(Coleoptera: Scarabaeidae) in Turfgrass

SEASONAL PHENOLOGY, HOST RANGE, AND MANAGEMENT OF Tomarus subtropicus (COLEOPTERA: SCARABAEIDAE) IN TURFGRASS

OLGA KOSTROMYTSKA

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

UNIVERSITY OF FLORIDA

2007

To My Dear Parents

ACKNOWLEDGMENTS

I would like to sincerely thank Dr. Eileen A. Buss, my supervisory committee chair, for her support and guidance and giving me an opportunity for my professional and personal growth.

She has been a role model of professional honesty, work ethic, enthusiasm and dedication balanced with love and care in the personal life, which will inspire me throughout my further career. Working with Dr. Buss was enjoyable and rewarding. I want to acknowledge my supervisory committee members (Drs. Ronald Cave and Catharine Mannion) for their contribution. I am grateful for the research sites, assistance and cooperation provided by Eugene

McDowell (Tony’s Pest Control), Pete Quartuccio (All-Service Pest Management), Pest

Solutions Plus, Nancy Palmer (Master Gardener), Greig Henry, Trish Woods, and companies that provided me with products: Bayer Environmental Science, Dow AgroScience, Arysta

LifeScience, and Becker Underwood. I would like to thank all of my lab partners who helped me

with my research: Cara Vazquez, Paul Ruppert, James Nichols, Ta-I Huang and Amin Cheikhi. I

would also like to thank Dr. D. Boucias, K. Nguyen, and L. Buss for their help.

TABLE OF CONTENTS

page

ACKNOWLEDGMENTS ...... 4

LIST OF TABLES...... 7

LIST OF FIGURES ...... 8

ABSTRACT...... 9

CHAPTER

1 INTRODUCTION ...... 11

2 BIOLOGY OF Tomarus subtropicus IN ST. AUGUSTINEGRASS IN SOUTHERN FLORIDA...... 18

Materials and Methods ...... 19 Seasonal Phenology...... 19 Adult Behaviors...... 20 Developmental Time ...... 22 Results...... 22 Seasonal Phenology...... 22 Adult Behaviors...... 24 Development Time ...... 25 Discussion...... 25

3 FEEDING HABITS OF Tomarus subtropicus LARVAE...... 39

Materials and Methods ...... 40 Effect of Soil Organic Matter Content on Survival and Growth of First Instars ...... 40 Survival, Growth and Development of T. subtropicus Larvae on Different Warm Season Grass Species...... 41 Results...... 43 Effect of Soil Organic Matter Content on Survival and Growth of First Instars ...... 43 Survival, Growth and Development of T. subtropicus Larvae on Different Warm Season Grass Species...... 43 Discussion...... 45

4 MANAGEMENT OF Tomarus subtropicus ...... 64

Introduction...... 64 Material and Methods...... 66 Evaluation of Insecticides for Preventive Control of T. subtropicus ...... 66 Curative Control of T. subtropicus...... 68 Infectivity of Five Nematode Species against Second and Third Instars...... 69

Reproduction of S. scarabei and S. glaseri in Third Instar T. subtropicus...... 70 Results...... 70 Insecticide Efficacy ...... 70 Nematode Infectivity against Second and Third Instar T. subtropicus...... 71 Reproduction of S. scarabei and S. glaseri in Third Instar T. subtropicus...... 71 Discussion...... 71

5 CONCLUSIONS ...... 81

LIST OF REFERENCES...... 83

BIOGRAPHICAL SKETCH ...... 94

LIST OF TABLES

Table page

2-1. Abundance and morphological parameters of different T. subtropicus life stages collected during blacklight trapping and soil sampling in 2005-2006...... 31

2-2. Air and soil temperatures and relative humidity (2005-2006) from the weather station KFLST. JA1 located in St. James City, FL...... 32

3-1. Clipping height for grasses used in the experiment...... 48

3-3. Third instar T. subtropicus survival and growth on different grass species...... 50

3-4. Statistics showing effect of time and grub feeding and their interaction on clippings collected during eight-week period of the 2005 experiment ...... 51

3-5. Tomarus subtropicus survival, development and growth on different species of warm season grasses when reared from the first instar...... 52

3-6. Statistics showing effect of time and grub feeding and their interaction on clippings collected during eight-week period of the 2006 experiment...... 53

3-7. Statistics showing effect of time and grub feeding and their interaction on turfgrass quality in 2006...... 54

4-1. Effectiveness of four insecticides tested for preventive control against T. subtropicus grubs in the greenhouse ...... 76

4-2. Mortality of first instar Tomarus subtropicus caused by selected insecticides in the field test...... 77

4-3. Effectiveness of four insecticides tested for curative control against T. subtropicus...... 78

LIST OF FIGURES

Figure page

2-1. Abundance of different life stages of T. subtropicus collected from soil samples in southwestern Florida in 2005...... 33

2-2. Abundance of different life stages of T. subtropicus collected from soil samples in southwestern Florida in 2006...... 34

2-3. Soil activity of adult T. subtropicus in relation to rainfall in 2005-2006...... 35

2-4. Flight activity of adult T. subtropicus during 2005-2006 in relation to rainfall...... 36

2-5. Developmental time from pupation to adult emergence at 23ºC, RH 87% and 14:10 hr L:D...... 37

2-6. Development time from oviposition to egg hatch to at 24ºC and 14:10 hr L:D...... 38

3-1. Reduction of root mass caused by third instar T. subtropicus feeding, 2005...... 55

3-2. Dry weight of St. Augustinegrass clippings collected weekly for 8 weeks, 2005...... 56

3-3. Dry weight of bahiagrass clippings collected weekly for 8 weeks, 2005...... 57

3-4. Dry weight of centipedegrass clippings collected weekly for 8 weeks, 2005...... 58

3-5. Dry weight of bermudagrass clippings collected weekly for 8 weeks, 2005...... 59

3-6. Dry weight of zoysiagrass clippings collected weekly for 8 weeks, 2005...... 60

3-7. Dry weight of ryegrass clippings collected weekly for 8 weeks, 2005...... 61

3-8. Reduction of root mass caused by third instar T. subtropicus feeding, 2006...... 62

3-9. Effect of grub feeding on total plant yield by warm season grasses, 2006 ...... 63

4-1. Infectivity of entomopathogenic nematodes against T. subtropicus second and third...... 79

4-2. Productivity of S. scarabei and S. glaseri reared on third instar T .subtropicus...... 80

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

SEASONAL PHENOLOGY, HOST RANGE AND MANAGEMENT OF Tomarus subtropicus (COLEOPTERA: SCARBAEIDAE) IN TURFGRASS

Olga Kostromytska

May 2007

Chair: E. A. Buss Major: Entomology and Nematology

Tomarus subtropicus is the most destructive pest of turfgrass in south Florida. Traditional

white grub management programs have not been successful against this species and severe turf

damage has resulted. Little is known about this scarab’s biology and management in turfgrass.

To study adult flight activity, blacklight traps were operated in Cape Coral and Punta Gorda, FL,

during 2005-2006. At the same time, soil sampling was conducted to determine which life stages

were present in the soil. Observations on adult feeding and mating behavior and oviposition were

conducted in the laboratory.

Warm season grasses (e.g., St. Augustinegrass (Stenotaphrum secundatum [Walt.]

Kuntze), ‘Tifway’ bermudagrass (Cynodon dactylon × transvaalensis Burtt-Davy), ‘Empire’

zoysiagrass (Zoysia japonica Steud.), centipedegrass (Erimochloa ophiuroides [Munro] Hack.),

‘Pensacola’ bahiagrass (Paspalum notatum Flugge), ‘Sea Dwarf’ seashore paspalum (Paspalum vaginatum Swartz.), and ‘Gulf’ annual ryegrass (Lolium multiflorum Lam.) were tested as potential hosts for T. subtropicus grubs in greenhouse no-choice tests in 2005 and 2006. Third instars (in 2005) and first instars (in 2006) were individually reared in pots with a single turfgrass species for 8 weeks. Grub weight gain and grass root reduction, quality and growth were measured. To determine the effect of soil organic matter on the growth and development of T.

subtropicus grubs, first instars were placed individually in seedling trays with sand or peat, with or without grass, and reared for 4 weeks. Grub initial and final weights, and final root weights were measured. In addition, several insecticides were tested as preventive or curative control options in field and greenhouse tests. The effectiveness of six nematode species against T. subtropicus second and third instars and nematode reproduction in third instars were determined in laboratory experiments.

My results suggest that T. subtropicus is univoltine, with peak adult flight activity in July–

August. Eggs are present from late June to early August, first instars occur in July – August, second instars are present in August – September, and most grubs are third instars by October.

All of the warm season turfgrasses tested were suitable hosts for T. subtropicus grubs.

Insecticide tests indicate that halofenozide, clothianidin and imidacloprid can provide preventive control if grubs are contacted by the products, and carbaryl and trichlorfon were satisfactory curative control products. The nematodes Steinernema scarabei, S. glaseri and Heterorhabditis bacteriophora were effective against T. subtropicus second instars.

CHAPTER 1 INTRODUCTION

The Scarabaeidae is a large and diverse beetle family which includes about 2,000 genera and 25,000 species (Leal 1998), many of which are economically important pests with worldwide distribution. The subfamilies that contain the major pests are Melolonthinae,

Rutelinae, Cetoniinae, and Dynastinae. Most plant damage is usually associated with the larvae

(e.g., white grubs), which typically feed on plant roots or tunnel in the soil disrupting the root system. Sometimes secondary damage from the digging of raccoons, skunks, armadillos, or birds looking for grubs to eat may also be a problem (Potter and Braman 1991). Adult feeding habits vary by species, but some adults can defoliate trees and shrubs (Phyllophaga spp., Popillia japonica Newman), feed on ripening fruits (Cotinis nitida (L.), Anomala spp.), attack plant

seedlings and stems (Dyscinetus morator (F.), Euetheola humilis rugiceps (LeConte),

Heteronychus morator (F.)) or do not feed at all (Cyclocephala spp.) (Hayes 1917, Gordon and

Anderson 1981, Watve et al. 1981, Foster et al. 1986, Spencer and Jarrat 1989, Matthiessen

1999, Harari et al. 2001, Logan et al. 2001, Ratcliffe 2003).

Many scarab pest species have generalist feeding habits. White grubs can be pests of

cranberries, soybeans, peanuts, potatoes, sweet potatoes, various fruits, vegetables, ornamental

plants and trees (Davis 1916; Hayes 1917; Litsinger et al. 1983, 2002; Donaldson et al. 1990;

Staines 1990; Oliveira et al. 2000; Harari et al. 2001; Zhang et al. 2003; Pardo-Locarno et al.

2005, Anitha et al. 2006; Jackson and Klein 2006). However, most of the reported white grub

damage has been associated with the Gramineae (Poaceae) and grubs are considered major pests

of lawns, sport and utility turf, golf courses, pastures, and agriculture crops, such as corn, rice,

wheat, and sugarcane (Litsinger et al. 1983, 2002; Cherry and Allsopp 1991; Lura and Nyren

1992; Flanders et al. 2000; Logan et al. 2001; Ansari et al. 2003; Drinkwater 2003; Pardo-

Locarno et al. 2005). In some cases, more damage occurs in newly developed agricultural fields

or those rotated with pastures (Watve et al. 1981, Matthiesen 1999).

White grubs have been known as major pests of turfgrass throughout the United States.

The life cycle and management of the white grub complex that infests cool season grasses have

been thoroughly studied, but similar studies in warm season turfgrasses have been limited. The

dominant species include Japanese beetle (P. japonica), northern masked chafer (Cyclocephala borealis Arrow), southern masked chafer (C. lurida Bland), oriental beetle (Exomala orientalis

(Waterhouse)), Asiatic garden beetle (Maladera castanea Arrow), European masked chafer

(Rizotrogus majalis Razoumowski), green June beetle (Cotinis nitida L.), black turfgrass

ataenius (Ataenius spretulus (Haldeman), and May – June beetles (Phyllophaga spp.)

(Niemczyk and Dunbar 1976, Tashiro 1987, Potter and Branam 1991, Cowles et al. 1999,

Bauernfeind 2001, Brandhorst-Hubbard et al. 2001, Grewal et al. 2001, Koppenhöfer et al.

2004). The most abundant and destructive turfgrass pests in Florida include C. lurida, C. nitida,

Euphoria sepulcralis (F.), Strategus antaeus (Drury), and Tomarus subtropicus (Blatchley)

(Reinert 1979, Buss 2006). Most species are univoltine and have three larval instars, but May or

June beetles need three years in the northern United States and about one to two years in the south to complete development. Several species (Anomala marginata (F.), A. innuba (F.), C. lurida, and Cyclocephala parallela (Casey)) which are univoltine in northern states have bimodal adult flight peaks in Florida (Forschler and Gardner 1991, Flanders et al. 2000, Buss

2006). The black turfgrass ataenius usually has two generations per year (Tashiro 1987, Vittum

1986, Potter and Branam 1991, Potter and Held 2002). Along coastlines and in subtropical and tropical climates, constant food availability may favor rapid development and insect abundance.

