...................... ................ 66
R results and D discussion ................ .............. ............................................ 68

C PRELIMINARY CHECKLIST OF ARTHROPODS ASSOCIATED WITH GOLF
C O U R SE T U R F G R A S S ............................................................................................7 1









D SCANNING ELECTRON MICROGRAPH PICTURES OF MOLE CRICKET
SE N S O R Y A R E A S ....................................................................................................7 3

L IST O F R E F E R E N C E S ...................................................................................................76

BIO GR APH ICAL SK ETCH .................................................................... ................ 87















LIST OF TABLES


Table page

2-1. Percent infection of Scapteriscus spp. mole crickets collected from sites treated with
Steinernem a scap terisci .......................................... ......................... ................ 24

3-1. Mean nematode mortality and percent of mole crickets infected with Steinernema
scapterisci after exposure for 24 h to various drenching solutions.......................34

3-2. Mean nematode mortality and infectivity after exposure for 24 h to various
drenching solutions. ............................ ............................................ 34

3-3. Mean number of mole crickets emerging from bermudagrass using various
drenching solutions in M ay 2003 ....................................................... ................ 35

3-4. Percent nematode infection from mole crickets exposed to treatment solutions 1, 5,
8, 12, or 24 h post infection ................................... ....................... ................ 35

4-1. Percent survival, infectivity, and days until death of S. scapterisci incubated in
solutions of insecticides for 24 h ......................................................... ................ 44

4-2. Average days until death and percent infectivity by S. scapterisci nematodes to mole
crickets exposed to insecticides........................................................... ................ 45

5-1. Oviposition of Scapteriscus mole crickets directly infected with different numbers
o f S scap terisci. ....................................................................................................... 5 9

5-2. Oviposition of Scapteriscus mole crickets in sand treated with S. scapterisci.......... 59

5-3. Response of Scapteriscus vicinus, S. borellii, and S. abbreviatus to Steinernema
scapterisci nem atodes versus sterilized sand. ..................................... ................ 59

5-4. Response of Scapteriscus vicinus to Steinernema scapterisci nematodes versus
pesticides treated sand .. ................................................. ............ 60

A-1. Ambient data collected from Gainesville Golf and Country Club on dates of mole
crickets collections. ........... ... .............. .... ........ ...... ............... 63

A-2. Ambient data collected from Ironwood Golf Course on dates of mole crickets
c o lle ctio n s ............................................................................................................ .. 6 4









A-3. Average turfgrass density ratings for treated and untreated plots on Gainesville Golf
and Country Club and Ironwood Golf Course. ................................... ................ 65















LIST OF FIGURES


Figure page

2-1. Linear pitfall trap used to collect m ole crickets. .................................. ................ 21

2-2. 3.8 L catch bucket inside 19 L bucket of linear pitfall trap..................................21

2-3. Mean monthly percent infection of mole crickets collected in pitfall traps at
Ironwood Golf Course from areas treated with Steinernema scapterisci ............22

2-4. Mean monthly percent infection of mole crickets collected in pitfall traps at
Gainesville Golf and Country Club from areas treated with Steinernema
sc ap te risc i................................................................................................................ 2 3

5-1. Distance tunneled through sand in Plexiglas arenas every 2 d by mole crickets
exposed to varying am ounts of nem atodes. ........................................ ................ 57

5-2. Comparison of total tunnel distances by male and female Scapteriscus spp. mole
crickets after exposure to S. scapterisci .............................................. ............... 58

5-3. Distance tunneled by mole crickets through sand treated with nematodes and
untreated sand in Plexiglas arenas over 48 h....................................... ................ 58

B-1. Average monthly infection rates at the Sarasota athletic field research site .......... 69

B-2. Average monthly infection rates at the Altamonte Springs athletic field research
site ........................................................................................................ ........ .. 6 6

D-1. SEM photograph of a female Scapteriscus vicinus antennal mid-section.............73

D-2. SEM photograph of a male Scapteriscus vicinus antennal mid-section...................74

D-3. SEM photograph of a female Scapteriscus vicinus mid-tarsal claw.........................74

D-4. SEM photograph of a female Scapteriscus vicinus labial palpomere.......................75















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

MANAGEMENT OF PEST MOLE CRICKETS USING THE INSECT PARASITIC
NEMATODE Steinernema scapterisci

By

Kathryn Ann Barbara

May 2005

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

Steinernema scapterisci Nguyen and Smart nematodes became established on two

golf courses in Gainesville, FL, when applied as an augmentative application, and moved

to untreated areas. It took about 4-8 wk post application for infection of mole crickets

(Scapteriscus spp.) to equal or exceed pretreatment levels. Infection levels in untreated

areas at least 80 m away from treated areas reached infection levels similar to the treated

areas in approximately 20 wk post application.

After a nematode application, mole crickets are frequently assayed to confirm

nematode establishment. However, the standard soap flush was suspected of providing

false negatives under field conditions. Thus, we examined the effect of several potential

flushing solutions on the survival and infectivity of S. scapterisci as well as flushing

ability under field conditions. Seventy percent of S. scapterisci died in the lemon dish

detergent solution, confirming that assays for nematode infection of soap-flushed mole

crickets are likely to be inaccurate. When sampling for mole crickets in areas where S.









scapterisci has been applied, a potential alternative to the standard soap drench is a dilute

permethrin drench.

Aqueous solutions of pesticides (acephate, bifenthrin, deltamethrin, fipronil and

imidacloprid) used in turfgrass to control mole crickets were tested for compatibility with

S. scapterisci in the laboratory. Survival of S. scapterisci was >95% in solutions of

acephate, bifenthrin and imidacloprid. Infectivity of S. scapterisci previously exposed to

insecticides was >60% in acephate and bifenthrin; however, infectivity was <40% in

imidacloprid. The entomopathogenic nematode was compatible with all insecticides

tested.

Both healthy and nematode-infected mole crickets had similar tunneling behavior.

Although not significant, crickets treated with 500 or 10,000 nematodes tunneled less

than uninfected crickets. Mole crickets did not appear to differentiate between untreated

and nematode-treated sand. Female crickets infected with nematodes were able to lay

eggs, and clutch size and egg chamber size were not significantly different than healthy

crickets. Crickets also laid eggs in sand treated with nematodes, suggesting that the

nematode treated sand was not a deterrent. Mole crickets in Y-tube tests did not

significantly choose untreated sand over sand treated with 500 or 10,000 nematodes.

When given a choice between field rates of S. scapterisci and acephate, bifenthrin,

deltamethrin, fipronil, or imidacloprid, crickets significantly chose nematodes over

insecticides.














CHAPTER 1
INTRODUCTION AND REVIEW OF LITERATURE

Mole Crickets

Exotic mole crickets (Scapteriscus spp.) are the most injurious insect pests of golf

courses, lawns, sod farms and pastures in Florida and throughout the southeastern United

States (Walker and Nickle 1981). Mole crickets damage turf by tunneling in the soil

which exposes and dries out roots and by direct root feeding. As a result, the turfgrass

thins and bare patches appear. Weeds may invade these patches, leading to increased

herbicide use. The tunneling and mounds that mole crickets make also disrupt the

playing surface on golf courses, especially the roll of the golf ball on greens.

Superintendents and golf course members typically have zero tolerance for damage

(Frank and Parkman 1999). Insecticide treatments are usually targeted against the most

destructive nymphal stages (Parkman and Frank 1996, 1998). Mole cricket damage and

cost of control in Florida in 1986 were estimated at $45 million with an additional $33

million in Alabama, Georgia, and South Carolina combined (Frank and Parkman 1999).

Estimates of annual expenditure on chemical insecticides are over $18 million in Florida

turf, and over $12 million in control costs (Hudson et al. 1997).

Mole crickets are omnivorous, feeding on animal as well as plant material. Studies

have indicated that the southern mole cricket, Scapteriscus borellii Giglio-Tos, is less

damaging than the tawny mole cricket, Scapteriscus vicinus Scudder. Scapteriscus

borellii is a predator and feeds mostly on other insects while S. vicinus is mainly

herbivorous (Matheny 1981, Matheny et al. 1981, Walker and Dong 1982). Dissection of









field trapped S. borellii showed that their gut contents contained 66% animal material and

only 2% of plant material, and the rest a combination of plant and animal material. In

contrast, 84% of the gut contents of the short-winged mole cricket, Scapteriscus

abbreviatus Scudder, and 88% of S. vicinus contained plant material (Hudson 1985).

Both S. vicinus and S. borellii are pests of tomato and strawberry fields in Florida

(Schuster and Price 1992), as well as many vegetables, peanut, sugarcane, tobacco, and

ornamentals such as coleus, chrysanthemum, and gypsophila. Among turfgrasses, S.

vicinus often injures bahiagrass and bermudagrass, whereas S. abbreviatus favors St.

Augustinegrass and bermudagrass. Mole crickets also feed on weeds such as pigweed

and Amaranthus spp. (Capinera and Leppla 2001).

There are ten mole cricket species in the continental United States, Hawaii, Puerto

Rico, and the Virgin Islands (Frank et al. 1998). All mole cricket species are not pests.

Most species are innocuous and some are rare. For example, in Britain Gryllotalpa

gryllotalpa (L.) has become so rare that efforts to reintroduce it to mainland England

have been proposed (Spinney 1995). In the United States the prairie mole cricket,

Gryllotalpa major Saussure, feeds on prairie vegetation and is restricted to four central

states due to habitat loss, though it once had a much wider distribution (Vaughn et al.

1993, Frank et al. 1998).

Mole crickets primarily live underground in excavated tunnels. The forelegs of the

mole cricket are flattened and expanded, enabling it to burrow quickly in sandy and

extremely dry soils. Mole crickets form vertical tunnels as well as horizontal galleries

just below the soil surface. Mole crickets usually occur in the top 20-25 cm of soil and

have been recorded to tunnel as deep as 75 cm (Hudson 1985). The depth of mole cricket









tunneling varies with temperature and moisture. Galleries made by the crickets can be

used as an indication of their presence in the turf. Adults and large nymphs occasionally

move about on the soil surface on warm nights with high humidity and are often attracted

to lights.

Male mole crickets produce loud songs (50-90 dB) after sunset by rasping a

stridulatory file on the forewing. Males use a funnel-shaped opening at the mouth of a

subterranean calling chamber, which amplifies the sound of their song. Calling chambers

are constructed each evening 10-20 min. before calling. The male mole cricket then

tunes these chambers to the frequency of his song (Forrest 1985), and calling lasts for

approximately 1 h after sunset. Songs function to attract flying and walking females.

The male's song intensity varies depending on male size and soil moisture (Forrest 1985).

The following information on the mole cricket life cycle was determined by Walker

(1985) and Frank and Parkman (1999). Adult S. vicinus fly in large numbers in early

spring, typically in March, but as early as February in Florida after warm winters. In

early summer, mole crickets mate (although some mating occurs the previous autumn),

and oviposit one or more clutches of 25-60 eggs. Females mate with males for < 24 h,

lay one or more egg clutches within 10-14 d after mating and then die. Eggs mature

within 3 wk in an incubation chamber in the soil. Nymphs hatch from eggs as early as

April, but continue to hatch from later deposited eggs through June. Nymphs develop for

about 5 mo and adults begin to appear in September. In some years there is a minor peak

of flight activity in the autumn, as early as August in the far south and as late as

December farther north if the weather remains warm. In most of the southern USA, the

spring activities occur in S. borellii about 3 wk later than in S. vicinus, but the autumnal









activities are concurrent. In south Florida, S. borellii has two generations during the

summer months, with a second peak in adult flight in July, following the first peak in

April. More S. borellii than S. vicinus overwinter as large nymphs. In contrast, all

developmental stages of S. abbreviatus occur throughout the year but with two peak

ovipositional periods, one in late spring and one in winter. Scapteriscus abbreviatus has

short, non-functional wings, cannot fly, and does not produce calling sounds. The peak

periods of damage caused by feeding of any species are when nymphs are abundant,

developing rapidly, and ingesting much food.

Management Practices

Chemical Control

Turf damage mitigation is often the top priority of turfgrass managers and this is

usually achieved by applying pesticides. Insecticide classes used to manage mole cricket

populations include carbamates, organophosphates, phenyl pyrazoles and pyrethroids.

Insecticides commonly applied to control mole crickets in lawns and golf courses are

expensive and not always effective while in pastures there is no economically feasible

control (Walker 1985). Several risks are associated with insecticide use for mole cricket

control. For example, insecticides often have a short residual and treated areas are

subject to reinfestation. Insecticides are usually non-specific and therefore non-target

insects, including natural enemies, are killed by applications. Insecticide-treated mole

crickets may die on the soil surface, which attracts birds and other insectivores and risks

secondary mortality of these organisms. Improper applications may contaminate

groundwater by runoff or seepage (Frank and Parkman 1999). Finally, insecticide use on

golf courses and athletic fields requires that play be suspended for the legally required









no-entry time after treatment to minimize human exposure. Scapteriscus mole crickets

are therefore good targets for classical biological control (Frank 1990).

Non-Chemical Control

Other options for non-chemical control of mole crickets exist (e.g., cultural,

physical or mechanical controls), but are not readily applicable to turfgrass and pastures

of the southeastern USA. Examples include tillage at appropriate times of year to expose

eggs and small nymphs to desiccation and flooding (Denisenko 1986, Sithole 1986).

Reducing mowing height acts as a mechanical control by killing any mole crickets on the

soil surface.

Biological Control

There are several natural enemies of precinctive and exotic pest mole crickets in the

USA. Birds, mammals, amphibians, reptiles and arthropods attack mole crickets. They

include birds, raccoons and armadillos, toads, snakes, carabid beetles and earwigs.

Several fungal pathogens have been isolated from mole crickets in Florida, including

Aspergillus, Beauveria, Isaria, Metarhizium, and Sorosporella (Boucias 1985, Pendland

and Boucias 1987). Several arthropods are also used as biological control agents of

Scapteriscus mole crickets, including the phonotactic tachinid fly Ormia depleta

(Wiedemann), the neotropical digger wasp Larra bicolor F., Megacephala tiger beetles,

Pasimachus carabid beetles, and wolf spiders (Hudson et al. 1988, Parkman et al. 1996).

Of these, 0. depleta and L. bicolor have been released in Florida.

In 1988, a Brazilian strain of 0. depleta was released in Florida to parasitize exotic

mole crickets. The female fly locates Scapteriscus spp. by mating songs and deposits

larvae on or near the host. The fly's seasonality limits its spread so it has been more

successful in central and south Florida. Golf course superintendents have reported that









counties in Florida with 0. depleta populations had significantly less mole cricket

damage than did counties that lacked 0. depleta (Frank et al. 1996). Ormia depleta

seems to overwinter more successfully in central Florida than in northern Florida,

perhaps because of milder winters in central locations (Walker et al. 1996).

The neotropical digger wasp L. bicolor parasitizes large nymphs and adults of mole

crickets (Frank and Parkman 1999). The wasp attacks and stings mole crickets on the

soil surface, causing paralysis. The wasp deposits an egg on the mole cricket near the

pronotum, and the neonate larva develops as an external parasitoid. The parasitized mole

cricket resumes normal activities and dies after a few weeks. Scapteriscus mole crickets

are the only known host ofL. bicolor (Frank et al. 1995). However, this wasp also feeds

on wildflower nectar which can be used to maintain or increase populations in an area.

Insect Parasitic Nematodes

Insect parasitic nematodes have recently been investigated for use against

subterranean and soil inhabiting pests (Kaya and Gaugler 1993). Insect parasitic

nematodes in the genera Steinernema and Heterorhabditis are potent biological control

agents that generally infect their hosts by entering natural openings such as the mouth,

spiracles, and anus (Shapiro and Lewis 1999). Insect parasitic nematodes have been used

to control mole crickets and other insect pests such as lepidopteran larvae (Epsky and

Capinera 1994, Shapiro and Lewis 1999), banded cucumber beetle (Creighton and

Fassuliotis 1985), sweet potato weevil (Jansson et al. 1993), pecan weevil larvae

(Shipiro-Ilan 2001), western cherry fruit fly (Patterson-Stark and Lacey 1999), Japanese

beetle grubs (Klein and Georgis 1992, Schroeder et al. 1993), and various Homopterans

(English-Loeb et al. 1999). The nematodes in these genera are mutualistically associated

with bacteria (Xenorhabdus spp. for steinernematids and Photorhabdus spp. for









heterorhabditids). Infective juvenile nematodes enter the host through an opening in the

arthropod. Once in the hemocoel, they release their symbiotic bacteria, which kill the

host and provide nematodes with nutrients and defense against secondary invaders

(Poinar 1990). The nematodes complete two to three generations within the arthropod

host, and then infective juveniles are released to search out new hosts (Kaya and Gaugler

1993). Several steinermatid and heterorhabditid nematodes are in current use, or are

being considered for use, as commercial biopesticides against soil insects in many

agricultural and horticultural systems (Gaugler and Kaya 1990, Kaya 1990, Georgis and

Gaugler 1991, Kaya and Gaugler 1993). Disadvantages in using insect parasitic

nematodes include sensitivity to ultraviolet light, desiccation, and insect/pathogen

interactions (Kaya and Gaugler 1993, English-Loeb et al. 1999). Certain nematode

species are highly effective against a particular pest, whereas others may be ineffective or

moderately effective against the same pest (Shapiro-Ilan 2001). The effectiveness of

entomopathogenic nematodes depends on matching the target pest species with the most

effective nematode.

