University of Nebraska - Lincoln DigitalCommons@University of Nebraska - Lincoln

Dissertations and Student Research in Entomology Entomology, Department of

8-2018 CHARACTERIZATION OF THE NATURAL ENEMY COMMUNITY, WITH EMPHASIS ON ENTOMOPATHOGENS, FOR MANAGEMENT OF WESTERN CORN ROOTWORM (DIABROTICA VIRGIFERA VIRGIFERA) IN WEST CENTRAL NEBRASKA Camila Oliveira-Hofman University of Nebraska-Lincoln

Follow this and additional works at: http://digitalcommons.unl.edu/entomologydiss Part of the Entomology Commons

Oliveira-Hofman, Camila, "CHARACTERIZATION OF THE NATURAL ENEMY COMMUNITY, WITH EMPHASIS ON ENTOMOPATHOGENS, FOR MANAGEMENT OF WESTERN CORN ROOTWORM (DIABROTICA VIRGIFERA VIRGIFERA) IN WEST CENTRAL NEBRASKA" (2018). Dissertations and Student Research in Entomology. 54. http://digitalcommons.unl.edu/entomologydiss/54

This Article is brought to you for free and open access by the Entomology, Department of at DigitalCommons@University of Nebraska - Lincoln. It has been accepted for inclusion in Dissertations and Student Research in Entomology by an authorized administrator of DigitalCommons@University of Nebraska - Lincoln.

CHARACTERIZATION OF THE NATURAL ENEMY COMMUNITY, WITH EMPHASIS ON

ENTOMOPATHOGENS, FOR MANAGEMENT OF WESTERN CORN ROOTWORM

(DIABROTICA VIRGIFERA VIRGIFERA)

IN WEST CENTRAL NEBRASKA

by

Camila Oliveira-Hofman

A DISSERTATION

Presented to the Faculty of

The Graduate College at the University of Nebraska

In Partial Fulfillment of Requirements

For the Degree of Doctor of Philosophy

Major: Entomology

Under the Supervision of Professors Julie A. Peterson and Lance J. Meinke

Lincoln, Nebraska

August, 2018

CHARACTERIZATION OF THE NATURAL ENEMY COMMUNITY, WITH EMPHASIS ON

ENTOMOPATHOGENS, FOR MANAGEMENT OF WESTERN CORN ROOTWORM

(DIABROTICA VIRGIFERA VIRGIFERA)

IN WEST CENTRAL NEBRASKA.

Camila Oliveira-Hofman, Ph.D.

University of Nebraska, 2018

Advisors: Julie A. Peterson and Lance J. Meinke

Diabrotica virgifera virgifera LeConte (Coleoptera: Chrysomelidae), the western corn rootworm (WCR), is a major pest of corn (Zea mays L.) in the United States and

Europe. WCR management options comprise mainly transgenic hybrids, insecticide applications and crop rotation. WCR is highly adaptable to management practices and field-evolved resistance to transgenic corn, insecticides and crop rotation in the United

States Corn Belt has been reported. Therefore, the motivation for this project was to look into alternative options for WCR management. The goal of this dissertation is to characterize the natural enemies from irrigated commercial cornfields in Nebraska and examine their potential as biological control agents of the WCR.

We surveyed five cornfields to document populations of arthropod predators, entomopathogenic fungi (EPF) and entomopathogenic nematodes (EPN). Yellow sticky cards and dry pitfalls captured a diverse community of above-ground natural enemies but their impact on WCR population dynamics is unlikely. In the laboratory, we isolated

EPF and EPN species from soil samples using a baiting technique with Galleria mellonella

L. and Tenebrio molitor L. Entomogenous fungi with a variety of ecological roles were

detected in every cornfield. Entomopathogenic fungi made up the majority of isolates, primarily represented by Metarhizium, but other genera of known and potential EPF include Beauveria, Penicillium, Pseudogymnoascus, and Purpureocillium. In the laboratory, forty-eight strains were screened against WCR larvae. Results showed that

Metarhizium anisopliae, M. robertsii, Pseudogymnoascus sp. and BotaniGard (Beauveria bassiana) caused mortality higher than the control and should be explored further in field studies. Six strains that were tested against the WCR can also infect prepupae of western bean cutworm (Striacosta albicosta Smith), another damaging pest of corn in

Nebraska.

We also determined that EPN strains of Heterorhabditis bacteriophora Poinar and Steinernema spp. are present in Western Nebraska cornfields. An inoculation project with commercial and New York strains of EPN did not cause significant mortality in WCR populations, potentially due to native Steinernema spp. being present in the control plots. Describing the natural enemy community from WCR-infested fields is a necessary first step in the exploration of biological control as a management tool against this devastating pest. i

Acknowledgements

To all the people that helped me through this journey, I would like to thank you. I am very lucky for the wonderful committee members that served as great mentors of my studies. First, I would like to thank my advisor Dr. Julie Peterson for taking me on as her first graduate student and for always believing in my potential. I would also like to thank Dr. Lance Meinke for serving as my co-advisor and providing consistent valuable advice in my studies. To Dr. Thomas Powers, I will forever treasure the moments sitting by the microscope and learning about nematodes from him. Dr. Gary Yuen thank you for all the mycology advice and for lending lab space for my studies. Dr. Robert Wright thank you for sharing all your knowledge with me all along.

To the current and past members from the Agroecosystems Laboratory: thank you so much for all the support and camaraderie throughout these years. Kayla Mollet and Débora Montezano; field and lab work were always fun with you around! Dotti

Dobson, Jonas Macedo, Vinícius Victor, Westen Archibald, Gavin Owen, Priscila da Luz,

Dr. Katharine Swoboda Bhattarai, Samantha Daniel, Alex Lehmann and Ethan Hoffart; I am very grateful for all your help throughout my doctorate.

I would like to thank Dr. Anthony Adesemoye, who served as a key mentor and collaborator of all my mycological work. Thank you for all the teachings and assistance with the fungal studies and for opening up your lab to me. Thanks to members of the

Adesemoye lab: Hsin-Ho Wei, Michael Eskelson, Dr. Lipi Parikh, Srikanth Kodati, and

Jessica Joedeman for assistance and friendship in the laboratory. I also want to thank ii

Bob Skates, Jeff Golus and all other members from the West Central Research and

Extension Center in North Platte for making it a great working environment.