The sugarcane grub, T. subtropicus, was scarce and not considered a pest in the 1930s in the United States (Blatchley 1930). It was distributed along the coastal regions of Alabama, central and southern Florida, Georgia, South Carolina and North Carolina by 1959 (Cartwright

1959). In 1972, a small sugarcane grub infestation and its damage were reported in a sugarcane field in Belle Glade (Palm Beach Co.) (Summers 1974). The population continued to grow rapidly, and by 1978, it had spread throughout the entire sugarcane growing area in Florida

(Prewitt and Summers 1981). This species could reduce sugarcane yield up to 40% and it was not profitable to harvest the crop if a large infestation was present (Sosa 1984). The species has since then been considered the “primary sugarcane pest of great economic significance” (Cherry

1985) and the biology and management of T. subtropicus have been examined primarily on

sugarcane (Miller and Bell 1981, Prewitt and Summers 1981, Watve et al. 1981, Cherry 1984a,

b, Sosa 1984, Cherry 1985, Boucias et al. 1986, Hall 1987, Cherry et al. 1990, Cherry 1991,

Cherry and Coale 1994, Cherry and Klein 1997). Although some variation exists in dates

between these studies, its univoltine life cycle could be summarized as the following. Adults are

usually active from April until August, with peak activity in June. Eggs and first instars are

prevalent in June and July. Second and third instars occur in September, and third instars are

present until April. Pupation occurs in April and May (Gordon and Anderson 1981, Miller and

Bell 1981, Sosa 1984, Cherry 1985, Hall 1987). Ovipositing females and larvae seem to prefer

muck soils (Gordon and Anderson 1981, Hall 1987). Adults feed on the stems of young

sugarcane plants in the field (Watve et al. 1981) or on sliced carrots in the laboratory (Cherry et al. 1990). Yet, little is known about the life history of T. subtropicus in turfgrass. In 1979, it was reported as a part of a species complex damaging bermudagrass (Cynodon dactylon (L.)) and St.

Augustinegrass (Stenotaphrum secundatum (Walt. Kuntze)) in residential lawns, industrial

plantings, and other turf areas in Miami, FL (Reinert 1979).

How to specifically manage white grubs in warm season grasses is poorly known, so a typical management program based on studies of scarab pests in cool season turfgrass was

assumed to be applicable. Cultural practices are important aspects of turf management; they

usually are low cost, environmentally friendly and take into account the biology of turfgrass and

the insects feeding on it. Cultural controls can be used to prevent or reduce a grub infestation by

altering the behavior of egg-laying adults, affecting egg and larval survival, and increasing grass

resistance. For example, withholding irrigation during adult flight may make non-irrigated sites

less attractive to adult females, decrease the survival of eggs and newly hatched first instars and

reduce the rate of grub development (Potter et al. 1996). However, fall irrigation helps the grass

to withstand the root feeding and does not have a significant effect on white grub reproduction

and development (Crutchfield et al. 1995). Fall fertilization may help the turfgrass to recover

after grubs stop feeding. Spring fertilization may increase the visible effect of grub damage

because it stimulates shoot growth and impedes root system formation (Crutchfield et al. 1995).

Mowing at an elevated cutting height (18 vs. 7.6 cm), and using aluminum sulfate to reduce soil

pH from 6.6 to 5.4 makes the root systems of cool season grasses more fibrous and less suitable

for grub feeding (Potter et al. 2000). Endophytic fungi, Neotyphodium spp. and Epichloë spp.

(Clavicipitaceae: Ascomycetes), associate mutualistically with cool season grasses and can

slightly increase grass resistance to white grubs (Koppenhöfer et al. 2003). Grub survival on

grass with endophytes is slightly lower than survival on grass without these fungi. Some other

practices have little or no effect on grub populations, such as spring application of lime, soil

compacting, or aerification before adult flight (Potter et al. 2000). The importance of cultural

practices in suppressing grubs in warm season grasses is poorly known. Flooding is a successful

method for T. subtropicus grub and pupal control in sugarcane (Cherry 1984a), but is not feasible

in turfgrass.

The natural enemy complex of damaging grubs in Florida has not been extensively

examined. Parasitoids of white grubs include members of Tiphiidae and Scoliidae. Several

species of both families live in Florida (Grissell 1977 a, b), but their hosts and effectiveness are

not known. However, up to 50% of T. subtropicus were reported to be parasitized by an

unindentified tiphiid in a highly infested sugarcane field (Prewitt and Summers 1981).

Entomopathogenic nematodes (e.g., Heterorhabditis bacteriophora (Poinar), H. zealandica

(Poinar), and Steinernema glaseri (Steiner)) can be effective biological control agents for some

white grub species (Koppenhöfer et al. 2002; Koppenhöfer and Fuzy 2003ab, Koppenhöfer et al.

2006). If the timing and application technique are correct, nematodes could be as effective as

chemical control. In countries where pesticides are strictly limited (e.g., Germany, Japan),

nematodes are widely marketed for white grub control in turf (Georgis et al. 2006). However, the

high cost of production and application, and inconsistent field effectiveness limit the use of

nematodes. In addition, susceptibility to nematodes varies greatly among scarab species and

stage of grub development (Koppenhöfer et al. 2004). A recently described species, Steinernema

scarabei Koppenhöfer, is highly pathogenic to several scarab species, and is claimed to be more

effective for white grub control than other species (Koppenhöfer and Fuzy 2003a,b). Laboratory experiments (Sosa and Beavers 1985) and field tests (Sosa and Hall 1989) on the effectiveness of

S. glaseri and S. feltia against T. subtropicus were conducted with sugarcane as a host plant.

Steinernema glaseri provided excellent control in the laboratory test (100% mortality) and was more effective than S. feltia (17% of mortality), but neither species significantly reduced the grub

population under field conditions (Sosa and Beavers 1985, Sosa and Hall 1989). Milky spore disease (Paenibacillus popilliae Dutky) is a naturally occurring and commercially available pathogen. However, only 1.9% of T. subtropicus larvae and 6.1% of Cyclocephala spp. larvae were infected with it in Florida sugarcane fields (Cherry and Boucias 1989). Entomopathogenic nematodes and imidacloprid and nematodes with Bacillus thuringiensis can work in synergy against white grubs, which makes nematodes compatible with IPM, although the synergistic effect varies depending on nematode species, grub species, and insecticide types and rates

(Koppenhöfer and Kaya 1997, Koppenhöfer et al. 2000, 2002).

Traditional control of white grubs is based on the use of preventive and curative insecticides. The goal of preventive control is to suppress grub populations before noticeable damage occurs, so insecticides are applied immediately after egg hatch. Clothianidin (Arena®,

Arysta LifeScience, San Francisco, CA), imidacloprid (Merit®, Bayer Environmental Science,

Raleigh, NC), and halofenozide (Mach 2®, Dow AgroScience, Indianapolis, IN) are used for preventive control. These insecticides have long residuals, target small grubs, and are plant systemic (Potter 1999). Curative control becomes necessary when turf damage is noticeable, which is usually concurrent with the presence of third instars and their intensive feeding on roots.

Current curative insecticides include a broad spectrum carbamate (carbaryl (Sevin®, Bayer

Environmental Science, Research Triangle Park, NC)) and organophosphate (trichlorfon

(Dylox®, Bayer Environmental Science, Research Triangle Park, NC )), which have short residual activity and can provide more than 70% control (Grewal et al. 2001, Koppenhöfer et al.

2005). Most references about the chemical control of T. subtropicus are out-of-date. Most of the organophosphates and carbamates (isazofos, fonofos, carbofuran) that were tested against this species were discontinued because of the Food Quality Protection Act of 1996 (Reinert 1979,

Watve et al. 1981). Also, insecticide trials were conducted against a species complex (T. subtropicus, C. parallela, S. antaeus, Phyllophaga spp.), rather than one species. However, it was mentioned that T. subtropicus was harder to control with insecticides than the other white

grub species (Watve et al. 1981, Reinert 1979).

Given the limited information about general white grub biology and management in warm

season turfgrasses and the growing importance of grubs in Florida, I sought to describe the

biology of T. subtropicus in St. Augustinegrass and develop an IPM program to suppress its

populations. This thesis is divided into chapters on the biology, host range, and management of

T. subtropicus in Florida.

CHAPTER 2 BIOLOGY OF Tomarus subtropicus IN ST. AUGUSTINEGRASS IN SOUTHERN FLORIDA

The sugarcane grub, Tomarus subtropicus Blatchley, is a severe pest of turfgrasses and sugarcane (Reinert 1979, Watve et al. 1981) in central and southern Florida. The larvae feed on roots near the soil surface, which reduces plant vigor, stunts growth, and kills plants. The phenology of white grub activity in sugarcane is known (Watve et al. 1981, Hall 1987, Cherry

1985), but control is limited to flooding (Cherry 1984a). To manage infestations in residential lawns, lawn care personnel apply preventive insecticides based on application timing for grubs in the northern United States and in sugarcane, but with limited success. Curative insecticides are applied after damage occurs and in severe cases damaged sod is replaced. Because of the field failures of preventive insecticides, it was suspected that the biology of T. subtropicus in St.

Augustinegrass lawns may differ from that reported in sugarcane fields, thus resulting in inappropriate application times.

Sugarcane fields and St. Augustinegrass lawns have very different characteristics, which could influence T. subtropicus larval growth. The soil in a sugarcane field is a typically rich, organic muck, poorly drained, and therefore saturated during most of the growing season

(McCollum et al. 1978, Belz et al. 1990). The soils in Florida lawns are mostly sandy and siliceous, with up to 30% of shell material and rock fragments (Henderson 1984a, b). Sugarcane is planted from August to January, harvested every 12-18 months (October - March) and replanted every 2-3 years. In contrast, turfgrass is a perennial planting and a continuous food source for insects. In spite of the similar climate across southern Florida, the microclimate of lawns with grass cut at ~7.5 cm and sugarcane fields with 1.2- 3.7 m-tall plants may be very different.

To begin developing an integrated pest management (IPM) program on turfgrass, the biology of T. subtropicus needed to be examined. Thus my objectives were to monitor its soil and flight activity in St. Augustinegrass lawns, identify any natural enemies, and describe the adult feeding, mating and ovipositing behaviors.

Materials and Methods

Seasonal Phenology

Soil sampling. To determine the seasonal phenology of T. subtropicus, four different St.

Augustinegrass residential lawns with existing infestations of T. subtropicus in Cape Coral (Lee

Co.) and Punta Gorda (Charlotte Co.) that were under professional care were sampled at each sampling date. Soil temperature (10 cm depth) and relative humidity were taken at the sampling sites on each date. Eight soil samples (30 cm², 12 cm deep in 2005; 30 cm², 20 cm deep in 2006) from each lawn were examined weekly from 31 May to 31 August 2005 and 1 June to 29

September 2006 and biweekly from 1 September to 30 November 2005 and 30 March to 30 May

2006. Samples were collected monthly from December 2005 to March 2006 because only third instars were present. The soil was visually inspected on site; all life stages of T. subtropicus and any natural enemies were collected, placed individually into Solo cups, and transported to the laboratory in a cooler. Adults and third instars were identified, and eggs, first and second instars were reared at least to the third instar to be identified. Grub head capsule diameter, the width and length of pupae and adults, and the weight of all individuals were measured.

To rear grubs, plastic pots (15 cm diameter) were filled with potting mix FaFard #2

(sphagnum peat moss, horticultural perlite, horticultural vermiculite, wetting agent, starter nutrients) and planted with St. Augustinegrass variety ‘Palmetto’. Because of their cannibalistic behavior, only one grub was placed in each pot. All pots were held in the greenhouse and air and soil temperatures were recorded. From December to March pots with grubs were placed on

heating mats to maintain the soil temperature above 10º C. Grubs were visually inspected under

a binocular dissecting scope for the presence of parasitoids, pathogens or nematodes. Nematodes were identified by Dr. Khoung Nguyen (Entomology and Nematology Department, UF), and the other pathogens were identified by Dr. Drion Boucias (Entomology and Nematology

Department, UF).

Blacklight trapping. One Circline ultraviolet blacklight (BioQuip®, Rancho Dominguez,

CA) was placed at each of two locations in Punta Gorda and one location in Cape Coral in 2005

(Figure 2-1). Four traps (two in Punta Gorda and two in Cape Coral) were operated in 2006.

Blacklight traps were hung 90 cm from the ground and were operated continuously from 1 April

to 30 November 2005 and 30 March to 29 September 2006. Beetles were collected once a week.

All trapped T. subtropicus adults were delivered to the laboratory, identified, sexed, counted and

used in further experiments. Sex of the beetles was determined externally by looking at the sixth

abdominal sternite, which is concave in males (Cherry 1985).