The entomopathogenic nematode Steinernema scapterisci Nguyen and Smart was

collected in Uruguay in pitfall-trapped Scapteriscus mole crickets in the 1980s. The

nematode was cultured and then released in Florida pastures in 1985, established

populations and was spread from the release site by infected Scapteriscus mole crickets

(Hudson et al. 1988, Nguyen and Smart 1990a, Parkman and Frank 1992, Frank 2001).

The nematode successfully kills adults and large nymphs of S. borellii and S. vicinus and

to a lesser extent S. abbreviatus. Small to medium nymphs of S. borellii and S. vicinus

are less frequently infected (Hudson and Nguyen 1989a).









Steinernema scapterisci live in moist soil and can survive without a host for at least

10 wk (Nguyen and Smart 1990a). Third stage infective juveniles can migrate up and

down 10 cm in the soil and invade mole cricket bodies through the mouth or spiracles

(Nguyen and Smart 1991). Once inside the mole cricket, the nematodes release bacteria

that feed on the hemolymph killing the mole cricket through septicemia; the nematodes

then eat the bacteria and reproduce inside the mole cricket resulting in mole cricket

mortality within 2-3 d. The nematodes then exit the body and infect other mole crickets

in the soil. Mole crickets can fly up to 8 km before dying (Walker 1985), thus spreading

nematodes to new sites. The use of S. scapterisci as a biopesticide would allow treatment

to hot spots rather than doing a broadcast application of synthetic insecticides. Anecdotal

evidence indicates that since the original release of the nematodes (ca. 1985), overall

mole cricket populations in Florida have declined (J.H. Frank, personal communication).

Augmentative nematode releases are likely to further reduce mole cricket populations.

Over the past 20 years, researchers have become very interested in changes in

behavior displayed by parasitized animals and whether or not they represent parasite or

host adaptations (Poulin 1995). Changes in behavior displayed by parasitized organisms

vary from slight shifts in the time spent performing a given activity to appearance of

drastically new and strange behaviors (Poulin 1994, Benton and Pritchard 1990).

Oftentimes these parasite induced behavioral modifications are simply pathological side

effects of parasite infection (Poulin 1995, Vance 1996, Williams et al. 2001). Studies

with the thrips Frankliniella occidentalis (Pergande) and the entomopathogenic nematode

Thripinema nicklewoodi Siddiqi demonstrated behavioral change in the host insect. The

behavioral changes displayed were reduced feeding and reduced fecundity ofF.









occidentalis when infected with T. nicklewoodi (Arthurs and Heinz 2003). The results

from this study proved useful in the biological control of thrips because T nicklewoodi

reduce populations by parasitizing F. occidentalis, reducing direct feeding damage, and

reducing the spread of tomato spotted wilt virus.

Nematodes also modify their host's behavior to increase their own fitness.

Parasitism of the beach hopper Talorchestia quoyana Milne-Edwards by mermithid

nematodes results in a greater burrowing depth of the host. Adult mermithids live in

water or moist soil; however their beach hopper host prefers dry terrestrial microhabitats.

This behavior modification of the host allows the nematode to mate and lay eggs in more

humid environments deeper in the soil (Poulin and Latham 2002). Maeyama et al. (1994)

observed that the ant Colobopsis sp. infected with Mermis nematodes displayed a suicidal

behavior by jumping into water and dying. This behavior is advantageous to the

nematode because they require water to reproduce. In order to increase contact with a

mate, the nematodes must emerge from the ants in water.

More than 30 species of nematodes are associated with insects and other

invertebrates (Poinar 1979, 1990; Kaya and Stock 1997; Lacey et al. 2001; Koppenhofer

and Fuzy 2003). The nematode families Allantonematidae, Heterorhabditidae,

Mermithidae, Tetradonematidae, Sphaerulariidae, and Steinernematidae are the focus of

much research because of their potential as biological control agents of insects (Lacey et

al. 2001). For example, inoculative releases of Deladenus siricidicola Bedding in New

Zealand and Australia has been a successful classical biological control agent of the

woodwasp Sirex noctilio F. (Bedding 1993). The mermithid Romanomermis culicivorax









Ross and Smith has been used as a successful, inundative biological control agent for

mosquito larval suppression (Petersen 1985).

Steinernematid and heterorhabditid nematodes are second only to Bacillus

ith1n iigiJel'i, Berliner in commercial sales at $2-3 million dollars annually (Georgis

1997). These nematodes infect a number of insect species yet pose no threat to plants,

vertebrates, and many invertebrates (Akhurst 1990, Kaya and Gaugler 1993). They can

be mass produced, formulated, and easily applied as biopesticides; they also have been

exempt from registration in many countries, are compatible with many pesticides, and are

amenable to genetic selection (Georgis and Kaya 1988, Kaya and Gaugler 1993, Georgis

and Manweiler 1994).

Nematodes in the families Steinernematidae and Heterorhabditidae are especially

efficacious against insects in soil and cryptic habitats (Lacey et al. 2001). These

nematodes have been used inundatively in many high value crop systems (Georgis and

Manweiler 1994, Koppenhofer 2000). Successful uses of these nematodes against

economically important pests include the citrus root weevil, Diaprepes abbreviatus (L.),

in citrus, the black vine weevil, Otiorhynchus sulcatus (F.), in nurseries and cranberries,

the peach borer moth, Carposina niponensis Walsingham, in apples, and the black

cutworm, Agrotis ipsilon (Hufnagel), in turfgrass (Lacey et al. 2001).

Objectives

Greater understanding of alternative controls, such as using insect parasitic

nematodes and how they fit into an integrated pest management program for mole

crickets in turfgrass, is needed. This research specifically assessed the effectiveness of

Steinernema scapterisci through a series of laboratory and field experiments. The

research included evaluating 1) the establishment of S. scapterisci in soil growing high






11


value golf course turfgrass, 2) effective drenching solutions in order to sample areas to

determine levels of nematode infected mole crickets, 3) compatibility of S. scapterisci

with insecticides, and 4) the effect of S. scapterisci infection on the reproductive and

tunneling behavior of pest mole crickets.














CHAPTER 2
ESTABLISHMENT AND SPREAD OF Steinernema scapterisci ON FLORIDA GOLF
COURSES

An integrated pest management program for pest mole crickets (Scapteriscus

abbreviatus, S. borellii, and S. vicinus) is being developed throughout the southeastern

United States. These insects damage turfgrass by tunneling and root feeding, that results

in large, irregular patches of dead turf throughout the year. Chemical control on golf

courses is still the primary means of preventive and curative control. Using resistant or

tolerant varieties or species of turfgrass, such as the bermudagrass hybrids 'TifSport' and

'Ormond', is possible but not common (Hudson 1986, Hanna and Hudson 1997, Braman

et al. 2000, Hanna et al. 2001, Reinert and Busey 2001). Cultural controls (i.e., adjusting

irrigation, fertilization, mowing heights, etc.) have not affected mole cricket populations

(Denisenko 1986, Frank et al. 1998, Frank and Parkman 1999. However, the introduction

and conservation of natural enemies that attack mole cricket adults is gaining momentum,

especially on pastures and Audubon International affiliated golf courses

The purpose of this study was to establish the efficacy of augmentative releases of

S. scapterisci against Scapteriscus spp. mole crickets on highly maintained golf courses

and athletic fields (Appendix B) as well as determining if subsequent applications of

nematodes would augment the nematode populations in the test area and increase the

percentage of mole crickets infected.









Materials and Methods

Study Sites

The establishment and spread of S. scapterisci was monitored on two golf courses

in Alachua Co., FL: Ironwood Golf Course and Gainesville Golf and Country Club.

Ironwood Golf Course (IGC) was an 18-hole city-owned public golf course built in 1964.

The roughs were bermudagrass (Cynodon dactylon Pers. x C. transvaalensis Burtt-Davy)

var. Tifway, mowed at 4.7 cm. Gainesville Golf and Country Club (GGCC) was an 18-

hole private course located 10.3 miles from Ironwood Golf Course. Gainesville Golf and

Country Club was built in 1962 and originally planted with bermudagrass var. Ormond

and the roughs were mowed at 3.2 cm. Ironwood Golf Course and GGCC had been

previously treated with S. scapterisci in the late-1980s and did not have any subsequent

treatments. Weather data, including minimum and maximum daily temperatures, relative

humidity, amount of monthly precipitation (from local weather stations located

approximately 1 mile from IGC and 9 miles from GGCC) and soil temperatures at 7.6 cm

below the soil surface, were documented on each collection date (Appendix A).

Mole Cricket Monitoring

Twenty hot spots of mole cricket activity were located in the roughs of ten fairways

on each golf course (two hot spots per hole). Linear pitfall traps (modified from

Lawrence 1982) were installed in the ground at least 80 m apart. Each golf course

fairway contained a trap located in one treated area and one untreated area.

Pitfall traps consisted of a 19 L plastic bucket placed in the ground and four PVC

(3 m long, 7.6 cm diameter) perpendicular arms with a 2.5 cm wide slit lengthwise along

the top. The arms were placed in the ground so the slits were flush with the soil surface

(Figure 2-1). The distal end of each arm was capped, so insects falling into an arm









eventually fell into a 3.8 L bucket containing approximately 3-5 cm of sand, located

within a 19 L bucket (Figure 2-2). Holes were drilled into the bottom of both buckets to

allow for water drainage. Traps on all sites were installed in September and October

2001.

Nematodes were released in the afternoon (approximately 1600) at Ironwood Golf

Course (31 October 2001) and in the morning (approximately 0700) at the Gainesville

Golf and Country Club (5 November 2001). Nematodes were applied in an aqueous

suspension of 1 billion nematodes/ 378.5 L of water applied using a boom sprayer

calibrated at 0.5 L/m2. The area treated was 20.1 x 20.1 m (0.04 ha) around one trap on

each fairway for both golf courses. All sites were irrigated with 0.6 cm of water before

and 0.6 cm after application. The pre-treatment dates for Gainesville Golf and Country

Club and Ironwood Golf Course were 11, 18, 25 October 2001.

Pitfall traps were used to monitor infection levels and mole cricket abundance

using methods similar to Parkman et al. (1993a, b). At each 24 h sampling period, the

buckets and arms were cleaned out and 3-5 cm of sand was placed into the inner bucket.

Traps were left for 24 h and adult and juvenile mole crickets with pronotal lengths > 4

mm (Hudson and Nguyen 1989a) were collected from traps and returned to the

laboratory. Crickets were tested for infection weekly for the initial 6 wk post-application

and one to two times a month thereafter for 1 yr on Gainesville Golf and Country Club

and 2 yr on Ironwood Golf Course. Turf quality (density, color) in the area immediately

surrounding the pitfall traps was assessed (1 = sparse or brown grass, 9 = dense or dark

green grass). Soil samples were taken from each golf course and soil texture was









analyzed using the sodium metaphosphate/hydrometer procedure. Traps were removed

from GGCC in October 2002 at the superintendent's request.

Laboratory Assay

Percent of mole crickets infected with nematodes caught in the traps was

monitored and tested in the laboratory. Mole crickets were placed individually in 20 ml

plastic scintillation vials (Fisher Scientific) with 1-2 drops of deionized water, capped

and labeled. Mole crickets were examined at 7 and 10 d after death under a dissecting

scope (10X) for the presence of nematodes. Steinernema scapterisci were identified by

Dr. Khuong Nguyen, Entomology and Nematology Department, University of Florida.

Statistical Analysis

Comparisons of infection rates between sites and years were subjected to analysis

of variance and Tukey's studentized range test or Student's t-test (SAS Institute 2001).

All comparisons were made at a 0.05 significance level. Non-transformed means plus or

minus one standard error of the monthly mean are presented.

Results and Discussion

Mole crickets infected with S. scapterisci were collected at both golf courses before

and after our augmentative applications. Infected mole crickets were only found in the

spring and fall of each year when the late instar and adults, the most susceptible life

stages to S. scapterisci infection, were present. Mean cumulative percentages ( SE) of

infection for mole cricket trap collections during the 2001-2003 field season from the

sites GGCC (22.1% + 10.5) and IGC (15.8% + 4.6) did not differ statistically (t = 2.00;

df= 2, 56; P > 0.0001). Monthly infection and baseline pretreatment infection levels for

IGC and GGCC are presented in Figures 2-3 and 2-4, respectively.









Pre-treatment infection rates at GGCC and IGC ranged from 10-15%. The

percentage of infected mole crickets in treated areas at GGCC exceeded pre-application

levels about 4-8 wk after application. The percentage of infected mole crickets in

untreated areas at both courses (>80 m from treated areas) equaled the percent infection

in treated areas after about 20 wk. Significantly fewer Scapteriscus spp. were collected

in year 2 than in year 1 at IGC (t = 2.47; df= 1, 37; P > 0.01). The S. scapterisci

population persisted throughout the entire study period, 1 yr for GGCC and 2 yr for IGC;

however at IGC the level of infection for year 2 was significantly less than year 1 (F =

6.63; df= 1, 37; P > 0.01). During this study fipronil and acephate were used on greens

of both courses; acephate was used in the roughs as a spot treatment on IGC. Data from

this study concur with Parkman et al. (1994) that more Scapteriscus adults were infected

with S. scapterisci than nymphs. Contrary to Parkman et al. (1994), overall infection of

S. borellii was lower than that of S. vicinus in this study. More S. vicinus were collected

in traps than S. borellii (Table 2-1).

Prior applications of S. scapterisci were made in 1988 and 1989 on Ironwood Golf

Course and Gainesville Golf and Country Club, respectively. Infection levels present in

mole crickets during the pre-treatment collections demonstrate that the nematodes can

survive in golf course soil where mole crickets are present and pesticides are used

regularly for 12 years. If the nematodes can persist for at least 12 years, then the

populations will probably last for many more years as effective biological control agents.

Steinernema scapterisci can survive for 10 wk in soil lacking mole crickets (Nguyen and

Smart 1990a), however if crickets are present the nematode recycles within the host in

soil environments and could survive for 12 years as demonstrated by our results. The









persistence of nematodes in the soil may reduce the populations of mole crickets leading

to an overall reduction of pesticide applications needed.

Establishment and persistence of S. scapterisci in pastures was evaluated by

Parkman et al. (1993a) for five years. Nematodes were applied as an aqueous solution

using a handheld watering can or by burying infected mole cricket cadavers at the test

sites. Crickets were collected using linear pitfall traps (Lawrence 1982) with electronic

callers located nearby to enhance local mole cricket populations. Based on their findings

S. scapterisci became established and persisted for five years at these pasture field sites.

Parkman et al. (1993b) also evaluated the efficacy of a single inoculative release of

S. scapterisci against mole crickets in pastures. Nematodes were applied in an aqueous

solution using a tractor-drawn chisel rig. Crickets were collected in pitfall traps placed at

50, 100, and 200 m from the center of the treated area. The nematode persisted at five of

the six sites and dispersed at least 150 m away from the initial treated area at three of the

sites (Parkman et al. 1993b). They found that inoculative releases of S. scapterisci were

an alternative to inundative releases for mole cricket suppression. Further tests (Parkman

et al. 1994) showed that S. scapterisci serves as an inoculative biological control agent

for Scapteriscus spp. mole crickets on golf courses and acted as a biopesticide for

relatively rapid suppression of pest populations (Parkman et al. 1993b). Establishment on

golf courses was not as successful as pastures; however there was a >27% pest population

reduction in areas where the nematode did persist (Parkman and Smart 1996).

Since its initial release in June 1985 in Alachua Co., Florida, S. scapterisci has

been non-commercially released and presumably established in at least 28 counties in

Florida. The nematode population established readily even after a single inoculative









release. The establishment of S. scapterisci has been demonstrated by strip and broadcast

spray applications. Mole crickets infected with S. scapterisci can fly several kilometers

before dying (Parkman and Frank 1992), thus potentially spreading nematodes to

uninfected sites. Therefore the use of S. scapterisci as a biopesticide would allow

treatment to hot spots rather than strict reliance on broadcast applications of synthetic

insecticides to provide long-term mole cricket suppression. However, the effect of

augmentative applications to hot spots of mole cricket activity has not been previously

determined.