To the Plant Pathology Department that was my second home in both my

Master’s and Ph.D., thank you for welcoming me in your laboratories. Thank you to the

Powers Laboratory for letting me learn from you. Thanks to Kris Powers, Tim Harris,

Rebecca Higgins and Lisa Sutton for the immense help and training with my nematode studies. To the Yuen lab, thank you for making me a member of your lab. Christy

Jochum, I am so thankful for the mycology training you gave me and your caring friendship in and out of the lab. Thank you to Joe Schenk and Tyler Juhlin for the technical assistance in the lab and to Jessica Walnut, Vivian Shi, Anthony Muhle, Rufus

Akinrinlola and Karen Silva for being great lab mates. I have very fond memories from these two lab groups and will carry their friendship with me.

I also want to acknowledge the Meinke Lab and their pivotal role in my studies.

Thank you to Emily Reinders, Dariane Souza, Jim Brown and Jordan Reinders for their assistance and advice on rearing rootworms and conducting experiments.

Thank you also to all past and current students, staff and faculty from the

Entomology Department. I am very proud to have my M.S. and Ph.D. from this department. I have learned so much from all of you and made many life-long friends.

Thank you to Dr. Gary Brewer and all professors for creating a culture that encourages life-work balance. I am very thankful for all the support everyone gave me during my pregnancy, maternity leave and motherhood journeys. iii

Thanks to all my off-campus collaborators that made my work possible. Dr.

Stefan Jaronski for sharing his expertise on insect pathology with me. I am also grateful to Dr. Shapiro–Ilan and Stacy Byrd for providing nematodes for my molecular studies.

And thank you to Dr. Elson Shields and Anthony Testa for giving me training on entomopathogenic nematodes and providing nematodes for my field studies.

My sincere gratitude also goes to all the farmers; David Schmitt, Donnie Hajek,

Alex Canning, Jason and Jarett Barnhill that let us conduct our experiments in their fields. And thanks to Larry Appel (in memoriam) for helping finding field sites. The UNL faculty and Staff at Brule and Grant, thank you for assisting in my project. Thank you to

Robert Klein, Toby Spiehs, Strahinja Stepanovic, and Jeremy Milander.

To my family, you are everything to me. To my wonderful husband Greg, I am so thankful I have you as my partner. You are the best husband and the best father to our daughter I could have asked for. Thank you for all the support and understanding throughout this long journey. Thank you for taking care of Emily on the nights and weekends when I had to work. Thanks to my sweet baby girl Emily for joining me in the final years of my studies and bringing so much joy to my life. I cherish the memories of her kicking away in my belly while I was in the lab, answering my comprehensive exams or analyzing data. And now, seeing her contagious smile everyday makes me the happiest person in the world. To my parents, Jose Claudio and Rosa Maria, and my sisters Luciane and Luana, you are my rock and I love you all so much. Thank you for the continuous support throughout my education. This degree is as much yours as it is mine. iv

This project was partially supported by the Nebraska Agricultural Experiment

Station with funding from the State of Nebraska and the Hatch Act (Accession Number

1007272) through the USDA National Institute of Food and Agriculture. Funding for

Chapter 4 and Appendix 2 was provided by the Nebraska Corn Board (Project ID 42057).

v

TABLE OF CONTENTS ACKNOWLEDGEMENTS ...... I TABLE OF CONTENTS ...... V LIST OF TABLES ...... VII LIST OF FIGURES ...... VIII CHAPTER 1: LITERATURE REVIEW ...... 1

WESTERN CORN ROOTWORM ...... 1 Biology and ecology ...... 1 Larval mortality factors ...... 3 CURRENT WCR MANAGEMENT PRACTICES ...... 6 Rotation ...... 7 Chemical control ...... 8 Host plant resistance ...... 10 BIOLOGICAL CONTROL OF WESTERN CORN ROOTWORM ...... 12 Arthropod natural enemies: Predators ...... 14 Arthropod natural enemies: Parasitoids ...... 16 Entomopathogenic fungi ...... 16 Entomopathogenic bacteria ...... 18 Entomopathogenic nematodes ...... 18 COMPATIBILITY OF BIOLOGICAL CONTROL WITH CURRENT WCR MANAGEMENT OPTIONS ...... 22 JUSTIFICATION ...... 27 RESEARCH OBJECTIVES ...... 27 REFERENCES ...... 29 CHAPTER 2: SURVEY FOR POTENTIAL ABOVE-GROUND PREDATORS OF THE WESTERN CORN ROOTWORM WITH EMPHASIS ON GROUND BEETLES (COLEOPTERA: CARABIDAE)...... 41 INTRODUCTION ...... 41 MATERIALS AND METHODS...... 44 RESULTS ...... 46 DISCUSSION ...... 48 REFERENCES ...... 54 TABLES ...... 59 FIGURES ...... 61 CHAPTER 3: CHARACTERIZATION OF POTENTIAL FUNGAL ENTOMOPATHOGENS FROM COMMERCIAL, IRRIGATED, CONTINUOUS CORNFIELDS...... 64

INTRODUCTION ...... 64 MATERIALS AND METHODS...... 66 RESULTS ...... 70 DISCUSSION ...... 73 REFERENCES ...... 82 TABLES ...... 92 FIGURES ...... 101 CHAPTER 4: SCREENING OF INSECT-ASSOCIATED FUNGI FROM NEBRASKA AGAINST WESTERN CORN ROOTWORM, DIABROTICA VIRGIFERA VIRGIFERA LECONTE...... 105

INTRODUCTION ...... 105 MATERIALS AND METHODS...... 107 RESULTS ...... 111 DISCUSSION ...... 112 vi

REFERENCES: ...... 118 TABLE ...... 124 CHAPTER 5: IDENTITY AND SEASONAL PATTERNS OF NEMATODES ASSOCIATED WITH INSECT CADAVERS FROM CORNFIELDS IN NEBRASKA...... 127 INTRODUCTION ...... 127 MATERIALS AND METHODS...... 130 RESULTS ...... 137 DISCUSSION ...... 139 REFERENCES ...... 149 TABLES ...... 157 FIGURES ...... 165 CHAPTER 6: SUMMARY AND CONCLUSIONS ...... 171

REFERENCES: ...... 179 APPENDIX 1. FIELD SITES LOCATION AND AGRONOMIC CHARACTERISTICS...... 184 APPENDIX 2: PRELIMINARY TEST OF ENTOMOPATHOGENIC FUNGI AGAINST THE WESTERN BEAN CUTWORM, STRIACOSTA ALBICOSTA SMITH...... 186

INTRODUCTION ...... 186 MATERIALS AND METHODS...... 186 RESULTS ...... 188 DISCUSSION ...... 189 REFERENCES: ...... 190 APPENDIX 3: CURRICULUM VITAE CAMILA OLIVEIRA HOFMAN ...... 192 vii

LIST OF TABLES

Table 2.1. Total abundance of predator taxa found on yellow sticky cards per field...... 59