Adult Behaviors

Adult maintenance. Clear 2 L plastic bottles were cut at ~5 and 9 cm from the top to

remove the funnel and create a cylindrical ovipositional container (9 cm diameter, 20 cm deep)

and a lid with ~5 cm diameter opening at the top. Fine mesh was glued to the lid opening to allow airflow and prevent beetle escape. The bottles were filled with Black Velvet peat, and planted with St. Augustinegrass variety ‘Palmetto’ plugs. One male-female pair was placed on the grass and provided with ~50 g of mango as a food source. The bottles were held in either a

Florida reach-in rearing chamber or in the greenhouse at 24 ± 3°C and a photoperiod of 16:8 and

14:10 (L:D), respectively. Beetles remained in the containers until death. The soil was examined

periodically for eggs, and mango and grass were replaced as needed.

Observations on feeding and mating behavior. Three methods were used to observe T. subtropicus mating behavior. In 2005, 12 field collected beetles were put in a deep container (50 cm wide, 100 cm long, and 50 cm deep) filled with autoclaved top soil (30 cm deep) and planted with St. Augustinegrass variety ‘Palmetto’. Observations were conducted for five consecutive nights, but no surface activities during the day or night were noticed. Therefore, beetles were placed in trays (50 cm wide, 100 cm long, and 12 cm deep) with 1 cm layer of mulch and provided fruits and 10% honey water for food. In 2006, plastic shoe boxes (19.5 cm wide, 27 cm long, and 9 cm deep) were filled with rectangular pieces of sod (St. Augustinegrass variety

‘Palmetto’). Four beetle pairs (male–female) were put in each container and their behavior was observed. All observations were conducted using a red light. Beetle activities, ambient temperature and humidity were recorded.

Impact of resource availability on oviposition. To determine if food availability affected oviposition, I conducted the following no-choice test. In 2006, ovipositional containers were assigned to four treatments (10 replicates for each treatment): 1) 50 g of mango and St.

Augustinegrass variety ‘Palmetto’, 2) only St. Augustinegrass, 3) only mango, and 4) neither mango nor grass. One male-female pair (1-3 days old) was randomly placed into each ovipositional container. Containers were placed in the greenhouse in a completely randomized design. Ambient temperature averaged 28 ± 4º C and relative humidity 75 ± 16%. Soil in the bottles was inspected for eggs every other day, and bottles were filled with fresh soil and new grass and mango. The number of eggs laid and time of beetle mortality were recorded. Dead females were dissected, the number of unlaid eggs was determined, and the length of the female elytra was measured. Data were analyzed using a two-way ANOVA for effects of food and grass

availability on female longevity and the number of eggs laid, and Tukey’s test was used for mean

separation (SAS Institute 2003).

Developmental Time

Pupation. To determine the length of the pupal stage, field collected grubs were reared in pots of St. Augustinegrass in the greenhouse. Prepupae were placed into artificial pupation

chambers containing autoclaved Black Velvet® peat soil compacted in 59 ml Solo cups, with

five 3-mm diameter holes cut into lids. Prepupae and pupae were checked daily and the duration

of pupation and ambient temperatures were recorded.

From oviposition to egg hatch. Twelve pairs of beetles were placed into individual

oviposition containers and provided with grass and food to enhance oviposition. Containers were

checked daily for egg presence. Fifty-six apparently healthy eggs laid on the same day were

chosen for determining egg developmental time. Eggs were placed onto moistened filter paper,

covered by another moistened filter paper in a plastic quatered Petri dishes (9 cm diameter), four

eggas per each dish and held at 24°C in a reach-in rearing chamber, and checked daily for

hatching.

First instar. The duration of the first instar was determined using 36 larvae that hatched on

the same day. Two grubs were put in the soil of 15 cm diameter plastic pots filled with potting mix and St. Augustinegrass variety ‘Palmetto’. Each grub was put into a hole (7 mm diameter,

10 cm deep) and covered with soil. Pots were inspected every other day. Grub developmental stage, mortality and ambient temperature were recorded.

Results

Seasonal Phenology

From the soil samples, I collected 70 adults, 147 eggs and 535 larvae in 2005, and 68

adults, 245 eggs, 578 larvae and 14 pupae in 2006. Most of the eggs (91%) were present from 29

June to 9 August in 2005 and from 22 June to 10 August in 2006 (Figure 2-1, 2-2). First instars

were prevalent from 12 July to 23 August during both years, second instars occurred from July

27 to 29 September, and third instars were in the soil from early August 2005 and 2006 until the

following April. Each instar could be easily distinguished by the head capsule measurement

(Table 2-1). The head capsule of T. subtropicus larvae also differed from Cyclocephala spp. and

Phyllophaga spp. that co-occurred in the soil by being pitted and a dark rusty red color, rather than smooth and tan colored.

The grub population within each sampling site was fairly clumped. I found more grubs on slopes along ditches, over-watered areas, and edges of yards and flower beds. Most specimens were collected in the thatch (~2-5 cm deep) from May to November. About half of the larvae were 10-15 cm deep from December to April.

No parasitoids were recovered from the soil samples. Four percent (19 grubs) of the grubs collected in 2005 were infected with Paenibacillus popilliae Dutky, and the majority of the infected grubs were third instars (only one second instar was infected). In 2006, 3% (17 grubs) of all the collected grubs were infected with Steinernema glaseri (Poinar). Air and soil temperatures varied seasonally during soil sampling (Table 2-2).

Most of the adults (93%) were collected in the soil from 1 June to 13 July 2005 and from

30 May to 12 July 2006 (Figure 2-3). Adult flight activity peaked from 20 July to 21 August in both years, and by then adults were absent in the soil (Figure 2-3). More males were collected in the soil than in blacklight traps. The sex ratios were 1.03:1 (70 males, 68 females) and 1:2.2 (71 males, 159 females) in soil and blacklight traps, respectively. Males appeared to be active in the soil and blacklight traps earlier than females, but females were more abundant at the end of the soil activity period. Only single adults were in the soil in May, pairs of males and females

occurred from June to early July, and females were ovipositing in the soil by mid-July. The

increasing number of T. subtropicus adults in the soil coincided with the first big peak of rainfall in early summer (Figure 2-3), and flight activity peaked for a short time after the peak of rainfall

(Figure 2- 4).

Adult Behaviors

Most adult activity was noticed between 1-3 A.M. Beetles did not appear on the surface if they were in a container with soil deeper than 10 cm. Mating occurred in the soil or mulch and lasted on average 90 min. The female started moving first after mating, dislodged the male, remained motionless for about 1 hour, and then started crawling actively on the surface or under the mulch. All flight attempts observed were made by males. If allowed to fly, beetles flew about

2-3 m, about 30 cm high, and crawled actively after the flight. Both females and males stridulated by moving elytra up and down during their crawling activity and when disturbed.

Beetles made tunnels and chambers in the sod roots provided. Beetles were never observed feeding on the surface. All feeding appeared to occur from underneath the fruits or tunneling directly in the fruit tissue.

The presence of St. Augustinegrass grass and 20 cm soil depth were necessary for oviposition. Of 82 females studied, 69 (84%) laid eggs in 2005. Ovipositing females laid on average 25 eggs; the maximum number of eggs laid by a single female was 48, and the minimum was one egg. The eggs were laid individually in earthen chambers 4×4 mm (approximately the size of a mature egg), 1-2 cm apart and at varying depths, although in the field most eggs were within the top 5-7 cm of soil, in the root zone and dispersed horizontally.

Both grass and mango availability had a significant effect on the number of eggs laid when compared to no food available (F = 11.58; df = 1, 39; P = 0.0016 and F = 4.20; df = 1, 33; P =

0.047, respectively). More eggs were laid if grass and mango were provided (32.4 ± 0.6) than

without food and grass (16.7 ± 2.4). The interaction between the two factors was not significant

(F = 0.14; df =1, 39; P = 0.7081) and was excluded from the analysis. Longevity of the females

in the containers with the grass (100.1 days) was greater than in the containers without grass

(76.5 days) (F = 8.42, df= 1, 39; P = 0.0062). The presence of mango did not have any

significant effect on female longevity.

Development Time

In the pots held in the greenhouse, most grubs ceased feeding in February and created an

oval or elongate cell by moving and pushing substrate around themselves. Cells were about 10-

20 cm deep and usually were built against the pot wall. Larvae in some cases moved out of the

cells and built a new cell. A portion of grubs in the greenhouse and in the field continued feeding

until March. However, all grubs pupated by the end of May. The development time from

pupation to adult emergence was 21.3 ± 0.2 days (range: 17 to 27 days) (Fig 2-5). It took 20.9 ±

0.3 days for males to emerge and 21.7 ± 0.3 days for females, but the duration of pupation for

males and females did not significantly differ (t = -1.8, df= 93, P = 0.0749). It took 16.2 days for eggs to hatch (range: 13-20 days) (Figure 2-5). Development from first instar to second instar averaged 22.3 days (range: 17 - 30 days) at a soil temperature averaging 23.1º C and air temperature 29.1ºC.

Discussion

Tomarus subtropicus is univoltine, with a similar developmental pattern in both sugarcane and St. Augustinegrass. The main difference is the timing of adult activity and oviposition, which is important for management timing. Adult flight peaked about a month later in turfgrasses than in sugarcane (Watve et al.1981, Hall 1987). Eggs and first instars similarly were prevalent in July – August in turfgrass, and in May – June in sugarcane (Cherry 1985). The variability in its developmental time between hosts may be related to differences in

environmental conditions within Florida, host plant nutritional quality, or the amount of food

available.

Scarab beetle activity is closely related to environmental conditions such as temperature,

(Barratt and Campbell 1982), rainfall (Gaylor and Frankie 1979, Potter 1981), and soil type

(Cherry and Coale 1989). According to the Southeast Regional Climate Center

(http://www.dnr.sc.gov/climate/sercc), the seasonal temperature fluctuation is similar near Belle

Glade where sugarcane is commonly grown and Cape Coral/Punta Gorda, where my collections were done. However, significant precipitation begins in June in Belle Glade, but starts in July in

Cape Coral and Punta Gorda, which corresponds with the start of T. subtropicus adult activity in

both locations. Rainfall could be an important cue for synchronizing adult emergence to

maximize mate location (Litsinger et al. 2002). The presence of organic matter in soil can

improve an insect’s diet and therefore enhance the growth and development of white grubs (King

1977). It is possible that the T. subtropicus that occur in sugarcane fields with high organic

matter content (68% as reported by Hall (1987)) could develop from third instars to pupae faster

than the grubs in lawns where soils are predominantly sandy with ≤ 5% organic content. Faster

development could contribute to earlier beetle flight in sugarcane compared to turfgrass.

Host plant quality and availability can also affect rates of larval development (Barratt

1982, Dosdall and Ulmer 2004, Moreau et al. 2006). For example, Leucopholis irrotata

(Chevrolat) needs >1 year to develop in unfertilized grasslands, but it is multivoltine when living

in fertilized sugarcane fields (Litsinger et al. 2002). Both sugarcane and St. Augustinegrass are

grown using fertilizer, but frequency and amount vary. The UF/IFAS recommendations for

sugarcane include no fertilization for sugarcane grown on muck soils, 33 kg/ha of N on sandy

muck, 123.3 kg/ha of N on mucky sand and 444.8 kg/ha of N on sandy soil (Rice et al. 2006). St.

Augustinegrass is fertilized about three times from spring to fall with up to 98 – 290 kg/ha of N

(Trenholm 2006). Grass is always available unless it goes dormant or dies, whereas sugarcane is

harvested and replanted every 3-4 years and can be rotated with other crops. On other hand, grass

is mowed regularly, which reduces rooting (Duble 1989, Turgeon 2002, Tucker et al. 2006).

Sugarcane produces about 20 times more dry root mass than St. Augustinegrass and sugarcane

root length is about 13 times longer than St. Augustinegrass (Moris 2005). So during the growing

season the amount of food for grubs is probably greater in sugarcane fields than in the lawns.

However sugarcane roots contain phenols, like ferulic acid and p-coumaric acid (Nut et al.

2004), which fortify plant tissue cell walls and make plants more resistant to insect feeding

(Bergvinson et al. 1997). The amount of these chemicals increases in response to grub feeding

(Nut et al. 2004), which suggests the possibility of induced chemical defenses in sugarcane. The

defensive chemistry of St. Augustinegrass is unknown.

The prolonged pattern of adult T. subtropicus activity (about two months in soil followed

by a two month flight period) observed in St. Augustinegrass seems unusual for phytophagous

North American scarabs. Because blacklight and soil sampling studies were done separately in

sugarcane, it is unclear whether a similar pattern is also present in sugarcane, but a prolonged

adult presence was previously reported (Watve and al.1981, Cherry 1985, Hall 1987). The early

soil activity period is likely a maturation period. Alternatively, it could be a specific pattern of

flight behavior.