From our results it is evident that a single augmentative application of S.

scapterisci is sufficient to enhance nematode populations throughout a local area (i.e., a

fairway). Augmentative applications of nematodes provided higher than baseline

infection levels for 17 mo post treatment at IGC and 8 mo post treatment at GGCC.

Infection levels fluctuated with the host population in our study. Mole cricket adults and

large nymphs are present in the spring and fall months of any given year therefore,

crickets infected with S. scapterisci were usually found in larger numbers during these

months. The reduction in mole cricket population levels in year 2 versus year 1 could be

attributed in part to the S. scapterisci applications suggesting that the augmentation of

nematode populations does help reduce the numbers of mole crickets present on highly

managed golf courses in north Florida. Levels of infection may also be influenced by

mole cricket age, activity, environmental conditions and predation by other arthropods

within a trap. Although a single application can establish nematode populations, an

augmentative application may be necessary to keep population levels high enough for

sufficient mole cricket control.









Mole crickets collected from the pitfall traps were subject to mortality from

organisms other than S. scapterisci. Organisms observed infecting mole crickets

included other species of insect parasitic nematodes (i.e., Heterorhabditis spp. and

Steinernema spp.), fungi (i.e., Beauvaria bassiana), predators (i.e., ground beetles,

spiders, earwigs, etc.), and other mole cricket parasitoids (Larra bicolor and Ormia

depleta).

Previous research (Gaugler and Boush 1978, Molyneux 1985, Hudson and Nguyen

1989b, Smith 1999) has shown that there are several abiotic factors that contribute to the

success (or failure) of insect parasitic nematodes when used as a biopesticide. Abiotic

factors can significantly limit the nematodes' effectiveness to move, locate and enter a

host (Smith 1999). Some of these abiotic factors include ultraviolet light, desiccation,

soil moisture, soil texture and type, soil temperature, soil pH, and agrichemical

compatibility (see Chapter 4). In this study applications were done in the early morning

and late afternoon and watered in immediately after application to avoid nematode

damage from UV light and desiccation. Moisture is required for nematodes to move

through the soil (Hudson and Nguyen 1989b). Ames (1990) observed that infective

juveniles of S. scapterisci can survive up to 13 wk at wilting point (15 bars moisture

tension) and survive better in sandy loam than pure sand. In this study both courses had

sandy loam soil and were irrigated regularly. Several of the fairways and roughs on both

courses where crickets were collected were subject to flooding or extremely wet

conditions due to rain or excess irrigation. Molyneux and Bedding (1984) observed that

very saturated soils can inhibit nematode mobility and decrease their survival by creating

anaerobic conditions.






20


Steinernema scapterisci killed between 15-20% (on average) of mole crickets

collected in linear pitfall traps. However, because mole crickets may die as soon as 48 h

post-infection, this value may be an underestimate of the true percentage of kill.

Nematodes can kill generation after generation and this combined effect may be

something similar to compound interest (Frank et al. 2002). Steinernema scapterisci can

be an effective part of an integrated pest management system on managed turfgrass if

applied in the proper manner and in suitable locations where nematodes can survive.

























Figure 2-1. Linear pitfall trap used to collect mole crickets.


Figure 2-2. 3.8 L catch bucket inside 19 L bucket of linear pitfall trap.











60
D Untreated
m Treated
50 27 4
19
40 34

> 91 184
30 -
o 3
4 13
-c20 78 4


10 10 78 56


0
OCT01 NOV01 DEC 01 JAN 02 FEB02 MAR 02 APR02 MAY 02 AUG02 OCT02 MAR 03

Month




Figure 2-3. Mean monthly ( SEM) percent infection of mole crickets collected in pitfall
traps at Ironwood Golf Course from areas treated with Steinernema
scapterisci. Only months with infection levels are presented. Untreated areas
received no S. scapterisci and were >80 m from treated areas. Dashed line
represents baseline pretreatment infection level. Data presented are for
Scapteriscus vicinus and Scapteriscus borellii combined. Total numbers of
mole crickets are presented above SEM bars.











D Untreated
* Treated


OCT01 NOV 01 DEC 01 MAR 02 APR 02 MAY 02
Month


Figure 2-4. Mean monthly ( SEM) percent infection of mole crickets collected in pitfall
traps at Gainesville Golf and Country Club from areas treated with
Steinernema scapterisci. Only months with infection levels < 0 are presented.
Untreated areas received no S. scapterisci and were >80 m from treated areas.
Dashed line represents baseline infection level. Data presented are for
Scapteriscus vicinus and Scapteriscus borellii combined. Total numbers of
mole crickets are presented above SEM bars.











Table 2-1. Percent infection (mean SEM) of Scapteriscus spp. mole crickets collected
from sites treated with Steinernema scapterisci.

Gainesville Golf and Country Ironwood Golf Course
Club (n=10 traps) 1 (n= 10 traps) 2
S. borellii
Nymphs 0(1) 0(15)
Adults 0(1) 3.3 2.6 (20)
Total 0 (2)* 1.6 + 1.3 (35)*
S. vicinus
Nymphs 7.7 4.0 (70) 9.7 3.4 (208)*
Adults 13.9 + 4.5 (163) 33.2 7.0 (313)
Total 10.8 3.0 (233) 11.9 2.5 (521)
Pairs of means within columns followed by asterisks are significantly different, t-test (P >
0.05). Numbers in parentheses are the total mole crickets collected.
1 GGCC: F= 8.57; df= 2,111; P= 0.0003
2 IGC: F= 14.05; df= 2,155; P< 0.0001














CHAPTER 3
SURVIVAL AND INFECTIVITY OF Steinernema scapterisci AFTER CONTACT
WITH SOIL DRENCH SOLUTIONS

Mole crickets are subterranean pests of turfgrass in Florida and much of the

southeastern United States (Walker and Nickle 1981, Walker 1985). Mole cricket

damage and cost of control in Florida in 1986 were estimated at $45 million with an

additional $33 million in Alabama, Georgia, and South Carolina combined (Frank and

Parkman 1999). Estimates of annual expenditure in 1996 were over $18 million for

insecticides in Florida turf, and over $12 million in control costs (Hudson et al. 1997).

Mole crickets damage turf by their tunneling in the soil, which exposes and dries out

roots and by direct root feeding. As a result, the turfgrass thins and bare patches appear.

The tunneling and mounds that mole crickets make also disrupt the playing surface on

golf courses, especially the roll of the golf ball on greens. Superintendents and golf

course members have little tolerance for damage (Frank and Parkman 1999). Insecticides

are usually targeted against the most destructive, nymphal stage. A more sustainable,

environmentally friendly management approach for mole cricket control is needed.

Several biological control agents have been investigated for control of Scapteriscus

spp. mole crickets in Florida (Hudson et al. 1988). One of these biological control agents

is an entomopathogenic nematode, Steinernema scapterisci. Steinernema scapterisci was

originally isolated from pitfall-trapped Scapteriscus vicinus in Uruguay in the 1980s

(Nguyen and Smart 1990b). The nematode was cultured and released in several Florida

counties in 1985, where it established a population, and was spread from the release site









by infected Scapteriscus mole crickets (Hudson et al. 1988, Parkman and Frank 1992).

The nematode kills the adult and late instar nymphs of Scapteriscus borellii and S.

vicinus, and to a lesser extent S. abbreviatus. Fewer small to medium-sized nymphs of S.

borellii and S. vicinus become infected (Nguyen 1988).

Several techniques have been used to sample mole crickets including counts of

dead nymphs and adults after insecticide applications (Short and Koehler 1979),

estimation of surface burrowing (Walker et al. 1982, Cobb and Mack 1989), pitfall

trapping (Lawrence 1982, Adjei et al. 2003), removal with a tractor mounted soil corer

(Williams and Shaw 1982), sound trapping (Walker 1985) and soil drenching (Short and

Koehler 1979, Walker 1979, Hudson 1989). However, results from each of these

techniques are often inconsistent (Short and Koehler 1979, Lawrence 1982, Hudson

1988). Comparisons of different methods have indicated that soil drenching with soap

solutions are the most practical and consistent at obtaining direct counts of mole crickets

(Short and Koehler 1979, Hudson 1988).

Soil drenching with a solution of 15 ml of lemon dishwashing detergent in 3.8 L of

water is inexpensive and commonly used by turfgrass managers to sample soil pests. Soil

drenches with soap solutions irritate mole crickets and force them out of the soil. Soap

flushes are often used for monitoring mole crickets to determine the size, age, and species

present, the relative population density over time, and for control timing. However, it

was suspected that soap flushes, when used to monitor mole crickets potentially infected

with S. scapterisci, might be lethal to the nematodes because we rarely found nematodes

in soap-flushed mole crickets (K.B. Nguyen and G.C. Smart, Entomology and

Nematology Dept., University of Florida, pers. comm.). Solutions such as pyrethroids,









ammonia, vinegar, Lysol, and other soap detergents have previously been tested as

potential soil drench solutions (Short and Koehler 1979).

This study was conducted to determine whether a standard soap detergent solution

affects S. scapterisci survival and infectivity in pest mole crickets. Potential alternatives

to the standard soap drench solution were also evaluated.

Materials and Methods

Nematodes and Mole Crickets

Steinernema scapterisci (Nematac S, Becker Underwood, Ames, IA) was stored

in a 70C cold room until use (<3 mo). Nematode viability was tested before each

application by dissolving a pinch (-10 mg) of Nematac S into water and observing

nematode shape and mobility under a light microscope. Healthy nematodes were opaque

in color and S-shaped with undulating movements. Dead or unhealthy nematodes were

translucent, straight, and lacked movement. The product was used if viability was >50%

and discarded if <50% viable.

Scapteriscus vicinus were collected from pitfall traps or sound traps in Alachua

Co., FL, and returned to the laboratory. Each mole cricket was placed in a 120-ml plastic

vial (Thornton Plastics Salt Lake City, UT) with sterilized sand and held for >14 d to

ensure health. Surviving mole crickets were used in this study. Mole crickets were

maintained at 230C with a photoperiod of 12:12 (L:D) and fed commercial cricket chow

(Purina, St. Louis, MO).

Bioassay

Nematode viability and infectivity were assessed after exposure to various

drenching materials. Steinernema scapterisci nematodes were extracted from Nematac









S using a modified Baermann technique (Nguyen and Smart 1990b). Steinernema

scapterisci nematodes were kept at a density of 10,000 infective juveniles in solutions of

water (control), lemon dishwashing detergent (Joy, Proctor and Gamble, Cincinnati,

OH), insecticidal soap (Safer Soap, Woodstream Corporation, Litiz, PA), and

permethrin (Spectracide Bug Stop, Spectrum Brands, St. Louis, MO) for test 1. The

mixtures were kept at room temperature in a 125-ml Erlenmeyer flask with 125-ml per

flask on a shaker at 65 rpm. There were five replicates for each treatment.

Concentrations were selected based on recommendations for flush extraction of mole

crickets in the field (Short and Koehler 1979) and label rates for mole cricket control.

After 24 h, 10-[l samples were taken from each treatment and placed on a microscope

slide. The numbers of living and dead nematodes were counted using a dissecting

microscope (10 x), three 10-[l counts were taken and averaged to determine percent

mortality for each replicate. Immobile nematodes were touched with a probe to determine

survival.

A second test was initiated to further test potential drench materials. Treatments

for test 2 included water (control), azadirachtin (Safer Brand BioNeem, Woodstream

Corporation, Litiz, PA), citrus oil (Green Sense, Garland, TX), garlic extract (Garlic

Barrier, Garlic Research Labs, Inc., Glendale, CA), lemon juice (ReaLemon, Mott's,

St. Louis, MO), permethrin (Spectracide Bug Stop, Spectrum Brands, St. Louis, MO)

and cyfluthrin (Bayer Advanced Lawn and Garden Complete Insect Killer, Bayer

Environmental Sciences, Montvale, NJ). Concentrations were selected based on label and

half label rates for mole cricket control. Methods from test 1 were repeated.









Nematode infectivity was assessed by filtering nematodes from above solutions and

adding 50 living infective juveniles (parasitic stage) to 120 ml plastic cups (Fisher

Scientific) containing 20 g sterilized sand, 4% deionized water, and one S. vicinus adult.

Dead mole crickets were dissected and the presence or absence of nematodes was

recorded.

The above solutions were tested for their effectiveness at flushing mole crickets at

the University of Florida G.C. Horn Turfgrass Research Unit in Gainesville, FL, on 20

and 28 May 2003. Each treatment from tests 1 and 2 (3.8 L of each solution) was applied

to areas of bermudagrass (Cynodon dactylon Pers. x C. transvaalensis Burtt-Davy) var.

Tifway, that had mole cricket damage (75 x 75 cm2). The numbers of adult and first

instar mole crickets emerging from the soil within 3 min were counted. Five replicates

for each solution were completed. Any turfgrass phytotoxicity was noted 1 h post

application and 1 wk post application.

The effect of nematode infected crickets exposed to soap solutions was also tested.

Scapteriscus abbreviatus adults were obtained from a lab colony at the University of

Florida Entomology and Nematology Department, Gainesville, FL, and were inoculated

with about 10,000 nematodes by applying a predetermined amount (approximately 150

[l) of concentrated nematode solution onto a piece of filter paper (Fisher #P8, 5.5 cm)

inside a petri dish with one S. abbreviatus adult. The mole cricket was allowed to

incubate in the petri dish for 1, 5, 8, 12 or 24 h (five mole crickets per treatment).

Scapteriscus abbreviatus was used because S. vicinus adults were unavailable at the time

of the test. All infected mole crickets were then dipped into a 118 ml Solo souffle cup

(Gainesville Paper Co., Gainesville, FL) containing the soapy water or soapy water









followed by a deionized water rinse for 5 sec. Untreated controls were healthy,

uninfected mole crickets dipped in water. Mole crickets were placed into 20-dram plastic

scintillation vials (Fisher Scientific, Pittsburgh, PA) and observed every 24 h for 10 d.

On day 10, mole crickets were dissected and the presence of nematodes was noted.

Statistical Analysis

Nematode mortality and field test data were subjected to an analysis of variance

(SAS Institute 2001). Treatments were compared to the control (water) using Dunnett's

means comparison method (a = 0.05). Nematode infectivity data were subjected to Chi-

square analysis (SAS Institute 2001). Treatments were compared to the control (water)

and the standard soap flush solution (4 ml lemon dish detergent/L water) using Dunnett's

means comparison method (alpha= 0.05). Nematode mortality data were transformed

using arcsine-square root transformation before statistical analysis; nontransformed data

are presented. Effects of nematode infected crickets exposed to soap solutions data were

subjected to ANOVA (SAS Institute 2001).

Results and Discussion

Permethrin at the label rate for mole cricket control caused significantly more

nematode mortality than resulted from water (Table 3-1). Nematodes exposed to all

treatments showed similar infectivity in mole crickets. Nematode mortality was similar

among all treatments in test 2 except citrus oil (Table 3-2). Nematodes surviving all

treatments, except azadirachtin and lemon juice, demonstrated a low percentage

infectivity of mole crickets, no significant treatment differences were.

In the field, insecticidal soap and the higher rate of permethrin flushed significantly

more mole crickets than water (Table 3-3). However, when all treatments were

compared to the standard lemon dish detergent, insecticidal soap and permethrin brought









a similar number of mole crickets to the surface (n = 55; F = 2.88; df= 10,54; P = 0.008).

None of the mixtures tested produced any noticeable phytotoxicity to the turf.

Soil drenches with a mixture of lemon dish detergent and water are commonly used

to monitor turfgrass insects such as mole crickets, chinch bugs (Blissus spp.), big-eyed

bugs (Geocoris spp.), and several species of caterpillars (Short and Koehler 1979,

Hudson 1989). Soil drenches are inexpensive and are not labor intensive when compared

with other methods of monitoring mole cricket populations. These other methods include

large linear pitfall traps (Lawrence 1982, Adjei et al. 2003), an emitter producing a

synthetic song of male mole crickets (Parkman and Frank 1993), and a soil-coring device

(Williams and Shaw 1982). Each method requires more than one person, and is labor

intensive or costly (Lawrence 1982, Williams and Shaw 1982).

Seventy percent of S. scapterisci died in the lemon dish detergent solution. Assays

for nematode infection of soap-flushed mole crickets, the method currently used by many

turfgrass managers, are likely to be inaccurate. Krishnayya and Grewal (2002) reported a

toxic effect of a common soap surfactant (Ajax) on S. feltiae Bovien nematodes. They

found a 24% mortality level of nematodes when incubated at 4, 24, 72, and 120 h

(Krishnayya and Grewal 2002). Kaya et al. (1995) reported an insecticidal soap (M-

Pede) adversely affected S. carpocapsae (Weiser) and Heterorhabditis bacteriophora

Poinar survival and infectivity. However, infectivity may not be affected if the

nematodes are combined with an insecticidal soap and applied immediately (Kaya et al.