Table 2.2. Carabidae species found in dry pitfalls per field...... 60

Table 3.1 Fungal isolates from irrigated cornfields identified by the internal transcriber spacer (ITS) and beta-tubulin (BT) regions...... 93

Table 3.2 Ecological functions associated with baited fungi...... 99

Table 4.1 Mean mortality of western corn rootworm larvae in soil and immersion- exposure assays with entomopathogenic fungi collected from cornfields in Western Nebraska...... 124

Table 5.1. General linear mixed model analysis of nematode detection in the survey project...... 157

Table 5.2. General linear mixed model analysis of nematode detection in the inoculation project...... 158

Table 5.3. Differences of treatment by sampling date least square means with Tukey adjustment for multiple comparisons. All values were considered non-significant (NS) if adjusted P > 0.05...... 159

Table 5.4. Differences of depth by sampling date least square means with Tukey adjustment for multiple comparisons. All values were considered non-significant (NS) if adjusted P > 0.05...... 160

viii

LIST OF FIGURES

Figure 2.1. Mean cumulative western corn rootworm (WCR) adults per emergence cage...... 61

Figure 2.2. Mean cumulative WCR adults per yellow sticky card...... 62

Figure 2.3. Most common Carabidae taxa in dry pitfalls...... 63

Fig. 3.1. Locations sampled in Keith and Perkins Counties in Nebraska in 2014 and 2015...... 102

Fig.3.2. Mean percent of cadavers infected with entomogenous fungi per arena in 2014...... 103

Fig.3.3. Mean cadaver infection with entomogenous fungi per arena in 2015...... 104

Fig 5.1. Diagram of EPN inoculation project...... 165

Figure. 5.2. Mean nematode detection per arena in survey sites in 2014 (a) and 2015 (b)...... 166

Figure 5.3. Mean nematode detection per treatment over sampling dates...... 167

Figure 5.4 Mean nematode detection (averaged across treatments) by sampling depth over collection periods...... 168

Figure 5.5. Cumulative WCR emergence distribution among treatments...... 169

Figure 5.6. Neighbor-joining tree of COI nucleotide sequence from 70 nematode specimens...... 170

Figure A1. Western bean cutworm mortality from entomopathogenic fungi treatments...... 191 1

CHAPTER 1: LITERATURE REVIEW

Western corn rootworm

The chrysomelid beetle Diabrotica virgifera virgifera LeConte, the western corn rootworm (WCR) is widely distributed east of the Rocky Mountains in the United States

(Meinke et al. 2009). The WCR is highly invasive and has become one of the most damaging pests of field corn in North America and Europe (Gray et al. 2009). The western corn rootworm is also Nebraska’s most prevalent rootworm species.

Corn rootworm, as the name implies, cause injury to corn (Zea mays L.) via root feeding by the pest’s larval stage. Root feeding and pruning cause a reduction in nutrient uptake and can lead to lodging of the plant (Kahler et al. 1985). Rootworm damage can negatively impact plant-water relations, decrease photosynthetic rate, and above ground biomass (Riedell 1990; Godfrey et al. 1993; Urías-López et al. 2000).

Reduced nutrients to the plant impact corn yield and lodged plants are difficult to harvest. Lodged plants may exhibit what is called “goosenecking” in which plants grow curved instead of straight up, making it hard to harvest grain (Purdue University, 2009).

Biology and ecology The life cycle of the WCR is tightly connected to corn phenology since corn is its primary host plant. A number of grasses can serve as alternate hosts for larvae, but only a few are comparable to corn as hosts e.g.: western wheatgrass, Pascopyrum smithii

(Rydb.); pubescent wheatgrass, Elytrigia intermedia (Host); side-oats grama, Bouteloua curtipendula Michx; quackgrass, Elytrigia repens L.; Rhodes grass, Chloris gayana Kunth; and fall panicum, Panicum dichotomiflorum Michx (Oyediran et al. 2004; Wilson and 2

Hibbard 2004). Pubescent wheatgrass produced an adult population size comparable to that obtained from corn (Oyediran et al. 2004). A number of other grasses also allow development to adulthood, but the emerging beetles are usually not comparable to beetles from corn in terms of size and weight (Clark and Hibbard 2004, Oyediran et al.

2004, Wilson and Hibbard 2004, Chege et al. 2005). The importance of these alternative hosts on WCR pest status in the field is uncertain (Clark and Hibbard 2004; Oyediran et al. 2004; Chege et al. 2005).

Corn seedlings exude carbon dioxide (CO2) and other volatiles that attract neonate larvae to start feeding (Hibbard and Bjostad 1988). Younger larvae feed on root hairs and tips, burrowing into the root and crown to continue feeding as they grow.

Initial feeding injury can lead to brown root tips, which is usually followed by tunneling marks and root pruning (Purdue University, 2009). Larvae go through three instars and each instar takes about one week to develop. Third instar WCRs create an earthen pupal cell in which they develop into adults. In the Corn Belt, pupation often occurs during the mid-June to late August period with a duration of around five to ten days at a temperature range of 18-30°C (Fisher 1986).

Adult emergence takes place during late June through September. Adults feed on silk and pollen, and on pollen of weeds and other crops after corn has matured

(Maredia and Landis 1993; Campbell and Meinke 2006). Males initiate emergence about

2-10 days before females, a phenomenon known as protandry (Mabry and Spencer

2003). Females are generally ready to mate when they emerge, and can mate multiple times but generally only mate once throughout their lifespan (Hill 1975; Branson 1987; 3

Hammack 1995; Moeser and Hibbard 2005; Marquardt and Krupke 2009). Studies show a wide range of oviposition averages from around 200-1000 eggs/female throughout their lifespan (Spencer et al. 2009). Females don’t oviposit eggs on the corn plants, but place them in the soil adjacent to the plant (Moeser and Hibbard 2005). The majority of eggs are oviposited in the top 10 cm of soil, but eggs can be found up to 30 cm deep

(Weiss et al. 1983; Gray and Tollefson 1988; Gray et al. 1992). Females tend to move deeper via soil cracks to oviposit in areas with adequate moisture (Gustin 1979; Kirk

1979; Weiss et al. 1983). In the Corn Belt, oviposition usually occurs from mid-July to early fall.

Eggs go through an extensive, obligate diapause through the winter and then enter into a quiescent stage until the temperature warms up to 11-12.7°C (Krysan 1978;

Gustin 1981; Krysan et al. 1984; Levine et al. 1992). After reaching this temperature threshold, post-diapause development begins and neonate larvae hatch after a few weeks depending on temperature conditions (Wilde 1971; Wilde et al. 1972; Gustin

1981; Krysan et al. 1984; Levine et al. 1992; Godfrey et al. 1995). In the U.S and Europe, initial egg hatch occurs approximately between late May to the beginning of June

(Meinke et al. 2009).