Litsinger (2002) described three major types of flight behavior of scarabs, regarding the

purpose of flight and use of feeding and mating sites by adults. The first type of adults (e.g.,

Anomala hueralis (Burmeister), Antitrogus parvulus (Britton), Holotrichia flachi (Brenske),

Lepidiota picticollis (Lea), and Lepidiota crinita (Brenske)) emerge reproductively mature and

they do not need time or food to complete development and oviposit. Males in this case usually

find the females and the females then fly to a nearby location to oviposit. The second type of

flight behavior (exhibited by Lepidiota negatoria (Blackburn) and Phyllopertha horticola (L.)) involves flight to a feeding site, but the purpose of the flight is not to feed, but rather to complete maturation and mate. Adults can feed, but it is not critical for oviposition and fecundity (Milne

1956, 1959, Milne and Laughlin 1956). Oviposition usually occurs shortly after mating near a host tree. The third type of flight behavior (e.g., Dermolepida albohitrum (Waterhouse)) involves flight of both sexes to a food source to gain resources to complete reproductive development and mate. Females fly to oviposit after mating and egg maturation, and they can repeat the cycle two or more times.

Tomarus subtropicus adults appear to exhibit behavior intermediate between the first and second types of flight behavior. Males outnumbered females in the soil samples and blacklights earlier in the season, probably completing pupal development faster than females. This phenomenon is known as protandric emergence and is common among Lepidoptera and

Coleoptera, including scarab species (Kelsey 1951, Barrat and Campbell 1982, Litsinger et al.

2002, Moreau et al. 2006). Lepidopterans with protandric emergence have an advantage of decreasing the duration of the prereproductive period in females (Fagerström and Wiklund

1982). In addition, adult males died sooner in the laboratory and were observed dead on lawns much earlier than females, which is also common among scarabs (Litsinger et al. 2002). The beetles demonstrated subterranean habits in the laboratory; mating was in the soil or mulch and feeding occurred beneath the fruits. In the field, no surface activity was observed from mid-June to mid-July, when beetles were found active in the soil. Adults of some other Dynastinae species are reported to have similar subterranean habits. For instance, Heteronychus arator F., the

African black beetle (Matthiessen 1999, Matthiessen and Learmonth 1998), and Tomarus

gibbosus, which is widely distributed in North America, mate in the soil similarly to T.

subtropicus (Hayes 1917).

Adult T. subtropicus were able to feed on fruits (e.g., bananas, apples, pears, peaches,

mangoes), but preferred over-ripe fruits, especially mangoes. The availability of grass and food

enhanced oviposition in the greenhouse, but females could still oviposit without having fed.

Similarly, longevity was not significantly affected by food presence. However, it is possible that

beetles fed on grass stems or crowns. In sugarcane fields, adults feed on young sugarcane plants

(Watve et al. 1981). Adults of several dynastine species (T. cuniculus (F.), T. gibbosus, Eutheola

humilis rugiceps (LeConte), Dyscinetus morator (F.), and H. arator) can attack stems, roots and

seedlings of the plants, and are reported as serious pests of corn, rice, sugarcane, sweet potato,

carrots, and radishes (Davis 1916, Hayes 1917, Gordon and Anderson 1981, Foster et al. 1986,

Matthiessen and Learmonth 1998, Matthiessen 1999). Thus, if T. subtropicus adults feed on

grass, they do not need to fly to feed, so flight might have only a localized dispersal function,

similar to the behavior of H. arator infesting pastures (Matthiessen 1999).

In summary, T. subtropicus has one generation per year in both sugarcane and St.

Augustinegrass lawns. However, adult flight, oviposition, and egg and first instar occurrence are

one month later in lawns. The life cycle of the species in St. Augustinegrass lawns can be

described as the following: Adults start emerging in May and remain in the soil until the end of

June. Flight activity peaks from mid July to mid August and most of the eggs and first instars,

which are targets for preventive control, are concurrently present in the soil. From late August to

September, second and third instars are present and damage becomes visible in heavily infested

or stressed lawns. By October most grubs are third instars and damage is most severe because

grub feeding coincides with the start of the dry season. The third instar lasts until April, when pupation occurs. Prolonged adult presence and late flight can partially explain the difficulty of preventively controlling T. subtropicus infestations.

Table 2-1. Abundance and morphological parameters of different T. subtropicus life stages collected during blacklight trapping and soil sampling in 2005-2006. Year Life stage No. Mean head Mean Mean width Mean length capsule width weight (mm ± (mm ± SEM) (mm ± SEM) (g ± SEM) SEM) 2005 First instars 112 2.54 ± 0.02 0.11 ± 0.01 - - Second instars 76 4.59 ± 0.04 0.79 ± 0.04 - - Third instars 347 8.06 ± 0.01 3.53 ± 0.05 - - Pupae 0 - - - - Female adults 141 - 0.96 ± 0.03 12.31 ± 0.02 22.48 ± 0.05 Male adults 95 - 0.92 ± 0.04 11.92 ± 0.01 21.88 ± 0.03 2006 First instars 158 2.54 ± 0.01 0.11 ± 0.01 - - Second instars 186 4.48 ± 0.01 0.79 ± 0.02 - - Third instars 234 7.99 ± 0.01 3.29 ± 0.06 - - Pupae 14 - 1.98 ± 0.10 12.82 ± 0.53 26.94 ± 0.68 Female adults 85 - 0.95 ± 0.02 12.25 ± 0.02 22.52 ± 0.03 Male adults 50 - 0.93 ± 0.03 11.88 ± 0.03 22.01 ± 0.02

Table 2-2. Air and soil temperatures and relative humidity (2005-2006) from the weather station KFLST. JA1 located in St. James City, FL. Year Month Max Min Mean Mean Precipitation, RH (%) Air T Air T Air T Soil mm (ºC) (ºC) (ºC) T (ºC) 2005 May 33.4 16.8 25.5 20 73.4 72.9 June 33.4 20.9 26.9 22.3 350.5 81.2 July 35.1 23.6 29.1 21.6 398.3 76.3 August 35.6 23.7 29.4 26.4 113.8 75.2 September 34.7 23.7 27.9 26.4 147.1 75.9 October 32.5 12.6 25.3 22.2 196.1 76.8 November 29.8 11.7 22.6 22.2 64.5 77.4 December 26.6 8.6 17.8 20.6 6.3 79.5 2006 January 27.7 6.9 18.6 17.8 9.7 76.4 February 28.0 5.1 17.6 15.6 33.5 75.4 March 28.9 9.6 20.9 18.3 6.9 70.7 April 31.6 14.9 23.9 20 1.3 71.5 May 33.6 16.6 25.5 21 38.9 70.8 June 35.1 21.7 27.7 24.8 178.8 75.2 July 34.8 23.1 27.8 24.6 269.7 78.4 August 35.7 23.9 28.7 25.9 124.5 75.3 September 34.4 21.2 27.9 24.4 130.0 76.0

60 Eggs

1st instars 50 2nd instars collected 40 3rd instars

Beetles

T. subtropicus subtropicus T. 30 33

20 Total number of Total number

0 l l l t n n u u u g g g g ep ep ep ct ct v v v v c Ju u -J -J u S S S O O o - 5-Jul Au Au -Sep -Oc No No De 7-Jun 4 1-J 8-Jun 12 19 26-J 2-Au 9- 6- 6 3- 0- 4 1- 8- 1- 8-N 5-Nov 2-No 9- 6- 31-May 1 2 2 1 23-A 30-Aug 1 2 27- 1 1 25-Oct 1 2 2

Figure 2-1. Abundance of different life stages of T. subtropicus collected from soil samples in southwestern Florida in 2005.

70 Beetles

Eggs 60 1st instars collected 50 2nd instars

40 3rd instars

T. subtropicus T. Pupae 34 30

20 Total number of number Total 10

0 r y n an eb ar u Jul -J -Jan F Ma M Apr Ma -Jun J -Jul Aug Aug 6 0 3- 7- 4- -May 5 9- 3 17- 31-Jul 4- 8- 1 3 13-Feb 27- 1 2 10-Apr 2 8 22- 1 1 2 11-Sep 25-Sep

Figure 2-2. Abundance of different life stages of T. subtropicus collected from soil samples in southwestern Florida in 2006.

18 25

16 Rainfall Soil sampling 20 14

15 10

35 10 6 Weekly rainfall, cm rainfall, Weekly Total no. of no.Total beetlescollected 4 5

0 0 5-Jul 6-Jul 2-Jan 8-Jun 1-Jun 8-Jun 2-Feb 5-Sep 2-Apr 6-Apr 2-Mar 12-Jul 21-Jul 13-Jul 20-Jul 27-Jul 4-Aug 3-Nov 3-Aug 18-Jan 4-May 16-Jun 22-Jun 15-Jun 22-Jun 29-Jun 18-Oct 18-Feb 29-Sep 13-Sep 27-Sep 24-Apr 13-Apr 20-Apr 27-Apr 18-Dec 18-Mar 11-Aug 18-Aug 29-Aug 18-Nov 10-Aug 17-Aug 24-Aug 31-Aug 15-May 30-May 11-May 18-May 25-May 2005 2006

Figure 2-3. Soil activity of adult T. subtropicus in relation to rainfall in 2005-2006.

40 25

35 Rainfall Black light trapping

20 30

25 15

10 36 15 Weekly rainfall, cm Weekly Total no.of beetles collected 10 5

0 0 6-Jul 5-Jul 2-Jan 1-Jun 8-Jun 8-Jun 2-Feb 5-Sep 6-Apr 2-Apr 13-Jul 20-Jul 27-Jul 2-Mar 12-Jul 21-Jul 3-Aug 3-Nov 4-Aug 4-May 18-Jan 15-Jun 22-Jun 29-Jun 16-Jun 22-Jun 18-Oct 13-Sep 27-Sep 18-Feb 29-Sep 13-Apr 20-Apr 27-Apr 24-Apr 18-Dec 18-Mar 10-Aug 17-Aug 24-Aug 31-Aug 18-Nov 11-Aug 18-Aug 29-Aug 11-May 18-May 25-May 15-May 30-May 2005 2006

Figure 2-4. Flight activity of adult T. subtropicus during 2005-2006 in relation to rainfall

. 16 Males

14 Females

Total number ofadults eclosed 4

0 123456789101112 Days after pupation

Figure 2-5. Developmental time from pupation to adult emergence at 23ºC, RH 87% and 14:10 hr L:D.

s 10

6 Total no.ofTotal egg 4

0 12345678910 Days after hatching

Figure 2-6. Development time from oviposition to egg hatch to at 24ºC and 14:10 hr L:D.

CHAPTER 3 FEEDING HABITS OF Tomarus subtropicus LARVAE

White grubs are the most damaging soil dwelling insects in turfgrass. Grubs typically feed either on live roots (Popillia japonica Newman, Rhizotrogus majalis (Razumowsky),

Cyclocephala spp., Phyllophaga spp.) or on soil organic matter (Cotinis nitida L., Euphoria

sepulcralis F.), damaging turf by direct feeding or tunneling (Ritcher 1966, Tashiro 1987,

Braman and Pendley 1993). Understanding the feeding preference of key grub pests helps to

explain pest distribution, abundance, development, and management using resistant turfgrasses.

Studies on the host range and behavior of phytophagous scarabs demonstrate that cool season grasses differ in their suitability as a white grub food source, and that plant chemistry and physical attributes affect their palatability (Sutherland 1971, King 1977, King et al. 1981, Potter et al. 1992, Crutchfield and Potter 1995). Grub development and growth can be significantly affected by host plant quality (King 1977, Dosdall and Ulmer 2004, Potter 1992 et al., Moreau et al. 2006). In addition, turfgrasses can mediate grub feeding behavior by attraction with plant

volatiles and certain concentrations of CO2 (King et al. 1981, Sutherland 1972), by repellency

with secondary metabolites (Sutherland et al. 1975), stimulation, or deterrence.

The host range and larval feeding habits of T. subtropicus are relatively unknown, with the exception of its ability to attack sugarcane (Gordon and Anderson 1981), bermudagrass and St.

Augustinegrass (Reinert 1979). Several other warm season turfgrass species are commonly used on residential lawns, sports fields, and golf courses, such as zoysiagrass (Zoysia spp.), common centipedegrass (Erimochloa ophiuroides [Munro] Hack), bahiagrass (Paspalum notatum Flugge) and seashore paspalum (Paspalum vaginatum Swartz), but their suitability as a food source for T. subtropicus has not been determined.

Some grub species feed on soil organic matter as first instars and then feed on roots as second or third instars (Litsinger et al. 2002). Tomarus subtropicus grubs were reported to prefer soils with high organic matter content (e.g., muck soils) in contrast with other species, which prefer sandy soils (Gordon and Anderson 1981, Hall 1987). Tomarus subtropicus females prefer soils with high organic matter content for oviposition (Cherry and Coale 1994). In contrast, I have observed T. subtropicus grubs can also occur in soils with low organic matter. It is unclear whether soil organic matter is crucial for growth and development of T. subtropicus.

In this greenhouse study, no-choice tests were designed 1) to determine the effect of soil organic matter on T. subtropicus growth and development and 2) to determine which of the warm season grasses commonly used in Florida can serve as hosts for T. subtropicus grubs.