1995). Nematodes cannot be stored in an insecticidal soap solution because without

aeration, nematode survival can be adversely affected (Kaya et al. 1995). The toxicity of









metal ions present in soap may be responsible for the high mortality in soap solutions

(Jaworska et al. 1994, Krishnayya and Grewal 2002).

Tests of exposure of nematode infected mole crickets to soap solutions show that

soap flush solutions do not greatly affect nematode infection at least 8 h post infection

(Table 3-4). The soap flush solutions may potentially kill nematodes in certain areas of

the body (i.e., mouth) and further testing should be done to determine this. Immediately

rinsing flushed mole crickets with clean water may potentially increase the accuracy of

determining nematode infection. The unavailability of S. vicinus at the time of

experimentation may have also led to inconsistent, low levels of infection. It is known

that S. scapterisci does not infect S. abbreviatus as successfully as S. vicinus or S. borellii

(Nguyen 1988).

Although permethrin solutions killed some nematodes in our experiments, S.

scapterisci infectivity was not compromised and field flushes successfully extracted mole

crickets from the soil. Short and Koehler (1979) reported that pyrethrins were the most

effective material, flushing a mean of 19 mole crickets/m2. Hudson (1988) compared

three sampling techniques, soil flushing with lemon dish detergent or synergized

pyrethrins and a tractor mounted soil corer. None of the methods gave significantly

different results. Our results from the field test show drenching solutions of permethrin

are useful in determining whether mole crickets collected in the field are infected with S.

scapterisci nematodes. A soil drench containing permethrin may be the best monitoring

tool to flush mole crickets to determine the presence of S. scapterisci.

However, there are disadvantages to using pyrethroids as soil drenches for mole

crickets. Pyrethroid drenches at the half or full label rate may cause more mole cricket









mortality than using a soap solution. Subsurface mortality of mole crickets can be as

high as 65% when using pyrethroids or similar insecticides (Ulagaraji 1974, 1975;

Walker 1979; Hudson 1988). Applicator exposure to insecticides is increased when

using a pyrethroid soil drench.

Soil drenches are effective, non labor-intensive methods to sample soil insect

populations. Soap detergent solutions, although inexpensive, may not accurately indicate

mole crickets infected with S. scapterisci. Permethrin solutions are less cost effective,

but are effective at flushing mole crickets potentially infected with nematodes.











Table 3-1. Mean nematode mortality and percent of mole crickets infected with
Steinernema scapterisci after exposure for 24 h to various drenching
solutions.


Rate


Mean % nematode
mortality ( SEM)1


Number of mole
crickets infected with
S. %q' iu/'/i% i (n=3)


Water n/a 2.0+ 1.4 2
Lemon Joy 15 ml/ 3.79 L 32.4 + 1.6 0
Insecticidal Soap 15 ml/ 3.79 L 40.0 + 5.5 1
Permethrin 18 ml/3.79 L 11.6+ 1.0 2
* Statistically significant values using Dunnett's method comparing treatments to water
1n 20; F= 34.58; df= 19, 3; P = <0.0001
2 More than 10 replicates are needed for statistical analysis.







Table 3-2. Mean nematode mortality and infectivity after exposure for 24 h to various
drenching solutions.


Treatment


Rate


Mean % nematode
mortality ( SEM)1


Water n/a 3.6 +2.2
Citrus Oil 15 ml/ 3.79 L 10.6 + 3.2
Cyfluthrin 7.5 ml/ 3.79 L 2.2 + 2.2
Cyfluthrin 15 ml/ 3.79 L 0.6 + 0.6
Garlic Extract 111 ml/ 3.79 L 0
Lemon Juice 15 ml/ 3.79 L 1.8 + 1.8
Azadirachtin 60 ml/ 3.79 L 0.8 + 0.8
Permethrin 9 ml/3.79 L 2.0 + 1.3
Permethrin 18 ml/ 3.79 L 0
n = 45; F 3.80; df =44, 8;P =0.0025
2 More than 10 replicates are needed for statistical analysis.


Number of mole crickets
infected with S.
, ,q'/'//l i2 (n=3)
1
1
1
1
1
0
0
1
1


Treatment












Table 3-3. Mean number of mole crickets emerging from bermudagrass using various
drenching solutions in May 2003.


Rate


Mean number of mole
crickets flushed ( SEM)


Water
Citrus oil
Cyfluthrin
Cyfluthrin
Garlic extract
Lemon juice
Azadirachtin
Permethrin
Permethrin
Insecticidal soap


n/a
15 ml/3.79 L
7.5 ml/ 3.79 L
15 ml/ 3.79 L
111 ml/ 3.79 L
15 ml/3.79 L
60 ml/ 3.79 L
9 ml/ 3.79 L
18 ml/3.79 L
15 ml/3.79 L


2.6 1.6
0.2 0.2
4.0 2.1
0.4 0.2
0.6 0.4
3.2 1.2
2.6 1.1
5.8 1.4*
5.4 1.3 *


Lemon joy 15 ml/3.79 L 4.6 2.1
* Means statistically significant values using Dunnett's method comparing treatments to
water n = 54; F= 2.88; df= 59, 10; P= 0.01


Table 3-4. Percent nematode infection from mole crickets exposed to treatment solutions
1, 5, 8, 12, or 24 h post infection.


Time Post Infection
Joy (15 ml/ 3.79 L)
Joy (15 ml/ 3.79 L) + H20 rinse
Control1


Ih 5h


8h 12h


0 40 60*
40 40 100*


n = 75;F= 6.77; df= 14, 2; P < 0.0001
* Means within columns statistically significant values when compared to control.
1 Control = uninfected, healthy mole crickets immersed in water.


Treatment


100*
100*
0














CHAPTER 4
INTEGRATION OF INSECT PARASITIC NEMATODES WITH INSECTICIDES FOR
CONTROL OF PEST MOLE CRICKETS

Appreciation of high-quality turfgrass has recently led to rapid growth in the golf

course and landscape management industries (Zimmerman and Cranshaw 1990).

Associated with this growth have been an increased number of pesticide applications and

environmental concerns (Zimmerman and Cranshaw 1990). Restrictive legislation has

resulted in a greater need for alternative control methods. Insect parasitic nematodes

provide acceptable control of several soil and thatch infesting pests, such as white grubs

(Coleoptera: Scarabaeidae), billbugs (Coleoptera: Curculionidae), cutworms

(Lepidoptera: Noctuidae), and mole crickets (Orthoptera: Gryllotalpidae) (Zimmerman

and Cranshaw 1990).

Insect parasitic nematodes could be effective in integrated pest management

programs as long-term suppressive agents used in combination with quick knockdown

products like commercially available insecticides. Effects of selected pesticides on

entomopathogenic nematodes other than S. scapterisci have been reported in aqueous

solutions (Prakasa et al. 1975, Hara and Kaya 1983a, Das and Divakar 1987, Rovesti and

Deseo 1990, Zimmerman and Cranshaw 1990, Ishibashi and Takii 1993, Gordon et al.

1996, Koppenhofer and Kaya 1998) and in insect hosts following pesticide exposure

(Hara and Kaya 1982, 1983b). Several studies have demonstrated that pesticides can

decrease survival and infectivity of several entomopathogenic nematode species in the

families Steinernematidae and Heterorhabditidae (Hara and Kaya 1982, 1983a, 1983b;









Forschler et al. 1990; Zimmerman and Cranshaw 1990; Head et al. 2000). However,

combinations of insecticides and insect parasitic nematodes have a synergistic effect on

nematode infection rates against white grubs (Koppenh6fer and Kaya 1998; Koppenh6fer

et al. 2000a, 2000b; Koppenh6fer et al. 2002). It is important to determine what

interactions, if any, might occur when certain insecticides are used together with S.

scapterisci.

Knowing which insecticides might affect nematode performance is important in the

development of integrated pest management programs. Our first objective was to assess

the compatibility of chemical and biological management of pest mole crickets. A

second objective was to determine if label rates or half-label rates of these insecticides

could increase nematode infection rates.

Materials and Methods

Steinernema scapterisci (Nematac S, Becker Underwood, Ames, IA) used in this

study was stored in a 70C cold room until use (< 3 mo). Nematode viability was tested

before each application by dissolving a pinch of Nematac S into water and observing

nematode shape and mobility under a light microscope. Healthy nematodes were opaque

in color and S-shaped with undulating movements. Dead or unhealthy nematodes were

translucent, straight, and lacked movement. The product was used if viability was >50%

and discarded if <50% viable.

Scapteriscus vicinus adults were collected from pitfall and sound traps in Alachua

County, FL, and returned to the laboratory. Each mole cricket was placed in a 120-ml

plastic vial (Thornton Plastics, Salt Lake City, UT) with sterilized sand and held for >14d

to ensure health; only mole crickets that appeared healthy were used in this study. Mole









crickets were maintained at 24-270C, with a photoperiod of 12:12 (L:D), and fed

commercial cricket chow (Purina, St. Louis, MO).

Five insecticides commonly used for mole cricket control were evaluated for their

effects on the entomopathogenic nematode S. scapterisci in the laboratory. They

included acephate (Orthene Turf, Tree and Ornamental Spray, Valent Professional

Products, Walnut Creek, CA), bifenthrin (Talstar GC Flowable, FMC Corporation,

Philadelphia, PA), imidacloprid (Nleiit' 75 WP, Bayer Environmental Science,

Montvale, NJ), fipronil (Chipco Choice granular, Bayer Environmental Science,

Montvale, NJ) and deltamethrin (DeltaGard T&O granular, Bayer Environmental

Science, Montvale, NJ). These commercially formulated insecticides were selected due

to their widespread use for mole cricket control in the southeastern United States.

However, only products or formulations that are typically mixed with water or watered in

were chosen for this test. Products were tested at half and full label rates.

Survival and Infectivity of S. scapterisci After Exposure to Pesticides

Nematode viability and infectivity were assessed after exposure to the pesticides.

To count nematodes, S. scapterisci were extracted from Nematac S using a modified

Baermann funnel technique (Nguyen and Smart 1990b). Approximately 40-50 g

Nematac S was placed on two layers of unscented, non-lotion, white facial tissue

(Puffs, Procter & Gamble, Cincinnati, OH). The tissue was then placed on top of a 1-

mm screen filter in a square plastic container (13 x 13 cm2) and moistened with deionized

water. Approximately 100-110 ml of deionized water was added to the plastic container.

The setup was then placed on a laboratory counter at 24-270C for 5-8 h, during which









time the living nematodes moved through the tissue and filter into the water below. The

nematodes were then counted by taking an average of three, 10-rl samples.

Steinernema scapterisci were kept at a density of 10,000 infective juveniles per 125

ml in aqueous solutions of water (control), half or full label rates of acephate (1 or 2 kg

AI/ha), bifenthrin (112 or 224 g AI/ha), or imidacloprid (275 or 451 g AI/ha). The

mixtures were kept at room temperature (24-27 C) in a 125-ml Erlenmeyer flask on a

rotator at 65 rpm. There were five replicates for each treatment. After 24 h, 10-rl

samples were taken from each flask and placed on a microscope slide. The numbers of

living and dead nematodes were counted using a dissecting microscope (10 x). Immobile

nematodes were touched with a minute probe to determine survival. Three 10-rl counts

were averaged to determine percent mortality for each replicate. Nematode infectivity

was assessed by removing nematodes from above solutions and adding 50 living infective

juveniles to 120 ml plastic cups (Fisher Scientific, Pittsburgh, PA) containing 50 g of

sterilized sand moistened with deionized water and one S. vicinus adult. After death,

mole crickets were dissected and the presence or absence of nematodes was recorded.

Nematode Infectivity After Exposure to Pesticide Treated Mole Crickets

Percent infectivity of nematodes when exposed to mole crickets treated with

insecticides was also assessed. Mole crickets were placed in 120 ml plastic cups

containing 50 g of sterilized sand that was treated with label or half label rates of

acephate, bifenthrin, imidacloprid, fipronil (140 or 280 g AI/ha), or deltamethrin (73 or

146 g AI/ha). Pesticides were applied to the top of the sand and mixed into the sand by

capping the cup and shaking by hand before adding the mole cricket. The mole crickets

were allowed to move through the insecticide treated sand for 24 h. Mole crickets were

then removed and placed into new cups containing 50 g of sterilized sand moistened with









deionized water and nematodes. To ensure mole cricket infection the number of

nematodes used was increased to 500 infective juvenile S. scapterisci. Mole crickets

were fed cricket chow twice a week. Dead mole crickets were removed from the sand

cups and placed individually in 20 ml plastic scintillation vials. Mole crickets were

examined 5-7 d after death under a dissecting scope (10 X) for the presence or absence of

nematodes. The test concluded at 47 d, by which time all of the mole crickets had died.

There were five replicates (mole crickets) for each treatment.

Statistical Analysis

Nematode mortality data were subjected to an analysis of variance (SAS Institute

2001). Treatment means were compared by Tukey-Kramer Honestly Significant

Difference means separation test (P= 0.05). Nematode infectivity data are presented as

number of crickets infected with S. scapterisci out of five total crickets tested. Nematode

mortality data were transformed using arcsine square root transformation before

statistical analysis; nontransformed data are presented.

Results and Discussion

None of the solutions tested reduced S. scapterisci survival (Table 1). Survival

ranged from 95.6% (bifenthrin, label rate) to 100% (imidacloprid, half label rate). The

average number of days until mole cricket death was not significantly different from the

control in any treatment except imidacloprid (Table 1; F= 1.83; df=10, 54; P<0.001).

None of the infectivity levels were significantly different from the control. Between 2

and 4 of the mole crickets tested became infected with the nematodes from the solutions

tested. Mole crickets treated with imidacloprid survived significantly longer than those

in the other treatments, but it was not significantly different from the control (Table 2).

The percent nematode infection of mole crickets previously exposed to solutions of









insecticides ranged from 40-100% (Table 2; F=14.78; df=10, 54; P<0.001). Infectivity

did not significantly differ among any of the treatments tested.

These results suggest that the pesticides tested can be successfully used with S.

scapterisci applications. Organophosphate insecticides can kill S. carpocapsae (Weiser)

infective juveniles (Dutky 1974), or as with carbamates, paralyze them (Hara and Kaya

1982, 1983a). The organophosphate used in our study, acephate, did not have a

significant effect on survival and infectivity of S. scapterisci adults. Our results indicate

that the pesticides tested in this study are compatible with S. scapterisci infective

juveniles.

Previous laboratory studies with other steinernematid nematodes (excluding S.

scapterisci) have demonstrated compatibility with pesticides (Ishibashi et al. 1987,

Zimmerman and Cranshaw 1990, Ishibashi and Takii 1993, Head et al. 2000). Results

from field collected mole crickets have demonstrated that S. scapterisci applied to

intensively managed bermudagrass (Cynodon spp.) in Gainesville, FL, in October 1989

persisted and were recovered in mole crickets collected in October 2001 (Frank et al.

2002). This evidence along with the results presented here support the compatibility of

using S. scapterisci with current mole cricket management strategies. Frank and

Parkman (1999) stated that the optimal use of S. scapterisci would be in the roughs of

golf courses and areas where tolerance for damage is higher and turf is less intensively

maintained. Our results show that an insecticide treated mole cricket has the potential of

being infected by nematodes and nematode reproduction does occur within the treated

host.









Although our study did not directly test tank-mixing effects, we suggest the

potential for tank mixing exists using the wettable or liquid/flowable insecticides tested in

this study and Nematac S. However, there are several possible limitations to tank-

mixing with S. scapterisci. First, chemicals for mole cricket control are usually applied

in early summer for control of young nymphs while S. scapterisci is applied in the spring

or fall for adult control. Irrigation is recommended before and after Nematac S is

applied, thus tank mixing is only possible with pesticides that require watering in.

Because nematodes are sensitive to extreme heat and ultraviolet light (Gaugler and Boush

1978), they should be applied at dawn or dusk, which may conflict with applicator work

hours. The pesticides tested may provide a quick knockdown for late instar and adult

mole crickets, and S. scapterisci may provide a more sustainable, long-term control of

mole crickets.

An insecticide that could modify an insect's behavior to stop or prevent feeding or

tunneling damage, and also inhibit the insect's ability to defend against parasitism or

infection, would be a valuable component of integrated pest management. For example,

imidacloprid is synergistic with S. glaseri (Steiner) or Heterorhabditis bacteriophora

Poinar against white grubs (Koppenh6fer et al. 2000a, 2000b). Imidacloprid disrupts a

grub's normal nerve function, which drastically reduces its activity, affects grooming and

evasive behaviors, and facilitates nematode attachment onto the cuticle (Koppenh6fer et

al. 2000a, 2000b). In our study, mole crickets treated with imidacloprid survived longer

than those treated with the other insecticides, but still died from nematode infection.