Larval mortality factors Larvae are the most damaging stage of this pest and for this reason larval mortality factors must be well understood in order to establish effective pest management programs. Besides pest management practices, larvae are subject to abiotic and biotic mortality factors in the soil. Primary abiotic factors include soil 4 conditions such as moisture, temperature and texture. These factors are interactive and different combinations of these conditions can have different impacts on WCR larval survival. For instance, larvae exposed to high temperatures had increased survival when they were simultaneously exposed to wet conditions, whereas desiccation rapidly occurred when larvae were exposed to high temperatures and low soil moisture

(MacDonald and Ellis 1990). Besides direct mortality, abiotic factors can also affect larval movement in the soil, which in turn can disrupt larval establishment on the host, leading to mortality (MacDonald and Ellis 1990). Larvae show reduced movement when soils are dry (12% moisture) or saturated (38% moisture); movement was greater at moderate conditions (24-30% moisture) (MacDonald and Ellis 1990).

WCR tend to prefer soils with higher clay content (Turpin and Peters 1971).

Larvae exposed to soil with high clay content (48.2%) had higher adult emergence than larvae from sandy soils that have lower clay content (4.8-11.5%) (Turpin and Peters

1971). Sand particles are thought to physically injure WCR larvae through abrasion, which increases desiccation of larvae (Turpin and Peters 1971), but this hypothesis has never been proven. Larvae in silt-clay and loam soils moved greater than three times the mean distance (>18 cm) of larvae in loamy-sand soil (6.1 cm mean) (MacDonald and Ellis

1990). Clay soils hold water better than sandy soils; this may be a reason why WCR larvae have a tendency to prefer clay soils.

When exposed to severe hypoxia during underwater submersion experiments,

WCR larvae took longer to reach mean time mortality (LT50) at lower temperatures (10 and 15°C) than at higher temperatures (20 and 25°C) (Hoback et al. 2002), meaning that, 5 larvae that experience flooding will die quicker in higher temperatures. But not all larval stages tolerate flooding equally. Third instars had approximately half (~26 hrs) the LT50 at 25°C in hypoxia than second instars (~56 hrs) exposed to the same conditions (Hoback et al. 2002). This suggests that if flooding occurs when larvae are older (3rd vs. 2nd instars) they will have a decreased chance of survival.

Biotic factors that contribute to WCR larval mortality include density-dependent factors such as resource competition, as well as density-independent factors such as presence of suitable hosts and natural enemies. The role of mortality by natural enemies is discussed in “Biological control of western corn rootworm” section.

Larval density-dependent factors can influence mortality in WCR populations.

High larval density is directly correlated with decreased survival to adulthood (Onstad et al. 2006). High WCR infestation rates (1200 and 2400 eggs/30.5 cm of row) resulted in lower percentage survival to adulthood, plus, adults were smaller and exhibited lower egg viability when compared to lower infestation rates (300 eggs and 600 eggs/30.5 cm of row) (Branson and Sutter 1985). In a regression analysis of six infestation levels (25,

50, 100, 300, 600, 1200 and 2400 eggs/30.5 cm of row) it was determined that density- dependent mortality began at approximately 851 eggs/30.5 cm of row (Hibbard et al.

2010). In the same study, a regression analysis with previously published data showed that density-dependent mortality began at approximately 800 eggs/30.5 cm of row

(Hibbard et al. 2010). WCR in this study were also subject to unknown density- independent mortality factors, which caused at least 91% mortality, even at lower densities (Hibbard et al. 2010). In low egg densities (25 eggs/plant), larvae develop 6 faster than at higher densities (75 eggs/plant) (Weiss et al. 1985). At these higher densities, the sex ratio of emerged adults was biased more towards males, probably due to the fact that males inherently develop faster than females and are able to complete development before resources become limiting (Weiss et al. 1985). Females emerging from the high density treatment weighed significantly less than females in the low density treatment (Weiss et al. 1985). Smaller females with head capsule width of 1.15 mm lay fewer eggs (mean 506.6 eggs/female), than bigger females with head capsule widths of 1.17-1.22 mm (mean 735.6-845.5 eggs/female) (Branson and Sutter 1985).

Presence of a suitable host is essential for larval survival. Neonates are thought to have a maximum of 84 h to establish on a host before dying (Strnad and Bergman

1987a; Branson 1989), but this doesn’t mean that larvae can’t die before this time period. Strnad and Bergman (1987) found that larval survival was the same after 12, 24 and 36 h of starvation. However, after 12 h of starvation, only ~75% of larvae were able to burrow into roots, significantly lower than 100% of larvae in the non-starved treatment (Strnad and Bergman 1987). This indicates that, although they can survive starvation for relatively long periods, WCR lose their capability to establish successfully on the host, which contributes to decreased rates of survival to adulthood (Branson

1989).

Current WCR management practices

Management strategies for WCR are mostly focused on controlling the larval stage, although adult control can be employed as well to reduce egg laying and egg 7 density in next season’s corn crop. Current management strategies include crop rotation with non-hosts, insecticides and transgenic hybrids.

Rotation Crop rotation is the most effective way to minimize the WCR population in a field. This cultural management strategy is based on two principles of the WCR life cycle described above: larvae only feed on corn and adults have a strong oviposition fidelity to cornfields (Mabry and Spencer 2003). The switch from corn to a non-host breaks the

WCR life cycle and reduces insect pressure in the field. WCR larvae that hatch in a non- host field are unable to feed and will likely die from starvation. Similarly, WCR adult females from neighboring fields will be less likely to oviposit in a non-host crop. If volunteer corn is present in the rotated field some WCR can survive, but populations will likely be low in corn the following year.

Corn rotation can also reduce costs associated with WCR management because first-year corn can often be planted without the use of soil insecticides (Roth, 1996) and without the use of WCR-Bt hybrids. Rotation also provides benefits beyond WCR control. Growers that adopt a rotation schedule instead of continuous corn may see an increase in corn yield (Roth, 1996). Additionally, corn followed by another crop requires less fertilizer and less tillage than continuous corn (Roth, 1996).

A typical rotation in the Midwest is the annual corn-soybean rotation. This rotation works well for corn rootworm management for the majority of growers, but there are some problem regions in East Illinois and West Indiana where WCR have developed resistance to rotation (Levine and Oloumi-Sadeghi 1996; Levine et al. 2002; 8

Gray et al. 2009). In Nebraska, no cases of this phenotype have been reported to date.