Materials and Methods

Effect of Soil Organic Matter Content on Survival and Growth of First Instars

To determine if soil organic matter could affect first instar T. subtropicus survival and growth, St. Augustinegrass variety ‘Palmetto’ was planted in seedling trays (8×8×8 cm cells) with either sand (48 cells) or Black Velvet peat (48 cells). Twenty-four more cells had peat without grass. All grubs (2-6 days old) were weighed and individual grubs were placed 2.5 cm deep covered with soil in each cell in the peat only treatment, and in half of the peat with grass

(24 cells). The other cells remained uninfested as controls. Each cell in a seedling tray was watered daily with 30 ml of water. After 1 month, cells were visually inspected, and surviving grubs were weighed. Grass roots were washed using a #10 sieve, oven dried in paper bags for 48 hrs at 55ºC, and dry weights of roots were obtained. Data were analyzed using an ANCOVA

(SAS Institute 2000) procedure with soil type as a factor, post-treatment grub weight as a dependent variable and initial weight of grubs as a covariate, and a two-way ANOVA with soil type and grub presence as factors and dry root weight as a dependent variable.

Survival, Growth and Development of T. subtropicus Larvae on Different Warm Season Grass Species

The 2005 experiment. Seven species (six warm season and one cool season turfgrasses) were used in the test, including: ‘Palmetto’ St. Augustinegrass (Stenotaphrum secundatum

[Walt.] Kuntze), ‘Tifway’ bermudagrass (Cynodon dactylon × transvaalensis Burtt-Davy),

‘Empire’ zoysiagrass (Zoysia japonica Steud.), common centipedegrass, ‘Pensacola’ bahiagrass,

‘Sea Dwarf’ seashore paspalum, and ‘Gulf’ annual ryegrass (Lolium multiflorum Lam.). Thirty

pots (15 cm diameter) were planted with each grass species. All warm season grasses were

propagated from plugs, and ryegrass was seeded at a rate of 0.05 kg/m². FaFard mix #2 was the

soil substrate. Grass was allowed ~2 months to establish. Grass was watered daily and fertilized

monthly with 48 kg of N per ha during establishment. Pots were arranged in a randomized

complete block design in the greenhouse.

Initial grub weights were obtained, then one recently molted (<7 days) third instar was put

in a shallow depression on the soil of half of the pots for each turfgrass species. Grubs were

allowed 10 minutes to dig into the soil, and those that failed to do so were replaced. Daylight was

supplemented with lights to provide a photoperiod of 16:8 (L: D). Pots were watered with 150 ml

every other day. Grass clippings were collected weekly, placed in paper bags, fresh weights were

taken within 2 hours, and clippings were then dried for 48 hours at 55ºC, and weighed. Clipping

height was different for each grass according to management recommendations (Turgeon 2002)

(Table 3-1). Grubs were removed from the pots after 8 weeks, and survival, larval weight and

weight gain were determined. Grass roots were washed using a #20 sieve and cut no more than 2

mm from the plant crown. Washed roots were placed into paper bags, oven dried for 48 hours

(55ºC), and dry root weight was recorded.

The 2006 experiment. The test was repeated with the same warm season turfgrasses, but

annual ryegrass was excluded because the grubs had poor survival in the previous experiment

and the grass grows poorly in high summer temperatures. Grass plugs (10 cm diameter) were

obtained from the UF research unit at Citra and planted into 10 cm plastic pots. Soil used in the

experiment was collected from the same site (94% sand, 4% clay, 2% silt and 1.6% organic matter). Only apparently healthy first instars (1-3 days old) that actively moved were weighed

immediately before introduction into the pots. A cylindrical hole (8 mm diameter and 5 cm

deep) was made in the soil in each pot, one grub was put into the hole and covered with soil.

Grass was watered daily and fertilized weekly (0.6 kg of N per ha). Four replicates per each grass species were arranged. Each replication included six infested and six uninfested pots which served as controls. Grass clippings were collected weekly, placed in paper bags, fresh weights were taken within 2 hours, and clippings were dried for 48 hours at 55ºC, and weighed. Grass color and density were visually assessed on a scale from 1 (yellow, sparse) to 9 (dark green, very dense) and total grass quality score was calculated by averaging the two scores. Grubs were removed from the pots after 8 weeks, and survival, larval weight weight gain and head capsule width were determined. Grass roots were washed using a #20 sieve and cut no more than 2 mm from the plant crown. Washed roots and remaining stems, stolons and leaves were placed into paper bags, oven dried for 48 hours, and dry root weight and dry total plant yield were recorded.

Statistics. The correlation between initial and final weights was tested before analysis. If the two variables were significantly correlated, ANCOVA GLM procedure (SAS Institute 2003) was used to analyze the effect treatments on grub final weight with correction for initial weight for all experiments. Survival data were arcsine square root transformed before analysis. Dry root weights were analyzed using ANOVA (SAS Institute 2003), and clipping weights and grass

visual evaluation data were analyzed using a repeated measure analysis (SAS Institute 2003)

with time as a repeated factor.

Results

Effect of Soil Organic Matter Content on Survival and Growth of First Instars

Initial grub weight was on average the same for all treatments (0.028 g) (Table 3-2). Only

two grubs (8%) that were reared on peat without grass survived and they remained first instars.

Grubs reared on peat with grass and sand with grass had 83% and 75% survival, respectively,

and all of these grubs developed into second instars. Grub final weight was slightly higher for grubs reared on peat with grass (0.87 ± 0.07) than for grubs reared on sand with grass (0.74 ±

0.08), although the presence or absence of soil organic matter did not significantly influence grub

final weights. As expected, grub feeding significantly reduced root mass compared to uninfested

cells (F = 77.2; df = 2, 85; P < 0.0001). In addition, more roots were consumed in the cells with

sand (final weight 0.31 ± 0.05g) than in the cells with peat (final weight 0.59 ± 0.05g).

Survival, Growth and Development of T. subtropicus Larvae on Different Warm Season Grass Species

The 2005 experiment. The average third instar initial weight was 1.97 (± 0.08)g and it

did not differ statistically among the treatments (F = 0.71; df = 6, 76; P = 0. 64). Initial grub

weight and final grub weight not significantly correlated (Pearson r = 0.37, P = 0.001). The

interaction term between grass species and initial weight was not statistically significant (F =

1.32; P = 0.26), thus the interaction was excluded from the model. Differences in grub final

weight were statistically significant among grasses (F = 12.67; P < 0.0001). Further series of t- tests of LS means revealed that the final grub weight in ryegrass was significantly lower than grub weight in any of the warm season grasses (F =11.33; P < 0.0001), except bermudagrass

(Table 3-3). Grub weight gain in bermudagrass was lower than the mean weight of other warm

season grasses (F = 6.12; df = 6, 76; P < 0.001). However, 60% of the grubs survived in bermudagrass and only 40 % of the grubs survived in ryegrass (Table 3-2). Analysis of grass dry root weight with and without grubs indicated that there was a significant root reduction in all pots with warm season grasses, but not in the pots with ryegrass (Figure 3-1).

Grass leaf growth changed differently among grass species over time. For bahiagrass and

St. Augustinegrass the main effect of grub presence was significant, whereas the main effect of time and interaction between these two factors was not significant (Table 3-4). Therefore, grass leaf growth did not change significantly over time in pots with or without grubs, but pots with grubs tended to yield less clippings. Clipping yield was significantly lower by the fifth week for

St. Augustinegrass and the sixth week for bahiagrass (Figures 3-2, 3-3). The main effects of both factors (time and grub presence) were significant for centipedegrass and bermudagrass, and the interaction was not significant (Table 3-4). Clipping yield of these grasses changed significantly over time in infested and non-infested pots, however uninfested pots yielded more clippings

(Figures 3-4, 3-5). For zoysiagrass, the main effect of grub presence was not significant although the main effects of time and the interaction of time and grub presence were significant (Table 3-

4). Thus, clipping weight changed over time differently in the pots with grubs and without grubs

(Figure 3-6). Similarly for ryegrass, the main effects of both factors and their interaction were significant (Table 3-4), so grass growth decreased over time, and the decrease occurred more rapidly in the pots with grubs (Figure 3-7). Seashore paspalum clipping weight varied slightly over time, but no effects of time, grub presence or their interaction were observed (Table 3-4).

The 2006 experiment. Initial first instar weight (on average 0.0308 ± 0.01 g) did not correlate with grub final weight (on average 2.7493 ± 0.86 g) (r = 0.11, P = 0.2212), and was not included in the analysis as a covariate. ANOVA (SAS Institute 2003) was used to analyze the

effect of grass species on final grub weight and weight gain. Final grub weight differed among grasses (F = 5.02; df = 5, 109; P = 0.0004). Grubs reared on seashore paspalum gained significantly more weight (weight gain ca. 126 times) than grubs reared on other grasses, and grubs in bermudagrass weighed less (weight gain ca. 70 times) than grubs reared on other grasses

(Table 3-5). Percent survival percent of grubs that reached the third instar and head capsule width did not differ among grasses (F = 1.64; df = 5, 23; P = 0.20; F = 2.17; df = 5, 23; P =

0.1034; and F = 1.73: df = 5; 109; P = 0.1347 respectively).

Grub root feeding caused significant root reduction of all grasses (F = 70.61; df = 6, 287; P

< 0.0001) (Figure 3-8). However, the total plant yield only for bahiagrass and bermudagrass was reduced (F = 22.81; df = 11, 287; P = 0.001) (Figure 3-9). Dry clipping weight collected of all turfgrass species (except centipedegrass) was not affected by grub feeding (Table 3-6). Grass quality ratings for St. Augustinegrass, bermudagrass, zoysiagrass and centipedegrass were reduced because of grub damage, but were not affected by grub presence in bahiagrass and seashore paspalum (Table 3-7). Differences in grass quality became apparent from the fourth week (St. Augustinegrass, bermudagrass), the sixth week (zoysia), and eighth week

(centipedegrass) after grub introduction. At the final evaluation most grubs (94%) were third instars and grass crowns could be easily pulled from the pots with grubs, but grass was green and alive in all of the pots.

Discussion

The first instars of several phytophagous scarab species begin feeding on soil organic matter, then feed on roots as second or third instars (Litsinger et al. 2002). However, the presence of grass roots was more important to larval T. subtropicus survival and development than soil organic matter content, but the organic matter may provide supplemental nutrients

(King 1977). Final T. subtropicus grub weights were not affected by the presence or absence of

soil organic matter, but more roots were consumed when the grass was grown in sand than in peat. This may indicate that compensatory feeding occurs in nutrient-poor soil types (Yang and

Joern 1994, Obermaier 1999, Berner 2005). Soil type or organic matter content may also influence the ability of turfgrass to recover from root damage (Sparling et al. 2006).

Tomarus subtropicus was previously reported to feed on sugarcane (Gordon and Anderson

1981), St. Augustinegrass, and bermudagrass (Reinert 1979). However, it can also successfully survive and develop on bahiagrass, centipedegrass, zoysiagrass and seashore paspalum. Growth was better on zoysiagrass and seashore paspalum than other turfgrasses if grubs were reared from the first instar. Tomarus subtropicus can cause damage on bermudagrass golf courses (Reirnert

1979), so despite the slower grub growth, bermudagrass may still be an acceptable host. Reduced grub growth may be influenced by a smaller root system in bermudagrass. Grub movement was also limited, so grubs could not migrate in search of food after the previous source had been exhausted.

Annual ryegrass appeared to be a poor host for T. subtropicus grubs. This grass species is a cool season grass and has a C3 carbon fixation cycle, whereas all other grass species are warm season grasses with a C4 carbon fixation cycle. Differences in the morphology and physiology of these two grass types (e.g., higher nutritional quality of C3 grasses, lower digestibility of C4 grasses because of larger amount of structural defense compounds, higher plant density of C4 grasses) led to the formulation of the C3 –C4 hypothesis, which states that a C3 plant is a better quality food for herbivores than C4 plants (Caswell et al. 1973, Barbehenn and Bernays 1992,

Scheirs 2001). However, this hypothesis was not tested in relation to root feeding insects. It is possible that annual ryegrass is nutritionally poorer than the warm season grasses to which T. subtropicus is adapted. But it is more likely that the physical structure of the grass root system

caused reduced feeding. Numerous fine roots of ryegrass were distributed through the entire pot

volume, creating adense mesh which probably interfered with grub movement. At evaluation,

live grubs were in chambers that were located near pot walls and not more than 5 cm deep. In

contrast, the root systems of all other grasses had a main thick root shoot, from which roots

branched closer to the pot walls and bottom and grubs were often found in the center of the pots

at different depths.

Root loss caused by grub herbivory does not necessarily result in reduced grass growth

(Potter et al. 1992, Crutchfield and Potter 1995, Braman and Raymer 2006). Moreover, a small amount of root herbivory may provoke foliage and root growth (Hamphries 1958, Bardgett et al.