Steinernema scapterisci are ambush nematodes, unlike the cruiser nematodes tested by

Koppenh6fer et al. (2000a, b). Pesticides which increase mole cricket activity, rather









than slow it down, may result in increased contact with ambush nematodes. However, if

grooming is also increased, then nematodes may not be able to attach to the host. The

half-label rates of acephate and deltamethrin had greater infectivity (Table 2).

A more integrated and sustainable management plan for mole crickets is possible

using biological, cultural and chemical control. Natural enemies have become

established in Florida and actively target adult mole crickets and large nymphs. Insect

parasitic nematodes (e.g., S. scapterisci) can be applied as biopesticides for large nymph

and adult mole cricket control in the spring and fall. Cultural controls such as using non-

attractive lights during adult flight periods, monitoring life stages using pitfall traps or

soap flushes in order to time control methods (Hudson 1985), and planting nectar sources

for parasitic insects (Frank et al. 1995) should also minimize infestations while selective

use of preventive and curative insecticides can quickly control young nymphs during the

summer.












Table 4-1. Percent survival, infectivity, and days until death of S. scapterisci incubated in solutions of insecticides for 24 h.

Common Name Trade name Rate Nematode survival Days until mole cricket Number of mole
(%) SEM (n=5) death SEM (n=5) crickets infected (n= 5)
Acephate Orthene TT&O Spray 1 kg AI/ha 96.3 + 1.6a 1.4 + 0.2b 4
2 kg AI/ha 97.5 1.2a 1.4 0.2b 3
Bifenthrin Talstar GC Flowable 112 g AI/ha 95.9 1.7a 2.8 1.0Ob 3
224 g AI/ha 95.6 1.8a 1.6 0.2b 3
Imidacloprid Merit 75 WP 275 g AI/ha 100.0 + Oa 23.2 8.9a 2
451 g AI/ha 98.2 1.8a 22.2 4.0a 2
Control (water) n/a n/a 99.8 0.2a 17.0 3.9ab 4
Mean standard error of the mean (SEM), means within columns followed by different letters are significantly different at a = 0.05
using Tukey's Honestly Significant Difference means separation test.













Table 4-2. Average days until death and percent infectivity by S. scapterisci nematodes to mole crickets exposed to insecticides.


Common Name


Trade name


Rate


Days until mole cricket
death SEM (n=5)


Number of mole crickets
infected (n= 5)


Acephate


Bifenthrin

Deltamethrin

Fipronil

Imidacloprid

Control (water)


Orthene TT&O Spray

Talstar GC Flowable

DeltaGard T&O G

Chipco Choice G

Merit 75 WP

n/a


1 kg AI/ha
2 kg AI/ha
112 g AI/ha
224 g AI/ha
73 g AI/ha
146 g AI/ha
140 g AI/ha
280 g AI/ha
275 g AI/ha
451 g AI/ha
n/a


1.0 + 0b
1.0 + 0b
1.0 + 0b
1.0 + 0b
1.0 + 0b
1.0 + 0b
1.0 + 0b
1.4 0.2b
25.6 9.7a
17.4 6.9a
39.6 5.0a














CHAPTER 5
EFFECT OF Steinernema scapterisci NGUYEN AND SMART EXPOSURE ON MOLE
CRICKET TUNNELING, OVIPOSITION, AND AVOIDANCE BEHAVIOR

Avoidance of nematodes has been observed in soil inhabiting insects (Thurston

1994, Wang et al. 2002, Zhou et al. 2002). Laboratory evaluation of four

entomopathogenic nematodes for control of subterranean termites revealed that

Heterorhabditis indica Poinar repelled termites at high concentrations (Wang et al.

2002), but length of repellency varied with nematode concentration. Ants are also

repelled by products produced by the bacteria within nematodes (Zhou et al. 2002). The

symbiotic bacteria Xenorhabdus nematophila and Photorhabdus luminescens of

Steinernema carpocapsae and Heterorhabditis bacteriophora respectively produce "ant-

deterrent factors (ADF)". These ADFs are compounds that deter scavengers and protect

the nematodes from being eaten during reproduction within host cadavers (Zhou et al.

2002). Nematodes have the ability to detect and avoid toxic compounds in their

environment (Hilliard et al. 2004). The nematode Caenorhabditis elegans has the ability

to avoid toxic compounds such as quinine (Hilliard et al. 2004).

Soil insects have been observed to sense and avoid areas treated with insecticides,

pathogens, or nematodes (Villani et al. 1994, Milner and Staples 1996, Villani et al. 1999,

Thompson 2004). Soil insects are often highly susceptible to pathogenic organisms in the

laboratory, however are rarely infected in the field, indicating a possible behavioral

component of microbial defensive tactics (Villani et al. 1999). Metarhizium anisopliae

(Metsch.) incorporated into soil is repellent to Japanese beetle grubs (Villani et al. 1994).









Termites also detect and avoid Metarhizium conidia in soil (Milner and Staples 1996).

Mole crickets are repelled by soil treated with Beauveria bassiana (Brandenburg 2002,

Villani et al. 2002). Thompson (2004) observed mole crickets avoiding soil treated with

the Beauvaria bassiana strain 10-22 and Talstar in the greenhouse.

This study determined how mole cricket tunneling changes over time after

exposure to varying numbers of S. scapterisci, as well as to determine whether healthy

mole crickets avoid nematode treated soil and how the oviposition behavior of infected

female mole crickets is affected. We wanted to ascertain if mole crickets can sense and

avoid nematodes over long distances (ca. 15 cm) and, when given a choice between

nematodes and pesticides, would the nematode treated areas be preferred over pesticide

treated areas. This information will help in developing a successful IPM program using

S. scapterisci and pesticides, as well as determine ways in which application of S.

scapterisci could be improved or modified to increase infection rates and reduce pesticide

rates.

Materials and Methods

Steinernema scapterisci (Nematac S, Becker Underwood, Ames, IA) used in this

study was stored in a 70C cold room until use (< 3 mo). The viability of the nematodes

was tested before each application by dissolving a pinch of Nematac S into water and

observing nematode shape and mobility under a light microscope. Healthy nematodes

are opaque in color and S-shaped with undulating movements. Dead or unhealthy

nematodes are translucent, straight, and lack movement. The product was used if

viability was >50% and discarded if <50% viable.









Tawny (Scapteriscus vicinus) and southern (Scapteriscus borellii) mole crickets

were collected from pitfall traps or sound traps in Alachua Co., FL, during the fall and

spring of 2003 and 2004 and returned to the laboratory. Each mole cricket was placed in

a 120-ml plastic vial (Thorton Plastics Salt Lake City, UT) with sterilized sand and held

for >14 d to ensure health. Mole crickets were maintained at 230C with a photoperiod of

12:12 (L: D) and fed commercial cricket chow (Purina). Scapteriscus abbreviatus were

obtained from a laboratory colony at the University of Florida Entomology and

Nematology Department and maintained under the same conditions described above.

Nematode Infection and Nematode Treated Areas Effect on Mole Cricket Tunneling
Behavior

Two dimensional tunneling behavior assays were conducted on Scapteriscus

vicinus adults that were either healthy or infected with Steinernema scapterisci. Plexiglas

containers (30.5 cm wide x 30.5 cm high x 2.5 cm deep) were filled with autoclaved

sand, moistened with deionized water, positioned vertically, kept in a dark area to

simulate a subterranean environment, and observed under a red light bulb (40W).

Ryegrass (Lolium multiflorum) was grown on top of the containers as a food source for S.

vicinus. After the grass established a root system approximately 10-15 cm in length, one

adult mole cricket previously exposed to nematodes (0, 500, or 10,000 infective

juveniles) was placed in a randomly selected area at the top of the container. Five males

and five females were tested for each treatment level. A small strip of Plexiglas was

placed in the top opening of the arena to prevent mole cricket escape. Tunnel dimensions

in the sand were measured at 1, 4, 8, 24, 48, 72, and 96 h then every 48 h after that for 10

d or until mole cricket death, whichever occurred first. Tunnel patterns were traced on

acephate paper, and then lengths were measured with a cloth measuring tape. Time that









tunnel construction slowed down or ceased was recorded. Daily tunnel distances were

determined by subtracting total tunnel distance from the previous day's tunnel distance.

Mole crickets were then removed from the arenas and assayed for nematode infection.

There were 10 replicates (crickets) for each treatment.

Additional tests were completed to determine whether healthy mole crickets could

sense and/or avoid nematode-treated soil. Autoclaved builder's sand was colored with

blue-powdered chalk (American Tool Companies, Inc., Wilmington, OH) and placed on

one half of a 30.5 cm wide x 30.5 cm high x 2.5 cm deep Plexiglas arena with the other

half containing sand colored with orange powder (American Tool Companies, Inc.,

Wilmington, OH). Ryegrass was grown on top of the containers to provide food for the

mole crickets. The blue colored sand was inoculated with a label rate (approximately

25,000 nematodes/cm2) of S. scapterisci by injecting approximately 100 ml of nematodes

in solution to one corner of the container using a graduated cylinder. To determine if

there were any deterrent effects of the chalk used to color the sand, control arenas were

setup similar to the treated arenas. Healthy mole crickets were placed in the control arena

with colored sand, however both sides of the arenas lacked nematodes. The controls

were treated exactly as the treatment arenas.

The containers were kept in a dark area to simulate subterranean conditions and

observed under red light (40 W). One mole cricket was placed on the untreated side of

the Plexiglas arena. Half of the replicates contained orange sand treated with nematodes

with untreated blue sand and the other half contained blue sand treated with nematodes

and untreated orange sand. Tunnels and movement of nematode infected sand was

compared to movement of untreated sand by looking at the tunnel patterns and movement









of the treated sand. Mole cricket behavior was observed at 1, 4, 8, 24, 48, 72, and 96 h

then every 48 h after that for 10 d or until mole cricket death, whichever occurred first.

Data were summarized into 48 h increments. The length of the tunnel system was

determined as previously described. After 10 d mole crickets were removed from the

arenas and assayed for nematode infection. There were ten replicates (mole crickets).

Mole crickets used in this study were dissected after 10 d (or after death, whichever

occurred first).

Oviposition Behavior of Mole Crickets Exposed to S. scapterisci

The oviposition behavior including number of eggs, clutch size, depth and width of

egg chambers, and time until oviposition was also observed in the previous tests. After

10 d the arenas were dismantled and egg clutches were removed. An indentation was

made in 55 mm Petri dishes filled with moistened sterilized sand by pressing a gloved

finger into the moistened sand. Eggs were carefully removed using feather-weight

forceps and placed into the depression. Petri dishes were covered and wrapped with

parafilm (Fisher Scientific, Pittsburgh, PA) to keep humidity levels high. Eggs were

monitored every 48 h until egg hatch.

Y-Tube Tests

Observation Chamber

Response of mole crickets to potentially lethal compounds was tested in a dual

choice observation chamber intended to simulate below ground tunnel conditions. Y-

tube arenas were constructed to evaluate the response of Scapteriscus borellii, S. vicinus,

and S. abbreviatus to Steinernema scapterisci or untreated sand, and the choice between

nematodes or pesticides. Arenas were constructed of 1.27 cm diameter plastic tubing

(Fisher Scientific, Pittsburgh, PA), the basal section was 15 cm long, and each arm was









10 cm long. The tubing was connected to form a Y using a 1.27 cm plastic y-connector

(Fisher Scientific, Pittsburgh, PA).

Autoclaved builder's sand was loosely packed in either arm of the Y-tube.

Comparisons consisted of 500 against 0 nematodes. Nematodes were added to the sand

using a pipet. Each arm was capped with a 20 ml plastic scintillation vial (Fisher

Scientific, Pittsburgh, PA). The 15 cm entry arm was left void of sand to facilitate mole

cricket movement. Ten replicates of S. borellii and S. vicinus were completed for each

level of nematodes; each replicate was completed using a different cricket. Experiments

were carried out at 23 C in dark conditions using red light (40W) to observe the mole

crickets. One adult S. borellii or S. vicinus was placed into the basal section of the Y-

tube. The cricket was observed as it moved through the tubing at the base and then chose

between the arms containing treatments. Choice was determined when a cricket traveled

completely through one arm and entered the scintillation vial at the end. Y-tubes were

cleaned with a 1% bleach solution between runs and re-used for nematode studies only.

Response to S. scapterisci or Pesticides

Five insecticides commonly used for pest mole cricket control were evaluated in

the laboratory against nematodes for their attractive or repellent characteristics to mole

crickets. The pesticides tested were acephate (0.91 kg AI/ha) (Orthene Turf, Tree and

Ornamental Spray, Valent Professional Products, Walnut Creek, CA), bifenthrin (224 g

AI/ha) (Talstar GC Flowable, FMC Corporation, Philadelphia, PA), imidacloprid (451 g

AI/ha) (Merit 75 WP, Bayer Environmental Science, Montvale, NJ), fipronil (280 g

AI/ha) (Chipco Choice, Bayer Environmental Science, Montvale, NJ) and deltamethrin

(146 g AI/ha) (DeltaGard T&O, Bayer Environmental Science, Montvale, NJ).

Autoclaved builder's sand was loosely packed in one arm of the Y-tube and 10,000









nematodes were added using a pipet. The second arm contained autoclaved builder's

sand mixed with 1 ml of the label rate of the above pesticides. Each arm was capped

with a 20 ml plastic scintillation vial (Fisher Scientific). The 15 cm basal entry arm was

empty. Seven replicates were completed for each insecticide; each replicate was

conducted using a naive cricket. Experiments were conducted using the same methods

described in the previous Y-tube tests. Y-tube arenas used in pesticide studies were

discarded after use to prevent contamination.

Statistical Analysis

Tunneling and oviposition data were subjected to an ANOVA (SAS Institute 2001)

and means were separated using the Tukey-Kramer Honestly Significant Difference

means separation test. Within each Y-tube test, the number of crickets responding to

each choice was analyzed by a chi-square test against the null hypothesis of a 1:1 ratio

(Steel and Torrie 1981).

Results

Effect of Nematode Infection and Nematode Treated Areas on Mole Cricket
Tunneling Behavior

Tunneling rate of mole crickets treated with 0, 500, 10,000 nematodes did not

significantly differ over time among the treatments (Figure 5-1). Total distance tunneled

over the entire 10 d period (29-44 cm) was not statistically significant among the three

treatments (F= 0.87; df= 2, 35; P= 0.43). Mole crickets treated with 500 or 10,000 S.

scapterisci tended to tunnel less (lower average daily distance) when compared to healthy

mole crickets (Figure 5-2). There were no significant differences between male and

female mole cricket tunneling when exposed to different levels of nematodes (F=1.073;

df= 5, 23; P=0.408). During the 10 d test, the eight mole crickets treated with 0, 500, or









10,000 nematodes had 0, 1, and 3 crickets respectively infected with nematodes. Tunnel

distances began to decrease after 6 d in crickets treated with 500 or 10,000 nematodes.

Mole crickets did not appear to differentiate between untreated and nematode-

treated sand (F= 1.93; df=l1, 79; P= 0.1684). Tunnel length did not differ in either side of

the arena at anytime during the test (Figure 5-3). There were no significant effects on

tunneling or viability of crickets in arenas with sand lacking nematodes and tinted with

orange or blue colored chalk (F=2.94; df=l, 49; P=0.5899). Several mole crickets did

not venture far from the top 10 cm, and these crickets were the first to die. All mole

crickets were infected with S. scapterisci.

Oviposition Behavior of Mole Crickets Exposed to S. scapterisci

Mole crickets infected with 500 or 10,000 nematodes were able to lay eggs, to

about the same soil depth, in the same chamber dimensions, and in a similar quantity to

uninfected females (Table 5-1). These behaviors also did not differ in treated vs.

untreated sand (500 or 10,000 nematodes) (Table 5-2). Healthy mole crickets and those

treated with 10,000 nematodes laid eggs between 1-6 d. Crickets treated with 500

nematodes laid eggs between 1-10 d. One female cricket treated with 500 nematodes

either buried or consumed her clutch of eggs during the study period. None of the eggs

survived due to mold growth.

Y-Tube Tests

None of the three mole cricket species tested demonstrated a significant choice

between arms treated with 500 or 10,000 S. scapterisci (S. borellii, S vicinus, or S.

abbreviatus; y2 = 2.4, 2.1, and 1.6 respectively for 500 nematodes; S. abbreviatus and S.

vicinus; 2 = 1.6, and 2.4, respectively for 10,000 nematodes). Most mole crickets tested

chose the arm that contained S. scapterisci when given a choice between S. scapterisci or









insecticides (Table 5-4). Crickets, when given a choice between S. scapterisci and

fipronil chose the arm containing fipronil. Fipronil granules did not readily dissolve in

water and the solution tended to stay at the very distal end of the arm reducing contact

with the mole cricket.