These rotation-resistant (RR) WCR adults exhibit less affinity to stay in corn, even when silk and pollen is still available (Levine et al. 2002, Pierce and Gray 2006). Movement out of corn to soybeans by RR-WCR peaks after corn reaches R2 (reproductive stage 2, or blister stage), hence cornfields are still very susceptible to WCR injury if corn is planted in the following year (Pierce and Gray 2006, 2007). The WCR adults that move away from corn can feed on soybean foliage and oviposit in soybean fields (Levine et al.

2002). Soybeans are a sub-optimal source of food when compared to corn and it seems that WCR females under stress may oviposit prematurely in soybean fields due to this physiological stress (Mabry and Spencer 2003). It was also found that RR-WCR carry a different gut microbiota that facilitates its tolerance of antiherbivore defenses produced by soybeans (Chu et al. 2013). This lowered fidelity to oviposit in corn can lead to root injury and yield losses in corn following soybeans. Management of RR-WCR includes using transgenic corn in a 2-year rotation, increasing the rotation schedule to a 3-year rotation and/or sampling for WCR in the soybean year (O’Neal et al. 2001, Onstad et al.

2003). The use of unbaited yellow sticky cards to quantify WCR populations in soybean can indicate whether or not rootworm protection will be needed in first-year corn

(O’Neal et al. 2001).

Chemical control Chemical control of the WCR is targeted against the larval or adult stages. To prevent or slow down larval feeding, a grower can opt for high-rate insecticidal seed treatments, planting-time insecticides, or post-planting insecticides. Seed treatments 9 used are neonicotinoids such as clothianidin and thiamethoxam (Eisley and Hammond

2008). Planting-time insecticides include: organophosphates (chlorethoxyphos, chlorpyrifos, and tebupirimfos) and pyrethroids (bifenthrin, cyfluthrin and tefluthrin)

(Eisley and Hammond 2008, Bledsoe and Obermeyer 2010, Krupke et al. 2014). Planting- time and post-emergence insecticides work best when they are applied when larvae are already present or right before larvae are present (Bledsoe and Obermeyer 2010).

Adult control of the WCR can be applied to stop adult feeding, but it is primarily used to reduce egg laying and hence reduce the following year’s WCR population. If silk feeding is intensive during pollen shed, silk clipping can interfere with pollination

(Maredia and Landis 1993). A variety of organophosphates and pyrethroids, and one oxadiazine (indoxacarb) can be used to control adult populations of the WCR (DeVries and Wright 2016, Krupke et al. 2014). To prevent pollination interference by the WCR, insecticide application should take place before 50% of pollination has occurred or when pollen is being shed and silks are clipped to ½ inch or less (Krupke et al. 2014).

Numerous cases of WCR insecticide resistance have been reported in the literature. In Nebraska, WCR adults collected from several counties throughout the state showed different levels of susceptibility to methyl parathion (organophosphate) and carbaryl (carbamate) (Meinke et al. 1998). The populations with the highest levels of resistance to these insecticides came from Phelps and York counties in South-Central NE where organophosphates and carbamates were consistently used for both adult and larval control (Meinke et al. 1998). Overuse of these insecticides led to failure of the adult management program (Meinke et al. 1998). Also in Nebraska, field populations 10 were shown to have aldrin (cyclodiene organochlorine) and bifenthrin (pyrethroid) resistance (Ball and Weekman 1963, Parimi et al. 2006, Pereira et al. 2015). Although resistance to insecticides, especially overused products, has been reported in some regions of the country, the same active ingredients may still be effective in other regions.

Host plant resistance Host plant resistance (HPR) to insects has three main categories: antibiosis, antixenosis and tolerance. Plants exhibiting antibiosis have detrimental effects on the insect’s biology and survival (Painter 1951). Antixenotic plants exhibit adverse effects on the insect’s behavior in a way that the insect exhibits non-preference for the plant

(Kogan and Ortman 1978). Lastly, tolerance is a category of HPR that unlike antixenosis and antibiosis does not affect the insect’s biology or behavior. Rather, tolerant plants can withstand and/or recover from insect damage when compared to susceptible plants

(Painter 1951). Hence, tolerance is a category that is defined from the plants’ perspective not the insects.

Prior to transgenics, or genetically engineered plants with gene(s) from other species, HPR to rootworm species was done via selection of naturally occurring antibiotic, antixenotic and tolerant traits. Corn plants are fairly susceptible to WCR feeding and since there are not many innate antibiotic and antixenotic characteristics available; WCR-HPR was primarily achieved via tolerance (Wilson and Peters 1973,

Riedell and Evenson 1993, Gray and Steffey 1998, Urias and Meinke 2001). Corn hybrids that have rapid root growth and large root systems may be preferred since they are 11 more tolerant to WCR damage (Ortman et al. 1974; Branson et al. 1982). However, hybrids with over-compensatory root regrowth can exhibit less vegetative biomass including kernel weight and therefore negatively impact yield (Godfrey et al. 1993;

Urías-López and Meinke 2001).

Host plant resistance for WCR today is largely achieved via transgenic Bacillus thuringiensis Berliner (Bt) corn hybrids. The B. thuringiensis bacterium produces delta- endotoxins, crystalline proteins that when ingested by certain, susceptible insects cause septicemia and death. Delta-endotoxin genes are transformed into corn plants that can in turn produce crystalline proteins in the plant’s tissues. Sub-lethal effects due to Bt feeding also occur, and WCR larvae show increased developmental time when they feed on Bt roots (Meissle et al. 2009, Petzold-Maxwell et al. 2013). Crystalline proteins are categorized as Cry proteins and are known for their specificity to target organisms. The available registered Cry toxins against the WCR are: Cry3Bb1, mCry3A, Cry34/35Ab1, and eCry3.1Ab (DiFonzo 2017).

In 2003, Cry3Bb1 was the first plant-incorporated rootworm-active protein registered by the EPA in the U.S. (tradename: YieldGard® Rootworm, USEPA 2010). Since then, there has been a shift over time to incorporate more than one WCR toxin per plant, also known as trait pyramiding. The concept is that if WCR develops resistance to one toxin, but is still susceptible to the other, death will still occur, and this will delay resistance evolution.