1999). In my study, grasses could tolerate feeding by third instars for a month, which caused

>50% root reduction. Increased foliage yield did not occur, but was significantly lower after 5

and 6 weeks of grub feeding for most of the grasses in 2005. However, the total plant yield at the

end of the experiment in 2006 only differed between the infested and uninfested pots planted

with bahiagrass and bermudagrass. Such factors as irrigation regime, frequency of mowing or

clipping, presence of other stresses, and grass age can contribute to foliage reduction (Ladd and

Burliff 1979, Potter 1982). Most of the feeding by third instars occurs during August and

September, which coincides with reduced rainfall and possibly reduced shoot and root growth.

Thus, damage is very pronounced during this season.

In conclusion, T. subtropicus exhibits a generalist feeding habit similar to other grub

species. All of the warm season turfgrasses that I examined are susceptible to T. subtropicus

grubs, and grubs can grow and develop successfully to third instar on all of the tested grasses.

Seashore paspalum may be more favorable for grub growth and development than other grasses tested.

Table 3-1. Clipping height for grasses used in the experiment. Grass species Common name Clipping height (cm) Cynodon dactylon × transvaalensis Bermudagrass 2.54 Erimochloa ophiuroides Centipedegrass 5.08 Lolium multiflorum Annual ryegrass 7.62 Paspalum notatum Bahiagrass 7.62 Paspalum vaginatum Sea shore paspalum 1.27 Stenotaphrum secundatum St. Augustinegrass 7.62 Zoysia japonica Zoysiagrass 5.08

Table 3-2. Tomarus subtropicus survival and mean grub and root weight in sand and peat, with and without grass Grub survival, Mean initial grub weight Mean final grub weight (g) Mean final root weight Treatment % (g) ± SEM ± SEM¹ (g)± SEM² Peat only, with grub 8 0.028 ± 0.002 0.11 + 0.03a - Grass w/peat,w/grub 83 0.028 ± 0.002 0.87 + 0.07b 0.59 + 0.05a Grass w/sand, w/grub 75 0.028 ± 0.002 0.74 + 0.08b 0.31 + 0.05b Grass w/peat - - 1.24 + 0.06c Grass w/sand - - 1.17 + 0.06c ¹ Means within columns with different letters are statistically different at α = 0.05 (F = 24.6; df = 2,71; P < 0.0001) ² Means within columns with different letters are statistically different at α = 0.05 (F = 77.2; df = 2, 85; P < 0.0001) 49

Table 3-3. Third instar T. subtropicus survival and growth on different grass species. Grass species % grub survival Initial grub weight Final grub weight (g) Weight gain (g) ± Root (g) ± SEM ± SEM¹ SEM² reduction (%) Bermudagrass 66.67 ± 6.67 1.73 ± 0.21 2.10 ± 0.19a 0.37 ± 0.27b 65 Centipedegrass 66.67 ± 24.0 1.90 ± 0.25 3.43 ± 0.12b 1.53 ± 0.24a 55 Ryegrass 40.00 ± 0.00 2.19 ± 0.72 1.96 ± 0.21a 0.06 ±0.36b 19 Bahiagrass 93.30 ± 6.67 1.81 ± 0.18 3.05 ± 0.19b 1.16 ± 0.22a 46 Seashore paspalum 86.67 ± 6.67 2.13 ± 0.19 3.57 ± 0.23b 1.51 ± 0.18a 56 St.Augustinegrass 66.67 ± 6.67 2.17 ± 0.18 3.18 ± 0.14b 0.94 ± 0.15a 53 Zoysiagrass 86.67 ± 6.67 1.95 ± 0.11 3.04 ± 0.11b 1.09 ± 0.18a 56 ¹ Means within columns with different letters are statistically different at α = 0.05 (F = 8.51; df = 6, 76; P < 0.0001) ² Means within columns with different letters are statistically different at α = 0.05 (F = 6.12; df = 6, 76; P < 0.0001) 50

Table 3-4. Statistics showing effect of time and grub feeding and their interaction on clippings collected during eight-week period of the 2005 experiment Effect of interaction of both Effect of time Effect of grub feeding factors Grass F df P F df P F df P Bahiagrass 5.28 7, 22 <0.01 5.11 1, 28 0.03 0.86 7, 22 0.55 Bermudagrass 45.48 7, 22 <0.01 12.68 1, 28 <0.01 1.05 7, 22 0.43 Centipedegrass 3.55 7, 22 0.01 4.3 1, 28 0.05 1.11 7, 22 0.39 Ryegrass 28.9 7, 22 <0.01 9.38 1, 28 0.02 3.21 7, 22 0.07 Seashore paspalum 3.15 7, 22 0.19 0.33 1, 28 0.57 0.66 7, 22 0.70 St. Augustinegrass 2.34 7, 22 0.06 10.26 1, 28 <0.01 2.29 7, 22 0.06 Zoysiagrass 5.68 7, 22 <0.01 1.64 1, 28 0.21 3.63 7, 22 0.01

Table 3-5. Tomarus subtropicus survival, development and growth on different species of warm season grasses when reared from the first instar. Grass species % grub % of grubs 3rd instar head Initial grub Final grub weight Grub weight gain survival reaching 3rd capsule width, weight (g) ± (g) (final weight/initial instar mm ± SEM SEM ± SEM¹ weight)² Bahiagrass 79.3 ± 7.9 75.0 ± 0.70 7.73 ± 0.05 0.028 ± 0.002 2.62 ± 0.20ab 103.7 ± 11.2ab

Bermudagrass 66.8 ± 6.7 46.0 ± 10.5 7.91 ± 0.08 0.034 ± 0.003 2.20 ± 0.24b 69.60 ± 8.3b Centipedegrass 91.5 ± 4.9 91.5 ± 4.90 7.51 ± 0.09 0.030 ± 0.002 2.43 ± 0.21b 86.10 ± 0.7ab Seashore paspalum 66.8 ± 6.7 66.8 ± 6.70 7.75 ± 0.06 0.031 ± 0.003 3.35 ± 0.20a 126.3 ± 15.6c St. Augustinegrass 87.3 ± 4.3 75.0 ± 10.7 7.88 ± 0.07 0.033 ± 0.002 2.62 ± 0.21ab 92.60 ± 7.9ab Zoysiagrass 70.8 ± 7.9 70.8 ± 7.90 7.83 ± 0.06 0.030 ± 0.002 3.13 ± 0.13a 113.9 ± 8.7ac ¹ Means within columns with different letters are statistically different at α = 0.05 (F = 5.08; df = 5, 109; P = 0.003) ² Means within columns with different letters are statistically different at α = 0.05 (F = 3.95; df = 5, 109; P= 0.002)

Table 3-6. Statistics showing effect of time and grub feeding and their interaction on clippings collected during eight-week period of the 2006 experiment. Grass species Grub Time Time*Grub F df P F df P F df P Bahiagrass 2.89 1, 46 0.10 8.47 3, 44 <0.01 1.65 3, 44 0.19 Bermudagrass 2.72 1, 46 0.11 10.31 3, 44 <0.01 2.87 3, 44 0.05 Centipedegrass 2.87 1, 46 0.02 9.69 3, 44 <0.01 0.26 3, 44 0.85 Seashore paspalum 0.01 1, 46 0.93 4.61 19.74 <0.01 0.66 3, 44 0.58 St. Augustinegrass 3.63 1, 46 0.06 4.78 3, 44 0.07 2.00 3, 44 0.13 Zoysiagrass 3.02 1, 46 0.06 1.63 3, 44 0.20 0.41 3, 44 0.87

Table 3-7. Statistics showing effect of time and grub feeding and their interaction on turfgrass quality in 2006. Grass species Grub Time Time*Grub F df P F df P F df P Bahiagrass 0.6 1, 46 0.44 3.90 3, 44 0.15 0.95 3, 44 0.43 Bermudagrass 9.08 1, 46 <0.01 17.07 3, 44 <0.01 5.54 3, 44 <0.01 Centipedegrass 5.71 1, 46 0.02 13.51 3, 44 <0.01 8.66 3, 44 <0.01 Seashore paspalum 0.04 1, 46 0.83 4.61 3, 44 0.01 0.37 3, 44 0.70 St. Augustinegrass 9.08 1, 46 <0.01 17.07 3, 44 <0.01 5.54 3, 44 <0.01 Zoysiagrass 32.23 1, 46 <0.01 14.08 3, 44 <0.01 9.82 3, 44 <0.01

4.5 with grub a without grub 4 a b 3.5 b b 3

2.5

a 2 a b b a b Dry rootDry weight, g

55 1.5

a 1 a a* 0.5

0 Bermudagrass Centipedegrass Ryegrass Bahiagrass Sea shore St. Zoysiagrass grass species paspalum Augustinegrass Figure 3-1. Reduction of root mass caused by third instar T. subtropicus feeding, 2005.

1.4

Pots with grubs 1.2 Pots without grubs a 1 b a a b b b 0.8 a a a a a

0.6 a a a a Clippings dry weight, g 56 0.4

0.2

12345678

Week after grub introduction

Figure 3-2. Dry weight of St. Augustinegrass clippings collected weekly for 8 weeks, 2005.

1.2

1 Pots with grubs Pots without grubs b b 0.8 a a a a b a a a a 0.6 a a a a a Clipping dry weight, g 0.4 57

0.2

12345678 0 Week after grub introduction

Figure 3-3. Dry weight of bahiagrass clippings collected weekly for 8 weeks, 2005.

0.7

0.6 Pots with grubs b b Pots without grubs

0.5 a b a b a a a a 0.4 a a a a a a

0.3 Clippings dry weight, g 58 0.2

0.1 12345678 Weeks after grub introduction 0

Figure 3-4. Dry weight of centipedegrass clippings collected weekly for 8 weeks, 2005.

0.9

0.8 Pots with grubs Pots without grubs b 0.7 a a 0.6

0.5 a b a a 0.4 a a

a a 0.3

Clippings dry weight, g a a 59 a b 0.2 a

0.1 12345678 Week after grub introduction 0

Figure 3-5. Dry weight of bermudagrass clippings collected weekly for 8 weeks, 2005.

0.9 Pots with grubs b Pots without grubs 0.8 a a a 0.7 a b aa a

0.6 a b a 0.5 a

a a 0.4 a Dry clippingDry weight, g 60 0.3

0.2

0.1 12345678 Week after grub introduction 0

Figure 3-6. Dry weight of zoysiagrass clippings collected weekly for 8 weeks, 2005.

1.6

1.4 aa Pots with grubs Pots without grubs

1.2 a a

a 0.8 b b a b a a 0.6 a a a a

Weightof grass clippings, g a 61 0.4

0.2 12345678 Week after grub introduction 0

Figure 3-7. Dry weight of ryegrass clippings collected weekly for 8 weeks, 2005.

7 Pots with grubs Pots without grubs b 6 b

b 5 a b 4 a b 3

Dryroot weight, g a

62 b 2 a

a 1 a Bahiagrass Bermudagrass Centipedegrass Seashore St. Augustinegrass Zoysiagrass Paspalum

Figure 3-8. Reduction of root mass caused by third instar T. subtropicus feeding, 2006.

16 Pots with grubs Pots without grubs a a

14 a

a 12 b b a a

10 a a¹ a a 8 Dry plant weight, g 63 6

Bahiagrass Bermudagrass Centipede Seashore paspalum St.Augustinegrass Zoysiagrass 0

Figure 3-9. Effect of grub feeding on total plant yield by warm season grasses, 2006

CHAPTER 4 MANAGEMENT OF Tomarus subtropicus

Introduction

White grubs that infest turfgrasses are commonly managed using insecticides (Potter

1998). Preventive insecticides (e.g., neonicotinoids, insect growth regulators) are typically

applied during oviposition or shortly after peak egg hatch. Curative insecticides traditionally are

applied after egg hatch, but they may control second and third instars when damage becomes

obvious (Potter and Held 2002). Close monitoring is required to properly time curative control

applications (Potter 1998). The efficacy of these insecticides and other management tools has

been tested for major pests such as the Japanese beetle (Cowles and Villani 1996, Cowles et al.

1999, Grewal et al. 2001, Mannion et al. 2001, Potter and Held 2002), northern masked chafers

(Grewal et al. 2001), European chafer (Cowles and Villani 1996, Cowles et al.1999), May / June

beetles (Cranshaw and Zimmerman 1989), Oriental beetle (Cowles and Villani 1996, Cowles et

al. 1999) and Asiatic garden beetle (Cowles et al. 1999) in cool season turfgrasses. However,

little is known about the management of white grubs in warm season grasses within the southeastern United States.

Tomarus subtropicus has been difficult for turfgrass pest managers in Florida to control.

The application timing of preventive insecticides has been based on recommendations from related studies in sugarcane. Despite the use of preventive insecticides, second and third instar feeding damage in St. Augustinegrass lawns becomes apparent as early as August, which often necessitates a curative insecticide application. It is possible that either the preventive insecticides typically used against white grubs are less effective against T. subtropicus or the application timing has been wrong.