Discussion

Although mole crickets have been observed avoiding entomopathogens such as

Beauveria bassiana and Metarhizium anisopliae (Villani et al. 1999, Brandenburg 2002,

Villani et al. 2002, Thompson 2004), they do not appear to avoid the insect parasitic

nematode S. scapterisci. In this study we sought to determine if infection of Scapteriscus

spp. with S. scapterisci caused any modification of tunneling behavior. Although the

results were not statistically significant, the crickets infected with 10,000 nematodes

tended to tunnel less than crickets treated with 500 nematodes and healthy crickets. The

shape and length of the tunnels formed by S. borellii concur with Brandenburg et al.

(2002). Both healthy and nematode-infected crickets typically formed an inverted y-

shaped tunnel with branching occurring lower than the surface of the arenas. Our results

show that crickets infected for >6 d may not tunnel as much or as deep as uninfected

mole crickets. This shallow tunneling behavior may make the mole crickets more

vulnerable to insecticides, management practices such as aeration, or increase

vulnerability to predators such as birds and armadillos.

Because insect parasitic nematodes are slower-acting than chemical pesticides the

mole crickets may continue to oviposit during the period of infection before death. Our

results indicate no significant differences in the ovipositional behavior of female crickets.

Females infected with nematodes may not have the energy or resources available for egg

laying because these resources are required for immune system defense and the female









may die before completing oviposition. Research on Beauveria bassiana has involved

the effects of infection on oviposition behavior of Lygus hesperus (Hemiptera: Miridae)

an important crop pest in the western United States (Noma and Strickler 2000). Lygus

hesperus oviposition is significantly reduced after inoculation with B. bassiana in the

laboratory. Nematodes in the genus Steinernema have also resulted in reduced overall

host fecundity (Belair and Boivin 1995, Kim et al. 2004).

Scapteriscus borellii adults laid a similar number of eggs in nematode treated and

untreated sand in the Plexiglas arenas, indicating a lack of repellency by the nematodes.

Thus, a female ovipositing in soil treated with Nematac S may then become infected,

although her offspring are not likely to be immediately affected. We were unable to

assess egg viability in these tests due to excessive mold development (none of the eggs

hatched).

Mole crickets have been observed avoiding insecticides (Brandenburg 2002;

Thompson 2004). If mole crickets can sense and avoid insecticides applied to tees,

greens, and fairways this may potentially force the cricket into the roughs where

nematode applications could be made. In our tests, when given a choice, mole crickets

chose sand treated with S. scapterisci over sand treated with the insecticides tested. The

sensory system of mole crickets is not well understood (Villani et al. 1999; Brandenburg

2002; Villani et al. 2002; Thompson 2004). Initial scanning electron microscope

photographs (Appendix D) indicate many hairs covering the antennae, tarsi, and palps.

These hairs may be due to the insect's subterranean habitat and not for sensory detection.

Sensory detection in mole crickets should be studied further. Knowledge of mole cricket

sensory detection would be useful in development of a mole cricket IPM program.









The fact that Scapteriscus borellii oviposited in a laboratory arena is also new

information. The common rearing method for S. abbreviatus in the University of Florida

colony in Gainesville, FL is in cylindrical 120-ml plastic vials with autoclaved sand.

This method was unsuccessful for the author when rearing S. borellii or S. vicinus;

however oviposition was observed in females placed in the Plexiglas sandwich arenas

used for this study. The larger arenas may provide more room for the crickets to move

around; also females could tunnel down 30.5 cm versus only several centimeters in the

plastic vials. The moisture level also remained relatively high in the Plexiglas arenas.

High humidity is necessary for successful mole cricket oviposition (Hertl et al. 2001).













o 0 nematodes
* 500 nematodes
E 10,000 nematodes


50



I 40



30
c
= 30
I-

Cu
S20
Q


2 4 6 8 10
Day

Figure 5-1. Distance (cm) tunneled ( SEM) through sand in Plexiglas arenas every 2 d
by mole crickets exposed to varying amounts of nematodes. ANOVA for
2,4,6,8, and 10 d are (F=0.45; df=2,34; P=0.64); (F=1.45; df=2,32; P=0.25);
(F=0.32; df=2,31; P=0.73); (F=0.61; df=2,31; P=0.55); (F=1.49; df=2,29;
P=0.24); respectively.













E Female
* Male


I 50
"o

40
I-
U-
30


20
I-


0 I
0 500 10000
Number of Nematodes

Figure 5-2. Comparison of total tunnel distances ( SEM) by male and female
Scapteriscus spp. mole crickets after exposure to S. scapterisci.


35


30

25 -

20 -

15 -

10

5

0
1 5 15 24


* Untreated


Time (h)


Figure 5-3. Distance tunneled ( SEM) by mole crickets through sand treated with
nematodes and untreated sand in Plexiglas arenas over 48 h.


Treated











Table 5-1. Oviposition of Scapteriscus mole crickets directly infected with different
numbers of S. scapterisci.
# Mole Width of egg Depth of egg
Treatment crickets that Number of eggs
(# nematodes) laid eggs (+ SEM) chamber (cm) chamber (cm)
(n=8) SEM) SEM)
0 5 17.25 + 5.36 2.81 + 1.07 12.93 + 4.14
500 6 19.25 5.01 2.74 0.91 11.34 2.71
10,000 4 12.63 + 4.85 1.70 + 0.66 7.99 + 3.26
1F= 0.14; df= 2,14; P= 0.87
2F= 0.35; df= 2,14; P= 0.67
3F= 1.82; df= 2,14; P= 0.20



Table 5-2. Oviposition of Scapteriscus mole crickets in sand treated with S. scapterisci.
# Mole Width of egg Depth of egg
Treatment crickets that Number of eggs chamber (cm) (+ chamber (cm)
laid eggs (+ SEM) 1SE)2 SEM)3
(n= 5)


Treated sand 3 33.5 + 17.99 4.5 0.42 15.5 + 1.
Untreated sand 5 24.0 25.45 5.0 + 0.60 19.5 2.


'F = 0.970; df =
2 F= 52.600; df
3F= 77.933; df=


2, 4; P
=2,4; P
=2,4;P=


89
68


0.506
=0.019
0.013


Table 5-3. Response of Scapteriscus vicinus, S. borellii, and S. abbreviatus to
Steinernema scapterisci nematodes versus sterilized sand.
Species tested # crickets selecting the y-tube arm1
S. scapterisci Sterilized sand %2
S. vicinus 3 7 2.1*
S. borellii 6 4 2.4*
S. abbreviatus 2 8 1.6**
All species combined 11 19 6.3
1Response of 10 individuals of each mole cricket species when given a choice between
two arms. *P<0.01, **P<0.005






60


Table 5-4. Response of Scapteriscus vicinus to Steinernema scapterisci nematodes
versus pesticides treated sand.
Pesticide tested Y-tube arm selected 1

S. scapterisci Insecticide No choice


Acephate
Bifenthrin
Deltamethrin
Fipronil


0


0


Imidacloprid 5 1
Response of 7 mole crickets when given a choice between two arms.
More than 10 replicates are needed for statistical analysis.














CHAPTER 6
SUMMARY AND CONCLUSIONS

Steinernema scapterisci is an insect parasitic nematode that has been formulated by

Becker Underwood as the biopesticide Nematac S. It differs from other commercially-

available nematodes used for mole cricket control by being able to reproduce in the host's

body and persisting in the soil after application. Thus, after being sprayed like a pesticide

to infested turfgrass, the nematodes may continually help to suppress mole crickets as a

natural enemy.

Nematac S was originally marketed for use on pastures because insecticides were

not a cost effective control option. To expand the marketing and use of Nematac S, golf

courses, athletic fields, parks, lawns and other turfgrass areas were also considered. The

research conducted in this dissertation is relevant primarily to the golf course industry,

but results may be extrapolated into the other turfgrass sites.

The intensity of management on Florida golf courses ranges from high on private

courses to somewhat lower on municipal or public courses. Golfers, golf pros, and

superintendents tend to have little tolerance for damage on their greens, tees, and

fairways. Roughs and driving ranges, however, may receive less attention and may

therefore be more ideal areas for introducing or conserving natural enemies. This

research has demonstrated that S. scapterisci can survive on intensively managed golf

courses for more than ten years. Most of the insecticides (e.g., acephate, bifenthrin,

deltamethrin, fipronil, imidacloprid) used against mole cricket nymphs and adults do not

reduce nematode survival or infectivity. But, products used to control plant parasitic









nematodes would undoubtedly also kill beneficial nematodes. Perhaps this particular

problem, which many golf courses deal with, is why insect parasitic nematodes have not

been heavily used on golf courses in the past.

The beneficial effects of S. scapterisci increase in an additive fashion over time.

Steinernema scapterisci can survive and reproduce within the soil and can be transported

to new areas via mole cricket movement; therefore its benefits are seen year after year,

unlike pesticides which need to be reapplied. Golf courses unknowingly benefit from

nematode populations by an overall reduction in mole cricket populations over time. The

reduction in mole cricket populations would then reduce the pesticide applications

required for mole crickets possibly to the level of applying hot spot applications only.

Steinernema scapterisci is an important part of any mole cricket integrated pest

management program.















APPENDIX A
AMBIENT DATA AND TURFGRASS QUALITY DATA COLLECTED AT
GAINESVILLE GOLF AND COUNTRY CLUB AND IRONWOOD GOLF COURSE

Table A-1. Ambient data collected from Gainesville Golf and Country Club on dates of
mole crickets collections.
Air Temp.1 Rel. Humid.1 Soil Temp.1 Rainfall2
Date (0C) (%) (0C) (cm)
October 2001 20.2 68.0 21.1 0.3
November 2001 15.3 60.0 14.6 2.7
December 2001 13.6 75.7 16.1 3.8
January 2002 12.8 64.5 13.3 13.3
February 2002 19.6 72.0 16.4 2.8
March 2002 24.1 80.0 15.3 8.3
April 2002 21.1 74.5 21.1 1.0
May 2002 21.0 79.3 20.4 4.0
July 2002 23.9 68.0 23.3 13.8
August 2002 23.3 82.2 23.9 26.5
September 2002 25.0 68.0 25.6 12.8
October 2002 22.6 74.5 22.5 4.2
1 Average of monthly data taken on each collection date. Soil temp taken at 7.6 cm below
surface.
2 Average monthly rainfall value from data collected at the Gainesville Regional Airport.












Table A-2. Ambient data collected from Ironwood Golf Course on dates of mole crickets
collections. n/a indicates data not collected for that date.
Air Temp.1 Rel. Humid.1 Soil Temp1. Rainfall2
Date (OC) (%) (OC) (cm)
October 2001 24.9 61.3 23.7 0.3
November 2001 18.9 64.8 16.1 2.7
December 2001 15.0 80.4 15.0 3.8
January 2002 n/a n/a 13.1 13.3
February 2002 17.9 84.0 16.7 2.8
March 2002 16.8 66.5 16.1 8.3
April 2002 23.9 72.6 22.2 1.0
May 2002 18.5 94.3 20.9 4.0
July 2002 23.3 93.0 23.3 13.8
August 2002 24.2 74.0 24.6 26.5
September 2002 25.1 56.0 23.3 12.8
October 2002 22.6 60.5 24.0 4.2
November 2002 11.7 78.5 11.1 13.3
December 2002 13.3 93.0 10.0 18.3
January 2003 3.1 68.0 7.22 0.5
February 2003 10.6 60.0 5.0 16.8
March 2003 21.1 68.5 17.2 19.5
April 2003 16.2 76.0 17.2 4.1
May 2003 19.3 74.0 20.6 5.1
1 Average of monthly data taken on each collection date. Soil temp taken at 7.6 cm below
surface.
2 Average monthly rainfall value from data collected at the Gainesville Regional Airport.










Table A-3. Average turfgrass density ratings for treated and untreated plots on
Gainesville Golf and Country Club and Ironwood Golf Course. l=poor;
5=acceptable; 9=excellent. n/a indicates data not taken.
Gainesville Golf and Ironwood Golf Course
Country Club
Collection Date treated untreated treated untreated
plots plots plots plots
October 2001 5.7 5.2 5.0 3.7
November 2001 7.7 7.6 6.2 4.4
December 2001 6.7 6.3 6.0 4.9
January 2002 5.7 5.7 6.0 4.8
February 2002 6.1 6.2 5.9 4.8
March 2002 6.8 5.9 5.0 4.3
April 2002 6.3 5.6 4.8 3.7
May 2002 6.5 5.4 5.4 3.8
July 2002 8.7 6.8 6.4 5.5
August 2002 5.3 5.5 5.9 5.1
September 2002 5.8 6.3 5.7 5.0
October 2002 4.9 4.6 5.6 5.0
November 2002 n/a n/a 5.2 5.1
December 2002 n/a n/a 6.2 4.8
January 2003 n/a n/a 5.3 3.4
February 2003 n/a n/a 5.5 4.6
March 2003 n/a n/a 4.6 3.6
April 2003 n/a n/a 4.6 3.6
May 2003 n/a n/a 4.5 3.4














APPENDIX B
DATA FROM ATHLETIC FIELD DEMONSTRATION SITES

Materials and Methods

Objective

A demonstration project was conducted on athletic field turfgrass. Methods were

similar to those in Chapter 2 of this dissertation; however fewer replicates were used due

to limited space. Nematodes are an important part of an athletic field IPM program

because they are a non-chemical way of reducing mole cricket populations. This in turn

reduces the exposure to children who use these fields.

Study Site

The establishment and spread of S. scapterisci was monitored on two baseball

fields located in Sarasota, FL (Twin Lakes Park) and Altamonte Springs, FL (Eastmonte

Park). Both baseball fields were open to the public. The Sarasota field was built around

1975 and the Altamonte Springs field was built around 1970. The Sarasota field was

common bermudagrass (Cynodon dactylon [L.] Pers.) and the Altamonte Springs field

was bermudagrass (Cynodon dactylon x C. transvaalensis) var. Tifway. Both parks were

mowed at 1.91 cm. Neither park had previously received Steinernema scapterisci

treatments.

Mole Cricket Monitoring

Three hot spots of mole cricket activity were located in each baseball outfield.

Linear pitfall traps (modified from Lawrence 1982, described in Chapter 2) were installed









in the ground at least 100 m apart. Each field contained two treated areas and one control

area. Traps on all sites were installed in September 2001.

Nematodes were released in the morning (approximately 0700). Nematodes were

applied in an aqueous suspension of 1 billion nematodes/ 378.5 L of water applied using

a boom sprayer calibrated at 40 L/1000 m2. The area treated was 20.1 x 20.1 m (404.7

m2) around each trap. Treated areas were irrigated with 0.6 cm of water before and 0.6

cm after application. The pre-treatment dates for Sarasota were 11,18 October 2001 and

11 October 2001 for Altamonte Springs. Nematodes were applied on 11 November 2001

at Sarasota and 25 October 2001 at Altamonte Springs. A second application of

nematodes was made on the Sarasota field 11 October 2002.

Pitfall traps were sampled to monitor infection levels and mole cricket abundance

using methods similar to Parkman et al. (1993a,b). At each 24 h sampling period the

buckets and arms were cleaned out and 3 to 5 cm of sand was placed into the inner

bucket. Traps were left for 24 h and all live mole crickets were collected. Mole crickets

were placed individually in 20 ml plastic scintillation vials (Fisher Scientific) with 1-2

drops of deionized water, capped and labeled and mailed via overnight courier delivery to

Gainesville, FL. The nematode doesn't move much in the soil and can't be identified

through soil samples, so adult and juvenile mole crickets with pronotal lengths > 4 mm

(Hudson and Nguyen 1989a) were collected from traps, using the same methods as in the

pretreatment sampling, and tested for infection weekly for the first 6 weeks post-

application and one to two times a month thereafter for 2 yr. Turf quality (density, color)

in the area immediately surrounding the pitfall traps was assessed (1 to 9 scale with 1 =

sparse or brown grass, 9 =dense or dark green grass). A second application of S.









scapterisci was made on 11 October 2002 at Sarasota due to low levels of infection in

collected mole crickets. The percentage of infected mole crickets was determined at 7

and 10 d after death under a dissecting scope (10 X). Steinernema scapterisci were

identified by Dr. Khuong Nguyen, Entomology and Nematology Department, University

of Florida.