Bt corn hybrids require a refuge, or in other words, non-rootworm Bt corn areas where susceptible individuals can survive. Refuge systems proposed for Bt hybrids is the 12 high-dose refuge, with high-dose being defined as 25x the toxin concentration to kill

99.99% of susceptible insects (USEPA 2001). However, none of the current WCR -Bt hybrids are registered as high-dose, which allowed for some insect survival on those hybrids (Andow et al. 2016). The moderate-dose approach together with the wide use of WCR- Bt hybrids has led to the development of field-evolved resistance. The first resistance case to rootworm-protected Bt corn was reported in Iowa in 2011 for

Cry3Bb1 hybrids (Gassmann et al. 2011). There was a strong relationship between the number of years Cry3Bb1 hybrids had been grown and resistance, or in other words, continuous Bt toxin pressure lead to increased WCR survival in problem fields. Cross- resistance between Cry3Bb1 and mCry3A or eCry3.1Ab has been reported but no cross- resistance between these proteins and Cry34/35 (Gassmann et al. 2014, Wangila et al.

2015, Zukoff et al. 2016). Field-evolved resistance to Cry3Bb1 or mCry3A has subsequently been reported from other Corn Belt states (Schrader et al. 2016, Wangila et al. 2015, Zukoff et al. 2016) as well as initial documentation of field-evolved WCR resistance to Cry34/35 (Gassmann et al. 2016, Ludwick et al. 2017).

Biological control of western corn rootworm

Biological control utilizes living natural enemies to manage pest populations.

Three types of biological control exist: classical, augmentation and conservation biological control. According to (Eilenberg et al. 2001) classical biological control is defined as: “The intentional introduction of an exotic, usually co-evolved, biological control agent for permanent establishment and long-term pest control.” Classical 13 biological control aims to restore the natural balance present in the pest’s native habitat that prevents pest outbreaks (Hajek, 2004).

Augmentation biological control involves both methods of inoculative and inundative control and is different from classical biological control in that it does not aim for permanent establishment of the biological control agent (Hajek, 2004). Inundative control is often referred to as a “biopesticide” since it is expected to cause high mortality in a short period (Hajek, 2004). These biopesticides are applied in high doses and only the released agents, not subsequent generations, cause mortality. Inundative biological control can be defined as: “The use of living organisms to control pests when control is achieved exclusively by the released organisms themselves’’ (Eilenberg et al.

2001). Inoculative biological control is different from inundative control in which the progeny from the released biological control organism can also inflict pest mortality

(Hajek, 2004). Inoculative biological control is defined as: “The intentional release of a living organism as a biological control agent with the expectation that it will multiply and control the pest for an extended period, but not permanently’’ (Eilenberg et al.

2001).

The last category is conservation biological control which is defined by Eilenberg et al. (2001) as: “Modification of the environment or existing practices to protect and enhance specific natural enemies or other organisms to reduce the effect of pests”.

Conservation biological control aims to enhance natural populations instead of introducing biological control agents. Here, knowledge of the factors that are preventing natural enemies from being effective biological control agents is necessary in order to 14 establish the program (Hajek, 2004). With this understanding, practicioners can reduce disturbances such as pesticides and lack of hosts/food by manupulating the habitat to benefit natural enemies (Landis et al. 2000).

Biological control of corn rootworm species is an underexplored area when compared to other available management practices. Several natural enemies are found to prey on or infect WCR (Kuhlmann and van der Burgt 1998, Toepfer et al. 2009); however, the extent of their impact on WCR population reduction is often unknown.

Biological control of WCR has been investigated in the U.S. and in Europe, but the adoption of biological control on commercial farms has been limited. There are five main groups of natural enemies of the WCR: arthropod natural enemy predators and parasitoids; and entomopathogenic fungi, bacteria, and nematodes. Biological control doesn’t have to be a stand-alone strategy to pest management. Its compatibility with other pest management methods is discussed in section 2.4.

Arthropod natural enemies: Predators A wide variety of arthropods have been observed feeding on WCR eggs and larvae. Main taxa include predatory mites (Mesostigmata), ground beetles (Carabidae), hister beetles (Histeridae), rove beetles (Staphylinidae), carpet beetles (Dermestidae), centipedes (Chilopoda), and spiders (Araneae) (Kuhlmann and van der Burgt 1998). The ant species Lasius neoniger Emery has also been linked to consuming WCR larvae (Kirk

1981).

Lundgren et al. (2009a) used polymerase chain reaction (PCR) to amplify WCR

DNA extracted from predatory taxa in plots artificially infested with WCR eggs. Out of 15

1550 specimens, 166 (~10%) were positive for WCR DNA in their guts. Most common specimens positive for WCR DNA included Acari (Chausseria sp. and other mites);

Lycosidae and other Aranae; Phalangium opilio L. (Opiliones); Scarites quadriceps

Chaudoir, Poecilus chalcites (Say) and other Carabidae species; and Staphylinidae species (Lundgren et al. 2009b). A significantly higher percentage of predators were found positive for WCR DNA when this pest was in the egg stage; however, frequency of consumption and consumption index were comparable between egg and larval stages

(Lundgren et al. 2009b). Eggs are probably consumed more because they are in the field for a longer period than larvae (Lundgren et al. 2009b). Predators with chewing mouthparts had a lower consumption index than predators with sucking mouthparts

(Lundgren et al. 2009b). This is probably due to the fact that WCR larvae exhibit a hemolymph defense that coagulates quickly in the predators’ mouthparts (Lundgren et al. 2009a). This sticky hemolymph triggers immediate cessation of feeding and intense cleaning of mouthparts (Lundgren 2009a). Predators in the study (Poecilus cupreus L. and Harpalus pensylvanicus De Geer (Carabidae) spent significantly more time cleaning their mouthparts than eating, and once exposed to the hemolymph defense of WCR, they were hesitant to feed again on WCR (Lundgren et al. 2009a). A diverse community of predators is shown to increase the frequency of WCR egg and larval predation

(Lundgren and Fergen 2014). This indicates that in a biological control program against the WCR, multiple predators may enhance mortality.

16

Arthropod natural enemies: Parasitoids Insect parasitoids are obligate parasites that develop inside (endoparasitoids) or outside (ectoparasitoids) an individual host and result in the host’s mortality.

Hymenoptera and Diptera (Insecta) are the two major orders with parasitoid members.

Parasitoids from the genus Celatoria spp. (Diptera: Tachinidae) lay eggs containing a fully developed first instar larva on Diabrotica spp. adults (Kuhlmann and van der Burgt

1998). Celatoria spp. have been studied to control rootworm populations, especially the species Celatoria compressa Wulp (Kuhlmann et al. 2005). There are several constraints with the rearing of Celatoria spp., including its low fecundity rates, specific temperature requirements, and the univoltine nature of the host (WCR), all of which makes it an impractical biological control agent for augmentation (Toepfer et al. 2005). Moreover, studies are needed to evaluate its potential for conservation biological control (Toepfer et al. 2008).