The preventive grub insecticides clothianidin, imidacloprid, and thiamethoxam are neonicotinoids and halofenozide is an insect molt accelerator. Neonicotinoids are the most widely used class of insecticides, their sales accounting for 11-15% of the total insecticide sales

worldwide (Tomizawa and Casida 2005). They are known to have strong insecticidal properties

yet are relatively safe to vertebrates, acting selectively on insects’ nicotinic acetylcholine

receptor, blocking it irreversibly (Brown et al. 2006, Ihara et al. 2006). Neonicotinoids are water

soluble, are quickly absorbed by plants, have prolonged (up to 3 months) residual activity, and

are systemic. They act if ingested, but have poor contact toxicity (Tomizawa and Casida 2003,

2005). Their persistence in the soil varies depending on soil type and temperature, but could be

as long as one year in Minnesota and about 3 months in turfgrass in Georgia (Cox 2001).

Halofenozide is a systemic insect growth regulator with no stomach or contact activity (Ware

and Whitacre 1993). It mimics the action of the insect molting hormone 20–hydroxyecdysone,

promoting premature molting which results in the loss of hemolymph and molting fluids, and

subsequently causes larval death from desiccation (Gardner and Branham 2001).

Insecticides used for curative control are broad-spectrum with short residuals, and include

organophosphates and carbamates. Many insecticides of these classes were canceled or uses

were restricted by the 1996 Food Quality Protection Act. Trichlorfon and carbaryl are the only

insecticides of these classes which are still available for curative control of white grubs in

turfgrass. Both carbaryl and trichlorfon act by inhibiting cholinesterase, but carbaryl has lower

dermal and oral mammalian toxicity than most organophosphates (Ware and Whitacre 1993).

Entomopathogenic nematodes have been an alternative to insecticides in insect management systems. The insecticidal ability of nematodes was first discovered in 1929, when

Steinernema glaseri (Steiner) was described and tested against Japanese beetle larvae (Glaser

and Fox 1930). Since then, nematodes have been used to control >200 insect species, such as

armyworms, cat fleas, cutworms, filth flies, flea beetles, German cockroaches, leaf miners, mole

crickets, phorid flies, root weevils, stem borers, webworms and white grubs (Klein 1993, Georgis

et al. 2006). Successful control with nematodes has been achieved only in limited systems: the

Diaprepes root weevil (Diaprepes abbreviatus L.) in citrus, the black vine weevil (Otiorhynchus

sulcatus F.) in cranberries, mint and greenhouses, billbugs (Sphenophorus spp.) in turf,

mushroom flies (Bradysia spp.) in mushrooms, artichoke plume moth (Platyptillia carduidactyla

(Riley)), and cockroaches in urban industrial environments (Georgis et al. 2006). Heterorhabditis spp., S. glaseri, and S. scarabei (Stock and Koppenhöfer) are the most effective nematode species against white grubs in turf (Klein 1993; Cappaert and Koppenhöfer 2003, Stock and

Koppenhöfer 2003; Koppenhöfer and Fuzy 2003 a, b; Koppenhöfer et al. 2006). However, white grub species and larval instars differ in their susceptibility to entomopathogenic nematodes

(Grewal et al. 2001, Koppenhöfer and Fuzy 2003, Koppenhöfer et al. 2004).

The objective of this study was to evaluate the efficacy of preventive and curative insecticides and entomopathogenic nematodes against T. subtropicus in a series of laboratory, greenhouse, and field tests.

Material and Methods

Evaluation of Insecticides for Preventive Control of T. subtropicus

Greenhouse experiment. Five insecticides were used in a greenhouse trial: imidacloprid

(Merit® 75 WP, Bayer Environmental Science, Research Triangle Park, NC), halofenozide

(Mach 2®, Dow AgroScience, Indianapolis, IN), clothianidin (Arena® 50WDG, Arysta Life

Science, San Francisco, CA), and imidacloprid plus bifenthrin (Allectus® SC, Bayer

Environmental Science, Research Triangle Park, NC) with four replications and a control (total:

24 pots). Plastic pots (15 cm diameter) were filled with potting mix FaFard #2 and St.

Augustinegrass variety ‘Palmetto’ was planted and allowed 4-5 weeks to establish.

Four active and apparently healthy first instars (1-3 days old) were placed individually into holes (7 mm in diameter, 10 cm deep) in each pot and covered with soil. Grubs were allowed to acclimate for 48 hours. Treatments were applied with a 1-m long, two-nozzle boom sprayer.

After 3 weeks the soil in the containers was inspected and grub survival was recorded. Maximum daily temperature averaged 37.9°C ± 1.4°C and minimum daily temperature averaged 25.1°C ±

1.9°C in the greenhouse. Data were analyzed by analysis of variance (ANOVA) for effects of type of insecticide.

Field test. First instar T. subtropicus were reared from eggs, and held 2-6 days in 20-ml

Solo cups containing soil and St. Augustinegrass in the laboratory at room temperature. Only apparently healthy, feeding and actively moving grubs were selected for the experiment. The formulated insecticides that were tested included imidacloprid (Merit® 75 WP), halofenozide

(Mach 2®), clothianidin (Arena®50WDG), and imidacloprid plus bifenthrin (Allectus®SC) and two experimental products NUP 06026 (Nufarm) and DPX E2Y45 (Dupont) (Table 4-2). The experiment was conducted at the U.F. Plant Science Unit in Citra, FL, from 14 June to 21 July

2006. A PVC ring (30 cm diameter, 10 cm high) was inserted 7 cm deep in the center of each St.

Augustinegrass variety ‘Floratam’ plot (1×1 m). Six holes 5 cm deep were made inside each ring and one grub was placed into each and covered with the soil. PVC rings were enclosed with white mesh to prevent escape or predation. Each treatment had four replications.

Treatments were applied 24 hours after grub introduction with a 1-m long, two nozzle boom sprayer. Control plots remained untreated. All plots were irrigated (0.64 cm) after

application. Rings were removed 30 days after application and soil and grass were destructively

sampled. The number of live, dead and morbid grubs was recorded.

The thatch layer was 1.7 cm, mowing height was 7.5 cm, air temperature during

application was 31ºC, soil temperature was 20ºC and relative humidity fluctuated 34% - 62%.

Constant wind speed was up to 3 km/ h with gusts up to 8 km/h.

Curative Control of T. subtropicus.

A field trial was conducted at the U.F. Plant Science Unit in Citra, FL, in October 2005 to

determine the curative potential of clothianidin, halofenozide, carbaryl, and trichlorfon. A PVC ring (30 cm diameter, 10 cm high) was inserted 7 cm deep in the center of each St.

Augustinegrass variety ‘Floratam’ plot (1×1 m). To minimize cannibalism, only three field-

collected third instar T. subtropicus were placed in each ring, after being held in pots of St.

Augustinegrass variety ‘Palmetto’ in the greenhouse for at least 2 wk to ensure health. Grubs

were placed on the soil surface and allowed 10 minutes to burrow in the soil. Grubs that failed to

burrow within 10 minutes were replaced. PVC rings were enclosed with white mesh to prevent

escape or predation. Each treatment had six replications.

Treatments were applied 48 hours after grub introduction with a 1-m long, two nozzle boom sprayer. Control plots remained untreated. All plots were irrigated (0.64 cm) after application. Rings were removed 7 days after treatment and soil and grass were destructively sampled. The number of live, dead and morbid grubs was recorded.

Air temperature was 27.5ºC at the beginning of application and 30.8ºC at the end of application, relative humidity was 70% at the beginning and 67% in the end, and soil temperature was 22.2ºC at a 10 cm depth. Wind speed began <3 km/h with gusts up to 9.7 km/h,

and by the end of application, wind speed was ~ 4.8 km/h with gusts 8.9 km/h. Grass height was

9.25 cm, and thatch thickness was 0.75 cm.

Statistics. All treatment mortality data were corrected for the control mortality using

Abbott’s formula (Abbott 1925) and arcsine square-root transformed before analysis. Data were

analyzed using one-way ANOVA (SAS Institute 2000) and means were separated using Tukey’s

test (α = 0.05).

Infectivity of Five Nematode Species against Second and Third Instars.

A laboratory experiment was conducted to compare the effectiveness of five nematode

species (Heterorhabditis bacteriophora (Poinar), H. zealandica (Poinar), H. megidis (Poinar), S.

glaseri, and S. scarabei against T. subtropicus second and third instars which were field collected

and held in colony >2 weeks before the test to ensure health. Heterorhabditis zealandica was

obtained from K. Nguyen’s colony at the University of Florida; Becker Underwood provided H.

bacteriophora and H. megidis; S. glaseri and S. scarabei were acquired from A. Koppenhöfer’s

colony at Rutgers University. Nematodes were stored for <30 days before use in the test.

Individual grubs were placed into 266-ml plastic Solo cups (53 cm²) containing sifted, autoclaved soil (95% sand, 5% clay with 4.4% organic matter content) and St. Augustinegrass variety ‘Palmetto’ plugs. Each treatment was applied at a rate of 500 IJ per grub, in 50 ml of distilled water, 48 h after grub introduction to the cups. The control cups received water only.

Each treatment was replicated four times with five grubs per replicate. Cups were held at room temperature (23-24ºC) and a photoperiod of 14:10 (L:D). Grub mortality and nematode presence were assessed 14 days after treatment. Data were analyzed using a two-way ANOVA (SAS

Institute 2000) and means were separated using Tukey’s test (α = 0.05).

Reproduction of S. scarabei and S. glaseri in Third Instar T. subtropicus.

Individual grubs were placed into 69-ml plastic Solo cups (53 cm²) containing sifted, autoclaved soil (95% sand, 5% clay with 4.4% organic matter content). Steinernema glaseri and

S. scarabei were applied with the rates 100, 200 or 500 IJ per grub with 5 ml of water. Cups were held at room temperature (23-24ºC) and a photoperiod of 14:10 (L:D). Cups were arranged in a completely randomized block design with 20 cups per treatment. Cups were checked daily for grub mortality. Nematodes from the cadavers were collected using emergence traps described by Koppenhöfer and Fuzy (2003), which consisted of lids of 3.5 ×1 cm Petri dish lined with filter paper and placed in a 15 × 1.5 cm Petri dish with 25 ml of tap water. Traps were observed daily until 7 days after the first nematode emergence. The number of emerged infective juveniles was estimated by counting five subsamples from the nematode suspension collected from each grub.

Results

Insecticide Efficacy

Grub mortality was significantly greater than the control for all of the preventive insecticides, except Allectus (F = 7.96; df = 4, 15; P = 0.0012) in the greenhouse test (Table 4-

1). Clothianidin and halofenozide provided 85% control and imidacloprid 70% control. Grubs in the halofenozide treatment had distinctive cuticle deformities and could be distinguished from other treatments.

All treatments in the preventive field test were statistically different from the control (F =

22.46 df = 8, 35; P = 0.0002) (Table 4-2), but high control mortality weakened the strength of the results. Carbaryl and trichlorfon killed significantly more (61% and 58%, respectively) third instar T. subtropicus (F = 8.36; df = 4, 29; P = 0.0006) than clothianidin and halofenozide in the curative field test (Table 4-3).

Nematode Infectivity against Second and Third Instar T. subtropicus

Nematode species, grub life stage and their interaction had significant effects on mortality

(F = 37.06; df = 11, 47; P < 0.001) (Figure 4-1). Grub mortality caused by all nematode species significantly differed from the control for second instars (F = 37.29; df = 5, 23; P < 0.001). The most virulent species was S. scarabei. Steinernema glaseri and H. bacteriophora caused more than 50% mortality, whereas mortality caused by H. megidis and H. zealandica was 45%.

Significantly more third instars treated with H. bacteriophora, S. glaseri and S. scarabei died,

compared to the control (F = 72.53; df = 5, 23; P < 0.001) (Figure 4-1). Second instars were more susceptible than third instars to all nematode species, except S. scarabei, which was

equally effective against both. None of the grubs in the control treatments died.

Reproduction of S. scarabei and S. glaseri in third instar T. subtropicus

The number of S. glaseri infective juveniles collected (on average 148,623 IJs from each

grub) was higher than the number of S. scarabei IJs (on average 81,022 IJs per grub) (F = 12.36;

df = 1; 90; P = 0.0007). Application rate did not significantly affect nematode productivity (F =

2.58, df = 2, 90; P = 0.0814) (Figure 4-2). Infective juveniles emerged from 65% (100 IJ

applied), 50% (200 IJs applied), and 80% (500 IJs applied) of grubs treated with S. scarabei, and

from 75% (100 IJ applied), 85% (200 IJs applied) and 95% (500 IJs applied) of grubs treated

with S. glaseri.

Discussion

Preventive insecticides can provide control of first instar T. subtropicus if the grubs are

exposed to the products and applications are correctly timed under controlled conditions. Many

interrelated factors can influence effectiveness of insecticides against soil insects under field

conditions, such as insecticide formulation, physical and chemical properties, soil parameters,

climatic conditions, insect physiology, phenology and behavior, and application error (Harris

1972, Villani and Wright 1990).

Insecticide mobility in the soil can affect whether or not an active ingredient can reach its target pest. Poor water solubility may decrease the dispersion of insecticides in soil, whereas a highly water soluble insecticide may pass beyond the root zone (Villani and Wright 1990).