Results and Discussion

It took approximately 8 wk for infection levels to reach 25% in the mole crickets

collected at the Sarasota site. Levels remained at or near this for about 1 yr post-initial

application. After the second application in October 2002 levels reached 25% (Figure B-

1) and remained at this level in the following spring mole cricket population (March and

April 2003). The baseball field had heavy rains and flooding as well as a non-functional

irrigation system in 2002. Infection level may have been higher if these factors had not

occurred. Very low levels of mole cricket damage/activity were observed in spring

through fall 2003 when compared to previous years. Levels of infection were also low in

the summer months due to the small size of the crickets present during these months (see

Chapter 2). A second application of nematodes may be necessary to reestablish

nematode populations high enough in the soil to result in infection levels of 15-30% in

collected mole crickets.

Data from the Altamonte Springs site was terminated early due to the very low

numbers of crickets (< 20) collected during the research period. The study on this site

was canceled in May 2002. Statistics presented in both graphs are comparisons of

monthly means (SAS Institute 2001).











Sarasota Athletic Field


40
35
30
25
on


/U
15
10
5
0


Month


Figure B-1. Average monthly infection rates at the Sarasota athletic field research site. Arrows indicate Steinernema scapterisci
applications. F=3.1212; df=24,92; P=0.0001


-T

t


I-^














Altamonte Springs


OCT 01 NOV 01 DEC 01 JAN 02 FEB 02 MAR 02 APR 02


MAY 02


Month


Figure B-2. Average monthly infection rates at the Altamonte Springs athletic field research site. Arrow indicates Steinernema
scapterisci applications. F=9.35; df=6,15; P=0.002














APPENDIX C
PRELIMINARY CHECKLIST OF ARTHROPODS ASSOCIATED WITH GOLF
COURSE TURFGRASS

Below is a preliminary checklist of arthropods found in linear pitfall traps located

on Gainesville Golf and Country Club and Ironwood Golf Course in Gainesville Florida

from November 2001 through October 2003. List is considered preliminary and

incomplete due to limited time in which the identifications could be completed.

Class Order Family: subfamily Genus: species
Insecta Coleoptera Carabidae Omophron labiatum
Pasimachus spp.
Curculionidae Sphenophorus spp.
Elateridae 3 A'/ itihn, squamiger
Geotrupidae Peltotrupes profundus
Histeridae
Lampyridae
Ptiliidae
Scarabaeidae Ataenius spp.
Cotinis nitida
Cyclocephala spp.
Phylophaga spp.
Tomarus subtropicus
Silvanidae
Staphylinidae
Tenebrionidae
Tenebrionidae:Diaperinae
Tenebrionidae:Alleculinae
Dermaptera Labiduridae Labidura riparia
Forficula auricularia
Lepidoptera Geometridae Mocis spp.
Noctuidae Herpetogramma phaeopteralis
Pyralidae Spodoptera frugiperda
Saturnidae
Arachnida Agelenidae
Anyphaenidae
Corinnidae
Dictynidae









Class Order Family: subfamily Genus: species
Arachnida Gnaphosidae
Hahniidae
Lycosidae
Mimetidae
Mysmenidae
Pisauidae
Tetrognathidae
Theridiidae
Zorocratidae














APPENDIX D
SCANNING ELECTRON MICROGRAPH PICTURES OF MOLE CRICKET
SENSORY AREAS

Scanning electron micrograph photographs were taken using a tungsten low

vacuum scanning electron microscope model JSM-5510LV (JEOL-USA, Peabody, MA)

to determine if any significant sensory hairs, pores, or other morphological aspects were

apparent on mole crickets. Pictures were taken of dried, curated mole crickets focusing

on the antenna of an adult male and one adult female, a female mid-tarsomere, and a

female labial palpomere. These photographs are only an initial look at mole cricket

sensory organs and more in-depth photographs should be taken for future study.





















Figure D-1. SEM photograph of a female Scapteriscus vicinus antennal mid-section.



























Figure D-2. SEM photograph of a male Scapteriscus vicinus antennal mid-section.


Figure D-3. SEM photograph of a female Scapteriscus vicinus mid-tarsal claw.



























Figure D-4. SEM photograph of a female Scapteriscus vicinus labial palpomere.















LIST OF REFERENCES


Adeji, M. B., J. H. Frank, and C. S. Gardner. 2003. Survey of pest mole cricket
(Orthoptera: Gryllotalpidae) activity in South-Central Florida. Florida Entomol. 86:
199-205.

Akhurst, R. J. 1990. Safety to nontarget invertebrates of nematodes of economically
important pests, pp. 233-240. In M. Laird, L. A. Lacey, and E. W. Davidson (eds.),
Safety of microbial insecticides. CRC Press, Boca Raton, FL.

Ames, L. M. 1990. The role of some abiotic factors in the survival and motility of
Steinernema scapterisci. M. S. Thesis. University of Florida, Gainesville, FL.

Arthurs, S. and K. M. Heinz. 2003. Thrips parasitic nematode Thripinema nicklewoodi
(Tylenchida: Allantonematidae) reduces feeding, reproductive fitness, and
tospsvirus transmission by its host, Frankliniella occidentalis (Thysanoptera:
Thripidae). Environ. Entomol. 32: 853-858.

Bedding, R. A. 1993. Biological control of Sirex noctilio using the nematode Deladenus
siricidicola. pp. 11-20. In R. Bedding, R. Akhurst, and H. Kaya (eds.), Nematodes
and the biological control of insect pests. CSIRO Publications, East Melbourne.

Belair, G. and G. Boivin. 1995. Evaluation of Steinernema carpocapsae Weiser for
control of carrot weevil adults, Listronotus oregonensis (LeConte) (Coleoptera:
Curculionidae), in organically grown carrots. Biocon. Sci. Tech. 5: 225-231.

Benton, M. J. and G. Pritchard. 1990. Mayfly locomotory responses to endoparasitic
infection and predator presence: the effects on predator encounter rate. Freshwater
Biol. 23: 363-371.

Boucias, D. G. 1985. Diseases. pp 3-10. In T.J. Walker (ed.), Mole crickets in Florida.
Florida Agric. Exp. Sta. Bull. 846.

Braman, S. K. 1993. Azadirachtin affects growth and survival of immature tawny mole
crickets (Orthoptera: Gryllotalpidae). Florida Entomol. 76: 526-530.

Braman, S. K., R. R. Duncan, W. W. Hanna, and W. G. Hudson. 2000. Evaluation of
turfgrasses for resistance to mole crickets (Orthoptera: Gryllotalpidae). HortScience
35: 665-668.

Brandenburg, R. L. 2002. Improving mole cricket management by targeting their
weaknesses. USGA Turf. and Environ. Res. 1: 1-8.









Brandenburg, R. L., Y. Xia, and A. S. Schoeman. 2002. Tunnel architectures of three
species of mole crickets (Orthoptera: Gryllotalpidae). Florida Entomol. 85: 383-
385.

Capinera, J. L. and N. C. Leppla. 2001. Pest Mole Crickets.
http://creatures.ifas.ufl.edu/ Accessed 12 February 2002.

Cobb, P. P. and T. P. Mack. 1989. A rating system for evaluating tawny mole cricket,
Scapteriscus vicinus Scudder, damage (Orthoptera: Gryllotalpidae). J. Entomol.
Sci. 24: 142-144.

Creighton, C. S. and G. Fassuliotis. 1985. Heterorhabditis sp. (Nematoda:
Heterorhabditidae): a nematode parasite isolated from the banded cucumber beetle
Diabrotica balteata. J. of Nematol. 17: 150-153.

Das, J. N. and B. J. Divakar. 1987. Compatibility of certain pesticides with DD-136
nematode. Plant Prot. Bull. 39: 20-21.

Denisenko, M. K. 1986. An effective method of controlling mole crickets. Zaschchita
Rastenii 10: 51.

Dutky, S. R. 1974. Nematode parasites. pp. 576-590. In F. G. Maxwell and F. A. Harris
(eds.), Proceedings of the Summer Institute on Biological Control of Plant Insects
and Diseases. Univ. Press of Mississippi, Jackson.

English-Loeb, G., M. Villani, T. Martinson, A. Forsline, and N. Consolie. 1999. Use
of entomopathogenic nematodes for control of grape phylloxera (Homoptera:
Phylloxeridae): a laboratory evaluation. Environ. Entomol 28: 890-894.

Epsky, N. D. and J. L. Capinera. 1994. Invasion efficiency as a measure of the efficacy
of the entomogenous nematode Steinernema carpocapsae (Rhabditida:
Steinernematidae). J. Econ. Entomol. 87: 366-370.

Forrest, T. G. 1985. Reproductive behavior. pp 3-10. In T. J. Walker (ed.), Mole crickets
in Florida. Florida Agric. Exp. Sta. Bull. 846.

Forschler, B. T., J. N. All, and W. A. Gardner. 1990. Steinernemafeltiae activity and
infectivity in response to herbicide exposure in aqueous and soil environments. J.
Inv. Path. 55: 375-379.

Frank, J. H. 1990. Mole crickets and other arthropod pests of turf and pasture. pp. 131-
139. In D. H. Habeck, F. D. Bennett, and J. H. Frank (eds.), Classical biological
control of insects and weeds in the southern United States. South. Coop. Ser. Bull.
355.

Frank, J. H. 2001. Statewide controls for mole crickets? Florida Turf Digest.
July/August 2001.









Frank, J. H., E. A. Buss, and K. A. Barbara. 2002. Compound interest beneficial
nematodes in turf: good for how many years against pest mole crickets? Florida
Turf Digest 19: 48-50.

Frank, J. H., T. R. Fasulo and D.E. Short. 1998. MCricket Knowledgebase. University
of Florida, Institute of Food and Agricultural Sciences. Available:
http://molecrickets.ifas.ufl.edu/index.htm Accessed: 6 December 2001.

Frank, J. H. and J. P. Parkman. 1999. Integrated pest management of pest mole
crickets with emphasis on the southeastern USA. Integrated Pest Management
Reviews 4: 39-52.

Frank, J. H., J. P. Parkman and F. D. Bennett. 1995. Larra bicolor (Hymenoptera:
Sphecidae), a biological control agent of Scapteriscus mole crickets (Orthoptera:
Gryllotalpidae) established in northern Florida. Fla. Entomol. 78: 619-623.

Frank, J.H., T.J. Walker and J.P. Parkman. 1996. The introduction, establishment,
and spread of Ormia depleta in Florida. Biol. Control 6: 368-377.

Gaugler, R. and G. M. Boush. 1978. Effects of ultraviolet radiation and sunlight on the
entomogenous nematode, Neoplectana carpocapsae. J. Inv. Pathol. 32:291-296.

Gaugler, R. and H. Kaya (eds.). 1990. Entomopathogenic nematodes in biological
control. CRC, Boca Raton, FL.

Georgis, R. 1997. Commercial prospects of microbial insecticides in agriculture. Proc.
Br. Crop. Prot. Council Symp. 68: 243-252.

Georgis, R. and R. Gaugler. 1991. Predictability in biological control using
entomopathogenic nematodes. J. Econ. Entomol. 84: 713-720.

Georgis, R., and H. K. Kaya. 1998. Formulation of entomopathogenic nematodes. pp.
289-308. In H. D. Burges (ed.), Formulation of microbial biopesticides. Kluwer
Academic, Dordrecht.

Georgis, R. and S. A. Manweiler. 1994. Entomopathogenic nematodes: a developing
biological control technology. Vol. 6, pp. 63-94. In Agricultural Zoology Reviews,
K. Evans, (ed.) Intercept, Andover.

Gordon, R., J. Chippett, and J. Tilley. 1996. Effects of two carbamates on infective
juveniles of Steinernema carpocapsae All strain and Steinernemafeltiae Umed
strain. J. Nematol. 28: 310-317.

Hanna, W. W., S. K. Braman, and W. G. Hudson. 2001. Bermudagrass hybrids just
say 'no' to mole crickets. Golf Course Mgmt. 69: 49-51.

Hanna, W. W., and W. G. Hudson. 1997. Genetic resistance to mole crickets in turf
bermudagrass. Turfgrass Trends 6: 17-18.









Hara, A. H. and H. K. Kaya. 1982. Effect of selected insecticides and nematicides on
the in vitro development of the entomogenous nematode, Neoplectana
carpocapsae. J. Nematol. 14: 486-491.

Hara, A. H. and H. K. Kaya. 1983a. Toxicity of selected organophosphate and
carbamate pesticides to infective juveniles of the entomogenous nematode,
Neoplectana carpocapsae (Rhabditida: Steinernematidae). Environ. Entomol. 12:
496-501.

Hara, A. H. and H. K. Kaya. 1983b. Development of the entomogenous nematode,
Neoplectana carpocapsae (Rhabditida: Steinernematidae), on insecticide-killed
beet armyworm (Lepidoptera: Noctuidae). J. Econ. Entomol. 76: 423-426.

Head, J., K. F. A. Walters, and S. Langton. 2000. The compatibility of the
entomopathogenic nematode, Steinernemafeltiae, and chemical insecticides for the
control of the South American leafminer, Liriomyza huidobrensis. BioControl 45:
345-353.

Hertl, P. T., R. L. Brandenburg, M. E. Barbercheck. 2001. Effect of soil moisture on
ovipositional behavior in the southern mole cricket (Orthoptera: Gryllotalpidae).
Environ. Entomol. 30: 466-473.

Hilliard, M. A., C. Bergamasco, S. Arbucci, R. H. A. Plasterk, and P. Bazzicalupo.
2004. Worms taste bitter: ASH neurons, QUI-1, GPA-3 and ODR-3 mediate
quinine avoidance in Caenorhabditis elegans. The EMBO Journal 23: 1101-1111.

Hudson, W. G. 1985. Other behavior, damage, and sampling. pp. 16-21. In T. J. Walker
(ed.), Mole crickets in Florida. Florida Agric. Exp. Sta. Bull. 846.

Hudson, W. G. 1986. Mole crickets (Orthoptera: Gryllotalpidae) damage to Hemarthria
altissima: resistance or nonpreference? J. Econ. Entomol. 79: 961-963.

Hudson, W. G. 1988. Field sampling for mole crickets (Orthoptera: Gryllotalpidae:
Scapteriscus): a comparison of techniques. Florida Entomol. 71: 214-216.

Hudson, W. G. 1989. Field sampling and population estimation of the tawny mole
cricket (Orthoptera: Gryllotalpidae). Florida Entomol. 72: 337-343.

Hudson, W. G., S. K. Braman, R. D. Oetting and B. L. Sparks. 1997. Ornamental,
lawn and turf insects. pp. 21-23. In D.G. Riley, G.K. Douce and R.M. McPherson
(eds.). Summary of losses from insect damage and costs of control in Georgia
1996, Georgia Agric. Exp. Sta. Spec. Publ. 91.

Hudson, W. G., J. H. Frank and J. L. Castner. 1988. Biological control of
Scapteriscus spp. mole crickets (Orthoptera: Gryllotalpidae) in Florida. Bull.
Entomol. Soc. Am. 34: 193-198.









Hudson, W. G. and K. B. Nguyen. 1989a. Infection of Scapteriscus vicinus (Orthoptera:
Gryllotalpidae) nymphs by Neoplectana Spp. (Rhabdita: Steinernematidae). Florida
Entomol. 72: 383-384.

Hudson, W. G. and K. B. Nguyen. 1989b. Effects of soil moisture, exposure time,
nematode age, and nematode density on laboratory infection of Scapteriscus
vicinus and S. acletus (Orthoptera: Gryllotalpidae) by Neoplectana sp. (Rhabditida:
Steinernematidae). Environ. Entomol. 18: 719-722.

Ishibashi, N., D. R. Choi, and E. Kondo. 1987. Integrated control of insects/nematodes
by mixing applications of steinernematid nematodes and chemicals. J. Nematol. 19:
531.

Ishibashi, N. and S. Takii. 1993. Effects of insecticides on movement, nictation, and
infectivity of Steinernema carpocapsae. J. Nematol. 25: 204-213.

Jansson, R. K., S. H. Lecrone and R. Gaugler. 1993. Field efficacy and persistence of
entomopathenogenic nematodes (Rhabditida: Steinernematidae, Heterorhabditidae)
for control of sweet potato weevil (Coleoptera: Apionidae) in southern Florida. J.
Econ. Entomol. 86: 1055-1063.

Jaworska, M., J. Sepiol, and P. A. D. Tomasik. 1994. Effect of metal ions on
Steinernema carpocapsae nematodes. pp. 227. In A. M., R. U. Ehlers, and J. P.
Masson (eds.),Genetics of entomopathogenic nematodes- Bacterium complexes.
Burnell, European Commission, Directorate-General XII, Science Research
Programme, Luxembourg, Belgium,.

Kaya, H. K. 1990. Soil ecology, pp. 189-198. In R. Gaugler and H.K. Kaya (eds.),
Entomopathenogenic nematodes in biological control. CRC, Boca Raton, FL.