Entomopathogenic fungi Entomopathogenic fungi (EPF) are present in ecosystems worldwide and play a role in keeping insect pests regulated. Infective spores can be present above and below ground and as opposed to other entomopathogens, they do not need to be eaten in order to be pathogenic (Hajek, 2004). EPF spores attach to the host’s cuticle, and then they germinate and penetrate the insect host (Hajek, 2004; Wraight et al. 2007). Fungi use both the force of mechanical pressure and digestive enzymes to pierce/digest the insect’s cuticle (Hajek, 2004). The fungus eventually takes over the host’s hemolymph and hemocoel and causes death. 17

The fungi Beauveria bassiana (Bals. -Criv.) Vuill. and Metarhizium anisopliae

(Metschn.) Sorokin infect WCR larvae and pupae (Kuhlmann and van der Burgt 1998).

These species are naturally occurring worldwide and can be used as part of a biological control program. Rudeen et al. (2013) found that M. anisopliae and B. bassiana were common in commercial cornfields in Iowa and that the WCR mortality caused by field- collected strains was comparable, if not greater, than commercially available strains.

Fungal strains can differ in their virulence to specific stages of WCR. A screen of twenty strains of M. anisopliae, B. bassiana and Beauveria brongniartii (Sacc.) Petch revealed varying levels of mortality (Pilz et al. 2007). In general, most M. anisopliae strains were more virulent than B. bassiana strains, but no strain induced greater than

50% larval mortality (Pilz et al. 2007). Adults were more susceptible to these strains than larvae, which the authors hypothesize that larvae are better adapted to living in the soil with naturally co-ocurring entomopathogens than adults. The most virulent M. anisopliae larval strain caused 47% mortality, but in comparison, the most virulent M. anisopliae strain for adults achieved 97% mortality (Pilz et al. 2007).

Indigenous B. bassiana was also found to infect adult Diabrotica spp. at emergence (Bruck and Lewis 2001). This means that emerging WCR adults are exposed to B. bassiana from the soil as they emerge, which causes adult mortality. Infection rates were low at 3.2%, but the researchers estimated that with their high level of ~2.6 million WCR beetles/hectare, there is a potential reduction in oviposition of 41 million eggs/hectare (Bruck and Lewis 2001). In addition, B. bassiana can also cause non-lethal effects. Infection of B. bassiana on mated females causes a 30% or greater reduction of 18 fecundity in the surviving beetles (Mulock and Chandler 2001a). Oviposition reduction can lessen WCR pressure in the following year’s corn crop.

Entomopathogenic bacteria Entomopathogenic bacteria can enter the host’s body via wounds but the primary entryway is via ingestion (Hajek, 2004). Bacteria proliferate inside the host’s hemocoel and kill the host via septicemia. The most well-known entomopathogenic bacteria, B. thuringiensis and Bacillus sphaericus Meyer and Neide, are from the spore- forming family Bacillaceae (Hajek, 2004). Non-spore forming entomopathogens include

Serratia entomophila Grimont and Serratia marcescens Bizio (Enterobacteriaceae).

Research on bacteria pathogenic to WCR comprises mainly the study of Bt strains that contain genes that have been incorporated into transgenic plants, but other bacterial species have not been studied in depth. Bacteria symbiotic with entomopathogenic nematodes are discussed in the next section.

Entomopathogenic nematodes Entomopathogenic nematodes (EPN) are natural enemies of soil arthropods that kill hosts within one to four days post-infection (Koppenhöfer 2007). Seven nematode families are considered to be entomopathogenic, but only Heterorhabditidae and

Steinernematidae (Nematoda: Rhabditida), which are obligate endoparasites of insect hosts, are consistently researched as biological control agents (Koppenhöfer 2007). The family Heterorhabditidae contains the single genus Heterorhabditis and

Steinernematidae has two genera: Steinernema and Neosteinernema (Stock and

Goodrich-Blair 2012). The most studied EPN species for biological control of WCR are 19

Steinernema feltiae (Filipvej), Steinernema carpocapsae (Weiser) and Heterorhabditis bacteriophora Poinar.

Third stage infective juveniles (IJ’s), or dauer juveniles, are the only life stage of the nematodes that can live outside the host in order to search for new ones. IJ’s infect insect hosts by penetrating thin areas of the cuticle or via natural openings such as the mouth, spiracles, genital pores and anus. Once inside the host’s hemocoel, IJ’s release mutualistic bacteria that induce host mortality via septicemia within 48 hours (Kaya and

Gaugler 1993). Steinernematidae are associated with bacteria from the genus

Xenorhabdus and Heterorhabditidae with Photorhabdus bacteria. EPN’s develop on the host’s metabolized tissue and the proliferated bacteria for 1-3 generations until host tissues are depleted and IJ’s need to seek a new host (Koppenhöfer 2007).

EPN’s are most effective against WCR in second and third instars (Jackson and

Brooks 1989; Journey and Ostlie 2000; Kurtz et al. 2009). The Mexican strain of S. carpocapsae infects all stages of the WCR besides eggs. However, based on the mean number of S. carpocapsae/instar, they found that third instar rootworms are 5x more susceptible than second instars and 75x times more susceptible than first instars and pupae (Jackson and Brooks 1995). Differences in infectivity are most likely due to bigger orifices/entry points in later instars (Jackson and Brooks 1995). It is important to point out that rootworm larvae only support one generation of nematode reproduction due to the small amount of larval host tissue (Jackson and Brooks 1995).

Different species of EPN’s have different strategies to forage for hosts. These strategies are known as cruise and ambush strategies. “Cruisers” actively search for the 20 host and because they are mobile they are more prone to encounter sedentary hosts.

Alternatively, “ambushers” are likely to encounter moving hosts as they can attach to the passing host. Species can also adapt a foraging strategy that is intermediate between cruising and ambushing (Lewis et al. 2006). Steinernema carpocapsae is an ambusher and H. bacteriophora is a cruiser (Grewal et al. 1993, 1994). However, S. feltiae exhibits both foraging behaviors, as a small subset of the population actively cruise for hosts but the majority of the population acts as ambushers (Gaugler et al.

1989).

IJ’s use chemo-, thermo- and mechanotaxis to locate and recognize insect hosts

(Lewis et al. 2006). In the case of the rootworm-EPN system, it was also found that nematodes are attracted to injured corn roots (Rasmann et al. 2005). The sesquiterpene

(E)-β-caryophyllene is a secondary compound present in the damaged roots of certain corn varieties (Rasmann et al. 2005). A European corn variety that contained (E)-β- caryophyllene strongly attracted Heterorhabditis megidis, which promoted secondary plant defense. (E)-β-caryophyllene treatments had five times the rate of WCR infection by H. megidis than the treatment that didn’t contain (E)-β-caryophyllene (Rasmann et al.