Imidacloprid, which is commonly used as a preventive grub insecticide, is water soluble with high mobility. These qualities make the compound easy for a plant to absorb and be systemic. On the other hand, it increases the leaching and dissolved run-off potential of imidacloprid (Cox

2001). High rainfall (up to 35 cm each month) in southern Florida during June and July could contribute to the rapid wash out of the insecticide from the turfgrass zone, and frequent mowing and clipping removal could eliminate the insecticide from within the turfgrass. Many sites infested with T. subtropicus are waterlogged or saturated with water and have slopes which, according to the product label, interfere with adequate active ingredient distribution and the vertical penetration of the insecticide.

Other studies have suggested that T. subtropicus is hard to control with insecticide rates that are lethal for other grub species (Reinert 1979, Watve et al. 1981). In the test of curative products even the most effective curative insecticides, carbaryl and trichlorfon, provided only up to 67% of control at their highest recommended rates. The lethal dose may be higher than for other smaller white grub species infesting turfgrass in Florida, but I did not test this. The size of

T. subtropicus grubs could be one explanation for this. The average weight of each of the T. subtropicus instars was 6 times as much as the average weight of the corresponding larval instars for Cyclocephala parallela (Cherry 1985). Moreover, adult flight and the majority of eggs and

first instars occur from July to August. If a preventive insecticide is applied about 1-1.5 months

earlier, the residual activity may be insufficient to kill large first instars.

The thick cuticle and physiology of the grub may be responsible for reduced susceptibility to insecticides. Reduced susceptibility to halofenozide and some organophosphates has been documented previously for European chafer, Rhizotrogus majalis (Razumowsky) (Villani et al.

1988, Cowles and Villani 1996, Cowles et al. 1999). To control the European chafer, twice the rate needed to control Popillia japonica (Newman) and Exomala orientalis (Waterhouse) is needed (Cowles and Villani 1996). European chafer is the only grub species that has exhibited localized resistance to cyclodiene insecticides (Tashiro et al. 1971). According to Harris (1972), the use of persistent insecticides favors resistance development in species with more than one generation per year, but use of insecticides with short residual activity (e.g., organophosphates and carbamates) favors the development of resistance by species with life cycles of one or more years.

Understanding how insects might behaviorally respond within an insecticide test is difficult. If insects are caged, they cannot escape and inevitably are contacted by the insecticides.

If tests are conducted on natural field populations, insects may be able to escape or avoid contact

with the treatments. Soil dwelling insects rely a lot on chemical cues in host finding because

their visual orientation is limited in the soil (Villani and Wright 1990, Villani et al. 1999). They are sensitive to CO2 (Galbreath 1988) and plant metabolites (Johnson and Gregory 2006), and insects are capable of distinguishing between positive and adverse olfactory stimuli (Scherer et al. 2003, Unoki et al. 2005, Liu and Davis 2006). Japanese beetle grubs can avoid contact with the entomopathogenic fungus Metarhizium anisopliae (Sorokin) by moving horizontally and vertically away from a treated area (Villani et al. 1994). Vertical movement of T. subtropicus

grubs was noticed during soil sampling; grubs were observed 10-15 cm deep in the soil in the

curative insecticides test.

Nematodes can be effectively used as an alternative or addition to chemical control of T.

subtropicus. The large size of the grubs provides the nematodes with enough resources for

reproduction and makes it easier to enter the host through the natural openings and recognize and

find the host (Bedding and Molyneux 1982, Wang and Gaugler 1998, Koppenhöfer and Fuzy

2004). Steinernema scarabei, which has shown high virulence to other grub species

(Koppenhöfer and Fuzy 2003b), caused the greatest mortality of T. subtropicus second and third

instars. However, a thick cuticle and strong immune system might reduce the susceptibility of

third instar T. subtropicus to Heterorhabditis spp. Second instars were more susceptible than

third instars to all nematode species, except S. scarabei, which was equally effective against

both. Effect of life stage on nematode infectivity varies among species (Koppenhöfer and Fuzy

2004). Early instars of Melolontha melolontha L., E. orientalis, and Phyllopertha horticola (L.) were reported to be more susceptible to Heterhabditis spp. and S. glaseri (Deseo et al. 1990,

Smits et al. 1994, Koppenhöfer and Fuzy 2004), whereas with S. scarabei and E. orientalis the opposite trend is observed, and in many cases there are no differences between instars in grub susceptibility to nematodes (e.g., Popillia japonica) (Koppenhöfer and Fuzy 2004). The mechanisms affecting the susceptibility of different grub developmental stages to nematodes are not fully understood, but insect behavior, maturity of immune system, size and cuticle thickness, in combination with nematode foraging strategies and ways to penetrate into an insect body, are possible factors (Smits et al. 1994, Koppenhöfer and Fuzy 2004).

In conclusion, for preventive control of T. subtropicus first instars, clothianidin, halofenozide and imidacloprid can be effectively used. Trichlorfon and carbaryl can be effective

against third instars and used as curative insecticides. Timing of insecticide application can be adjusted to achieve better control with these insecticides. Preventive insecticides are more likely to succeed from mid June to early July when a majority of eggs and first instars are present. An early curative application (late July - August) before damage becomes obvious could be more effective than waiting until turf damage is severe. The nematodes S. glaseri, S. scarabei and H. bacteriophora could be applied in August to target second instars.

Table 4-1. Effectiveness of four insecticides tested for preventive control against T. subtropicus grubs in the greenhouse Treatments AI Rates(kg AI/ha) Mean no. of dead grubs % mortality± SEM % corrected mortality² ± SEM Allectus SC Imidacloprid + 0.48 1.0 ± 1.4a¹ 25.0 ± 35.4 12.5 ± 12.50a³ bifenthrin 0.23 Arena 50WDG Clothianidin 0.28 3.5 ± 0.6b 87.5 ± 14.4 85.4 ± 8.60b Mach 2SC Halofenozide 4.88 3.5 ± 0.6b 87.5 ± 14.3 85.4 ± 8.60b Merit 75WP Imidacloprid 0.45 3.0 ± 1.5b 75.0 ± 28.9 70.7 ± 17.2b Control - - 1.0 ± 0.8a 25.0 ± 20.4 0a ¹ Means within columns with different letters are statistically different at α =0.05 (F = 7.96; df = 4, 19; P = 0.0012). ² Abbott’s formula was used to correct for control mortality. ³ Means within columns with different letters are statistically different at α =0.05 (F = 12.56; df = 4, 19; P = 0.0001)

Table 4-2. Mortality of first instar Tomarus subtropicus caused by selected insecticides in the field test. Product Formulation Rate,L/ha % mortality % corrected mortality¹ DPX E2Y45 1.67 SC 0.58 75 47**³ DPX E2Y45 1.67 SC 1.17 100 100** DPX E2Y45 1.67 SC 2.34 100 100** Merit 75 WP 0.47 100 100** Mach 2 2 SC 9.35 100 100** Arena 50 WDG 0.58 100 100** Allectus SC 5.26 100 100** NUP 06026 FF 1.91 100 100** Control ~ 54 0 ¹ Abbott's formula was used to correct for control mortality. ² Means marked with * are statistically different from the control (without Abbott's correction) at α = 0.05 (F = 5.94; df =8, 35; P = 0.0002) .³ Means marked with ** are statistically different from the control (with Abbott's correction) at α = 0.05 (F = 22.46; df =8, 35; P = 0.0001)

Table 4-3. Effectiveness of four insecticides tested for curative control against T. subtropicus. Treatment AI Rates Mean # of dead % Mortality ± % Corrected (kg AI /ha) grubs SEM Mortality² ±SEM Arena Clothianidin 0.28 0.7 ± 0.3 22.2 ± 27.2 11.1 ± 11.1 50WDG Dylox 80 Trichlorfon 11.44 2.0 ± 0.2*¹ 66.7 ± 21.1 61.1 ± 9.3**³ Mach 2SL Halofenozide 4.88 0.7 ± 0.2 22.2 ± 17.2 11.1 ± 7.0

Sevin SL Carbaryl 9.15 2.0 ± 0.4* 66.7 ± 29.8 58.3 ± 16.0** Control - - 0.5 ± 0.2 16.7 ± 18.2 0 ¹ Means followed by * differ statistically from control mean at α = 0.05 (F = 7.08; df = 4, 29; P = 0.0006). ² Mortality was corrected using Abbott’s formula.³ Mean marked with ** differ statistically from the control at α = 0.05 (F = 8.36; df = 4, 29; P = 0.0002)

a´ 100 A

90 2nd instar A 3rd instar 80 b

70 A bc 60 c c 50 b´ A

40 A b´

Grub mortality, % Grub mortality, B 30

79 B 20 c´ 10

c´ d c ´ 0 B

S. glaseri S. scarabei H. bacteriophora H. megidis H. zealandica Control

Nematode species Figure 4-1. Infectivity of entomopathogenic nematodes against T. subtropicus second and third

¹ Means marked with different letters differ statistically at α = 0.005 (F = 37.29; df = 5, 23; P< 0.001) ² Letter with ´ represent differences among treatments for third instars (F = 72.53; df = 5, 23; P< 0.001) ³ Capital letters indicate statistically significant differences between instars within a treatment

250000

200000 S.scarabei

S.glaseri

150000

100000 No.of nematodes collected nematodes No.of 80

50000

100 200 500 Rates of nematodes applied 0

Figure 4-2. Productivity of S. scarabei and S. glaseri reared on third instar T .subtropicus.

CHAPTER 5 CONCLUSIONS

As a result of this work, I have confirmed that T. subtropicus has a univoltine life cycle,

and its development on turfgrass appears similar to its growth on sugarcane in most aspects. The

major difference is that peak adult activity in St. Augustinegrass lawns occurs one month later

than in sugarcane (mid-July to mid-August vs. May to June, respectively). This is important from

a management perspective because preventive insecticides (e.g., halofenozide, imidacloprid) that

commercial applicators have used in May and June do not seem to provide the necessary control

of T. subtropicus larvae. A later application, in early July, may increase T. subtropicus larval

mortality and reduce turfgrass damage in late summer and early fall. Adult activity in the soil

coincided with increased rainfall and flight peaked about one month afterwards, so applicators

may be able to use rainfall as an indicator to time a preventive application. Turfgrass managers

would like to treat grubs earlier in the season to also control Cyclocephala spp. and Phyllophaga

spp., but two preventive applications may be needed for maximum turf protection where all of

these pests may coincide.

Curative applications against third instar T. subtropicus with either carbaryl or trichlorfon

are still effective control options, but I suggest monitoring lawns in August to determine grub

presence and decide if treatment is worthwhile. Several entomopathogenic nematodes could also

be considered in a grub management program, but only H. bacteriophora and H. megidis are

commercially available of all the nematode species I tested. Because second instars appear to be

more susceptible to nematodes, an application should be applied in August or early September.

Tomarus subtropicus was previously reported to feed on the roots of sugarcane, St.

Augustinegrass, and bermudagrass, but I have expanded its known host range to include centipedegrass, zoysiagrass, bahiagrass, and seashore paspalum. Varietal differences remain

unknown. One third instar can feed in a 6-inch diameter area for 5-6 weeks before turf vigor obviously declines. The turf can remain green and healthy-looking for weeks if adequately watered and fertilized, but lack most of its root system. Any additional stress, such as drought, can then kill the turf quickly. This reinforces the need for monitoring grub density in August, before the rainy season in Florida ends, because most lawn damage is apparent by the end of

September and seems to occur virtually overnight.

I enjoyed working on this project because I had an opportunity to learn different skills for laboratory and field research. The project involved knowledge of insect ecology, seasonal phenology, insect-host interactions, insect natural enemies and chemical and biological control.

My field work improved my understanding of the residential lawn ecosystem and the practical needs of the lawn care industry. Information obtained as a result of my work can improve management of the species. However, some of my attempts were not successful. My observations on adult flight and behavior in the field were conducted when beetles were active in the soil, but were too early to observe for nightly flight; nocturnal observations might have yielded new information if done in mid-July. The field test to evaluate preventive insecticides where I introduced grubs into the plots, did not work well because high mortality of the grubs in the control plots. The first instars may have been susceptible to environmental stresses and/or predation. A field test on naturally occurring populations could have been done instead, although grub density would be variable and finding good research sites and cooperators is difficult.

Knowing what I know now, I wish I had done more observations on beetle behavior in the field and laboratory, conducted a preventive control field test at several application times (May, June and early July) and tested several ornamental grasses and plants as potential hosts in addition to turfgrasses.

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BIOGRAPHICAL SKETCH

Olga Kostromytska was born, grew up and attended school in Khmelnytsky, Ukraine. She studied psychology in the Chernovtsy State University, where she obtained her Specialist degree in 1997. Olga continued studying at the department of Social and Developmental Psychology of the Kiev National University and worked simultaneously as counseling psychologist at Kiev

Gymnasium. She moved to the United States in 2002. She became involved in landscape entomology in August 2004 working part time as a technician at Landscape Entomology Lab under Dr. Buss’s supervision. She started studying toward her master’s degree in January 2005.