Kaya, H. K., T. M. Burlando, H. Y. Choo, and G. S. Thurston. 1995. Integration of
entomopathogenic nematodes with Bacillus id//l i/'gie//,i, or pesticidal soap for
control of insect pests. Biol. Contr. 5: 432-441.

Kaya, H. K. and R. Gaugler. 1993. Entomopathenogenic nematodes. Annu. Rev.
Entomol. 38: 181-206.

Kaya, H. K. and S. P. Stock. 1997. Techniques in insect nematology. pp 281-324. In:
Lacey, L. A, (ed.) Techniques in insect pathology. London, Academic Press.

Kim, H. H., H. Y. Choo, H. K. Kaya, D. W. Lee, S. M. Lee, H. Y. Jeon. 2004.
Steinernema carpocapsae (Rhabditida: Steinernematidae) as a biological control
agent against the fungus gnat Bradysia agrestis (Diptera: Sciaridae) in propagation
houses. Biocon. Sci. Tech. 14: 171-183.

Klein, M. G. and R. Georgis. 1992. Persistence of control of Japanese beetle
(Coleoptera: Scarabaeidae) larvae with steinernematid and heterorhabditid
nematodes. J. Econ. Entomol. 85: 727-730.









Koppenh6fer, A. M. 2000. Nematodes. pp. 283-301. In L. A. Lacey and H. K. Kaya
(eds.), Field manual of techniques in invertebrate pathology: application and
evaluation of pathogens for control of insects and other invertebrate pests. Kluwer
Academic, Dordrecht.

Koppenh6fer, A. M., I. M. Brown, R. Gaugler, P. S. Grewal, H. K. Kaya, and M. G.
Klein. 2000a. Synergism of entomopathogenic nematodes and imidacloprid against
white grubs: greenhouse and field evaluation. Biol. Control 19: 245-251.

Koppenh6fer, A. M., R. S. Cowles, E. A. Cowles, E. M. Fuzy, and L. Baumgartner.
2002. Comparison of neonicotinoid insecticides as synergists for entomopathogenic
nematodes. Biol. Control 24: 90-97.

Koppenh6fer, A. M. and E. M. Fuzy. 2003. Effects of turfgrass endophytes
(Clavicipitaceae: Ascomycetes) on white grub (Coleoptera: Scarabaeidae) control
by the entomopathogenic nematode Heterorhabditis bacteriophora (Rhabditida:
Heterorhabditidae). Environ. Entomol. 32: 392-396.

Koppenh6fer, A. M., P. A. Grewal, and H. K. Kaya. 2000b. Synergism of
imidacloprid and entomopathogenic nematodes against white grubs: the
mechanism. Entomol. Exp. App. 94: 283-293.

Koppenh6fer, A. M. and H. K. Kaya. 1998. Synergism of imidacloprid and an
entomopathogenic nematode: a novel approach to white grub (Coleoptera:
Scarabaeidae) control in turfgrass. J. Econ. Entomol. 91: 618-623.

Krishnayya, P. V. and P. S. Grewal. 2002. Effect of neem and selected fungicides on
viability and virulence of the entomopathogenic nematode Steinernemafeltiae.
Biocon. Sci. Tech. 12: 259-266.

Lacey, L. A., R. Frutos, H. K. Kaya and P. Vail. 2001. Insect pathogens as biological
control agents: do they have a future? Biol. Control 21: 230-248.

Lawrence, K. 0. 1982. A linear pitfall trap for mole crickets and other soil arthropods.
Florida Entomol. 65: 376-377.

Maeyama, T., M. Terayama and T. Matsumoto. 1994. The abnormal behavior of
Colobopsis sp. (Hymenoptera: Formicidae) parasitized by Mermis (Nematoda) in
Papua New Guinea. Sociobiol. 24: 115-119.

Matheny, Jr., E. L. 1981. Contrasting feeding habits of pest mole cricket species. J.
Econ. Entomol. 74: 444-445.

Matheny, Jr., E. L., A. Tsedeke and B. J. Smittle. 1981. Feeding response of mole
cricket nymphs (Orthoptera: Gryllotalpidae: Scapteriscus) to radiolabeled grasses
with, and without, alternative foods available. J. Georgia Entomol. Soc. 16: 492-
495.









Milner, R. J. and J. A. Staples. 1996. Biological control of termites: results and
experiences within a CSIRO project in Australia. Biocontrol. Sci. Tech. 6: 3-9.

Molyneux, A. S. 1985. Survival of infective juveniles ofHeterorhabditis spp., and
Steinernema spp. (Nematoda: Rhabditida) at various temperatures and their
subsequent infectivity for insects. Revue Nematol. 8: 165-170.

Molyneux, A. S. and R. A. Bedding. 1984. Influence of soil texture and moisture on the
infectivity of Heterorhabditis sp. Dl and Steinernema glaseri for larvae of the
sheep blowfly, Lucilia cuprina. Nematologica 30: 359-365.

Nguyen, K. B. 1988. A new nematode parasite of mole crickets: its taxonomy, biology
and potential for biological control. University of Florida Ph.D. Dissertation,
Gainesville, FL.

Nguyen, K. B. and G. C. Smart, Jr. 1990a. Preliminary studies on survival of
Steinernema scapterisci in soil. Soil and Crop Sci. Soc. Florida, Proc. 49: 230-233.

Nguyen, K. B. and G. C. Smart, Jr. 1990b. Steinernema scapterisci n. sp. (Rhabditida:
Steinernematidae) J. Nematol. 22:187-199.

Nguyen, K. B. and G. C. Smart, Jr. 1991. Mode of entry and sites of development of
Steinernema scaptersici in mole crickets. J. Nematol. 23: 267-268.

Noma, T. and K. Strickler. 2000. Effects of Beauveria bassiana on Lygus hesperus
(Hemiptera: Miridae) feeding and oviposition. Environ. Entomol. 29: 394-402.

Parkman, J. P. and J. H. Frank. 1992. Infection of sound-trapped mole crickets,
Scapteriscus spp. by Steinernema scapterisci. Florida Entomol. 75: 163-165.

Parkman, J. P. and J. H. Frank. 1993. Use of a sound trap to inoculate Steinernema
scapterisci (Rhabditida: Steinernematidae) into pest mole cricket populations
(Orthoptera: Gryllotalpidae). Florida Entomol. 76:75-82.

Parkman, J. P. and J. H. Frank. 1996. Effects of fenxoycarb on reproduction of the
shortwinged mole cricket, Scapteriscus abbreviatus (Orthoptera: Gryllotalpidae).
Florida Entomol. 79: 87-91.

Parkman, J. P. and J. H. Frank. 1998. Development and reproduction of mole crickets
(Orthoptera: Gryllotalpidae) after treatments with hydroprene and pyriproxyfen. J.
Econ. Entomol. 91: 392-397.

Parkman, J. P., J. H. Frank, K. B. Nguyen, and G. C. Smart. 1993a. Dispersal of
Steinernema scapterisci (Rhabdita: Steinernematidae) after inoculative applications
for mole cricket (Orthoptera: Gryllotalpidae) control in pastures. Biol. Control 3:
226-232.









Parkman, J. P., J. H. Frank, K. B. Nguyen, and G. C. Smart, Jr. 1994. Innoculative
release of Steinernema scapterisci (Rhabdita: Steinernematidae) to suppress pest
mole crickets (Orthoptera: Gryllotalpidae) on golf courses. Environ. Entomol. 23:
1331-1337.

Parkman, J. P., J. H. Frank, T. J. Walker and D. J. Schuster. 1996. Classical
biological control of Scapteriscus spp. (Orthoptera: Gryllotalpidae) in Florida.
Environ. Entomol. 25: 1415-1420.

Parkman, J. P., W. G. Hudson, J. H. Frank, K. B. Nguyen, and G. C. Smart, Jr.
1993b. Establishment and persistence of Steinernema scapterisci (Rhabdita:
Steinernematidae) in field populations of Scapteriscus spp. mole crickets
(Orthoptera: Gryllotalpidae). J. Entomol. Sci. 28:182-190.

Parkman, J. P. and G. C. Smart. 1996. Entomopathogenic nematodes, a case study:
introduction of Steinernema scapterisci in Florida. Biocontrl. Sci. and Tech. 6: 413-
419.

Patterson-Stark, J. E. and L. A. Lacey. 1999. Susceptibility of western cherry fruit fly
(Diptera: Tachinidae) to five species of entomopathogenic nematodes in laboratory
studies. J. Invertebr. Pathol. 74: 206-208.

Pendland, J. C. and D. G. Boucias. 1987. The hyphomycete Sorosporella-
Syngliocladium from mole cricket, Scapteriscus vicinus. Mycopathologia 99: 25-
30.

Petersen, J. J. 1985. Nematode parasites. pp. 110-122. In H. C. Chapman (ed.),
Biological control of mosquitoes. Am. Mosq. Control Assoc. Bull. 6.

Poinar, G. 0. 1979. Nematodes for biological control of insects. CRC Press, Boca
Raton, FL.

Poinar, G. 0. 1990. Biology and taxonomy of Steinernematidae and Heterorhabditidae,
pp. 23-62. In R. Gaugler and H. K. Kaya (eds.), Entomopathogenic nematodes in
biological control. CRC, Boca Raton, FL.

Poulin, R. 1994. Meta-analysis of parasite-induced behavioral changes. Animal Behav.
48: 137-146.

Poulin, R. 1995. "Adaptive" changes in the behaviour of parasitized animals: a critical
review. Int. J. for Parasitol. 25: 1371-1383.

Poulin, R. and D. M. Latham. 2002. Parasitism and the burrowing depth of the beach
hopper Talorchestia quoyana (Amphipoda: Talitridae). Animal Behav. 63: 269-
275.









Prakasa, R., P. K. Das, and G. Padhi. 1975. Note on compatibility of DD-136
(Neoplectana dutkyi), an insect parasitic nematode with some insecticides and
fertilizers. Indian J. Agric. Sci. 45: 275-277.

Reinert, J. A. and P. Busey. 2001. Host resistance to tawny mole cricket, Scapteriscus
vicinus, in bermudagrass, Cynodon spp. Inter. Turfgrass Soc. Res. Journal 9: 793-
797.

Rovesti, L. and K. V. Dese6. 1990. Compatibility of chemical pesticides with the
entomopathogenic nematodes, Steinernema carpocapsae Weiser and S. feltiae
Filipjev (Nematoda: Steinemematidae). Nematologica 36: 237-245.

SAS Institute. 2001. JMP Start Statisitics User's Guide. Second ed., SAS Inst. Inc.,
Cary, NC.

Schroeder, P. C., M. G. Villani, C. S. Ferguson, J. P. Nyrop and E. J. Shields. 1993.
Behavioral interactions between Japanese beetle (Coleoptera: Scarabaeidae) grubs
and an entomopathogenic nematode (Nematoda: Heterorhabditidae) within turf
microcosms. Environ. Entomol. 22: 595-600.

Schuster, D. J. and J. F. Price. 1992. Seedling feeding damage and preference of
Scapteriscus spp. mole crickets (Orthoptera: Gryllotalpidae) associated with
horticultural crops in west-central Florida. Florida Entomol. 75: 115-119.

Shapiro, D. I. and E. E. Lewis. 1999. Comparison of entomopathogenic nematodes
infectivity from infected hosts versus aqueous suspension. Environ. Entomol. 28:
907-911.

Shapiro-Ilan, D.I., 2001. Virulence of entomopathogenic nematodes to pecan weevil
larvae, Curculio caryae (Coleoptera: Curculionidae), in the laboratory. J. Econ.
Entomol. 94: 7-13.

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

Sithole, C. E. 1986. Mole cricket (Gryllotalpa africana). Zimbabwe Agric. J. 83: 21-22.

Smith, K. 1999. Factors affecting efficacy, pp. 37-46. In S. Polavarapu (ed.),
Proceedings, Optimal Use of Insecticidal Nematodes in Pest Management, 28-30
August 1999, New Brunswick, NJ.

Spinney, L. 1995. High-scoring crickets return for a second inning. New Scientist 146: 6.

Steel, R. G. D. and J. H. Torrie. 1980. Principles and procedures of statistics: a
biometrical approach. 2nd ed. McGraw-Hill, New York.









Thompson, S. R. 2004. Tunneling responses of mole crickets (Orthoptera:
Gryllotalpidae) to the entomopathogenic fungus, Beauveria bassiana (Balsamo)
Vuillemin. Masters Thesis, North Carolina State University. Raleigh, NC.

Thurston, G. S., W. N. Yule, and G. B. Dunphy. 1994. Explanations for the low
susceptibility of Leptinotarsa decemlineata to Steinernema carpocapsae. Biol.
Control 4: 53-58.

Ulagaraj, S. M. 1974. Mole crickets: ecology, behavior, and acoustical communication
(Orthoptera: Gryllotalpidae: Scapteriscus) Ph.D. Dissertation, Univ. Florida,
Gainesville. 72 pp.

Ulagaraj, S. M. 1975. Mole crickets: ecology, behavior, and dispersal flight (Orthoptera:
Gryllotalpidae: Scapteriscus). J. Econ. Entomol. 4: 265-273.

Vance, S. A. 1996. The effect of the mermithid parasite Gasteromermis sp. (Nematoda:
Mermithidae) on the drift behavior of its mayfly host, Baetis bicaudatus
(Ephemeroptera: Baetidae): a trade-off between avoiding predators and locating
food. Can. J. Zool. 74: 1907-1913.

Vaughn, C.C., S.M. Glenn and I. Butler. 1993. Characterization of prairie mole cricket
chorusing sites in Oklahoma. Am. Midl. Nat 130: 364-71.

Villani, M. G., L. L. Allee, A. Diaz, and P. S. Robbins. 1999. Adaptive strategies of
edaphic arthropods. Annu. Rev. Entomol. 44: 233-256.

Villani, M. G., L. L. Allee, L. Preston-Wilsey, N. Consolie, Y. Xia, and R. L.
Brandenburg. 2002. Use of radiography and tunnel castings for observing mole
cricket (Orthoptera: Gryllotalpidae) behavior in soil. Amer. Entomol. 48: 42-50.

Villani, M. G., S. R. Krueger, P. C. Schroeder, F. Consolie, and N. H. Consolie.
1994. Soil application effects ofMetarhizium anisopliae on Japanese beetle
(Coleoptera: Scarabaeidae) behavior and survival in turfgrass microcosms.
Environ. Entomol. 23: 502-513.

Walker, S. L. 1979. Population estimation, damage evaluation and behavioral studies on
mole crickets Scapteriscus vicinus and S. acletus (Orthoptera: Gryllotalpidae).
Masters Thesis, University of Florida, Gainesville.

Walker, T. J. 1985. Systematics and life cycles. pp 3-10. In T.J. Walker (ed.) Mole
Crickets in Florida. Florida Agric. Exp. Sta. Bull. 846.

Walker, T. J. and N. Dong. 1982. Mole crickets and pasture grasses: Damage by
Scapteriscus vicinus, but not S. acletus (Orthoptera: Gryllotalpidae). Fla. Entomol.
65: 300-306.

Walker, T. J., R. C. Littell, and N. Dong. 1982. Which mole crickets damage
bahiagrass pastures? Florida Entomol. 65: 110-116.









Walker, T. J. and D. A. Nickle. 1981. Introduction and spread of pest mole crickets:
Scapteriscus vinicus and S. acletus reexamined. Ann. Entomol. Soc. Am. 74: 158-
163.

Walker, T. J., J. P. Parkman, J. H. Frank and D. J. Schuster. 1996. Seasonality of
Ormia depleta and limits to its spread. Biol. Control 6: 378-383.

Wang, C., J. E. Powell, and K. B. Nguyen. 2002. Laboratory evaluations of four
entomopathogenic nematodes for control of subterranean termites (Isoptera:
Rhinotermitidae). Environ. Entomol. 31: 381-387.

Williams, J. J. and L. N. Shaw. 1982. A soil corer for sampling mole crickets. Florida
Entomol. 65: 192-194.

Williams, J. K., C. R. Townsend and R. Poulin. 2001. Mermithid nematode infections
and drift in the mayfly Deleatidium spp. (Ephemeroptera). J. Parasitol. 87: 1225-
1227.

Zhou, X., H. K. Kaya, K. Heungens, and H. Goodrich-Blair. 2002. Response of ants
to a deterrent factors) produced by the symbiotic bacteria of entomopathogenic
nematodes. Appl. Environ. Microbiol. 68: 6202-6209.

Zimmerman, R. J. and W. S. Cranshaw. 1990. Compatibility of three entomogenous
nematodes (Rhabditida) in aqueous solutions of pesticides used in turfgrass
maintenance. J. Econ. Entomol. 83: 97-100.