2005). Interestingly, modern corn hybrids in the U.S., seemed to have accidently lost (E)-

β-caryophyllene during breeding programs (Rasmann et al. 2005).

Field studies have shown that EPN applications can add an extra layer of protection to corn roots. Wright et al. (1993) applied a rate of 1.2 and 2.5 x 109 IJ’s of S. carpocapsae per hectare via center-pivot irrigation. In this study, they found that at the higher rate of IJ’s, nematodes could be just as effective as chlorpyrifos in reducing WCR 21 adult emergence. Similarly, S. carpocapsae and H. bacteriophora treatments (200,000

IJ/plant) were just as effective as terbufos in suppressing beetle emergence (Jackson

1996). Heterorhabditis bacteriophora applied at a rate of 3.8 x 109 IJ/hectare was comparable to tefluthrin and clothianidin-coated seed treatments in reducing WCR emergence (Pilz et al. 2009). Chlorpyrifos, terbufos (organophosphates), clothianidin

(neonicotinoid), and tefluthrin (pyrethroid) are common insecticides utilized for larval

WCR treatment. Collectively, these studies indicate that if well timed, EPN treatments can be used as an alternative to WCR pesticides in problem fields. However, improvements in EPN production costs still have to be made to turn that into reality.

In another study, S. feltiae, H. bacteriophora and H. megidis were applied to test plots via two methods: direct spray at sowing time (2.8 x 109 IJ/hectare) or spray when

WCR larvae were expected to be at second instar (3.4 x 109 IJ/hectare) (Toepfer et al.

2008). All three EPN species showed moderate to high levels of WCR control; when they were applied at sowing time, H. bacteriophora showed 81% reduction in WCR emergence, while the S. feltiae and H. megidis only accounted for 36% and 49% reduction, respectively, in adult emergence (Toepfer et al. 2008). Moreover, when EPN’s were applied when second instars of the WCR were present, both H. bacteriophora and

H. megidis were statistically similar in reducing WCR adult emergence (75% and 69%, respectively), but S. feltiae only provided 32% emergence reduction (Toepfer et al.

2008a). Even though these species differed in suppressing WCR populations, they were equally successful in protecting roots from damage regardless of which application method was used (Toepfer et al. 2008). In this study they infested 150 WCR eggs/ plant, 22 which is regarded as a low rootworm density; however, it is possible that EPN efficacy is different under high WCR pressure. Europe’s WCR densities are often much lower than

U.S. WCR densities and therefore, EPNs may hold greater potential in Europe.

Traditional studies with EPN releases have treated EPN’s as biopesticides (Shields

2015a). Biopesticides act in a similar fashion as pesticides, in which a high-dose application of a biological control agent is applied inundatively in the field and it is expected to cause high mortality of the pest within a short time period (Hajek, 2004).

Inundative releases of EPN’s use commercial strains that have lost their ability to persist in the environment (Shields 2015). Few researchers have looked into inoculation, rather than inundation, of EPN in the environment. A recent study showed that it is possible for S. carpocapsae and S. feltiae to reduce populations of the alfalfa snout beetle

(Otiorhynchus ligustici (L.) (Coleoptera: Curculionidae) in long-term alfalfa-corn and alfalfa-soybean rotations (Shields and Testa 2015a). This research team has also shown the persistence of these EPN species in vineyards against grubs of the Japanese beetle

Popillia japonica Newman (Coleoptera: Scarabaeidae) (Shields 2015a). The goal here is to introduce EPN strains that are able to persist in the environment, throughout multiple generations, even in low density host populations and provide long-term pest control (Shields 2015a).

Compatibility of biological control with current WCR management options

The management of WCR requires a complex, multi-strategy approach. Tactics adopted by growers are often over-used and ineffective after less than a decade due to field-evolved resistance. There is a need to develop new technologies against the WCR 23 that could be efficacious in the long run. More research needs to be done in integrating control strategies so that we have an ecologically and economically sound integrated pest management strategy against this highly destructive pest. Integrating biological control into the scope of management practices against the WCR might help us suppress population growth of this pest.

Some researchers have studied tri-trophic interactions between Bt, WCR and entomopathogens. Growth chamber and laboratory studies determined that a combination of fungal (B. bassiana and Metarhizium brunneum Petch) and nematode (S. carpocapsae, S. glaseri, H. bacteriophora) entomopathogens acted in an additive way with expressing Cry34/35Ab1 to reduce WCR larval densites, and survival to adulthood

(Petzold-Maxwell et al. 2012a). Because Bt proteins caused a delay of WCR development and the pathogens increased mortality, the pathogens had a longer window of time to infect the larvae; hence they acted independently but in an additive manner (Petzold-

Maxwell et al. 2012a). This additive effect may go away in WCR populations that display complete resistance to Cry3Bb1 and do not exhibit developmental delays (Wangila and

Meinke 2017). In the field, the addition of M. brunneum, S. carpocapsae, and H. bacteriophora in plots with Cry34/35Ab1 hybrids increased yield when compared to the

Bt only treatment, despite the fact that the entomopathogens did not decrease survival of WCR (Petzold–Maxwell et al. 2013). Moreover, they found that root injury in the Bt + entomopathogens treatment was only reduced when WCR abundance was high.

Conversely, root injury in the non-Bt + entomopathogens treatment was reduced when

WCR abundance was low (Petzold-Maxwell et al. 2013). The explanation for these 24 results is that at high WCR abundance, entomopathogens alone cannot protect roots from injury without another management practice, while at low WCR abundance, entomopathogens alone can give roots a good level of protection (Petzold-Maxwell et al. 2013). The addition of entomopathogenic nematodes, such as S. carpocapsae, may not increase mortality rates of WCR larvae when used in conjunction with the

Cry34/35Ab1 Bt hybrid treatment (Rudeen and Gassmann 2013). Steinernema carpocapsae are more infective in later instars of WCR and since larvae feeding on

Cry34/35Ab1 had delayed development, the larvae were not in the growth stage most susceptible to the nematode (Rudeen and Gassmann 2013).

A combination of a Cry3Bb1 Bt hybrid and entomopathogenic fungi can also be effective against WCR. Metarhizium anisopliae successfully infected WCR larvae and adults that fed on Cry3Bb1 (Meissle et al. 2009). Both WCR larval and adult stages were equally susceptible to fungal infection regardless if they fed on Cry3Bb1 or non-Bt diet

(Meissle et al. 2009). Since there is no interaction between M. anisopliae and Cry3Bb1 on WCR mortality, this result implies an additive effect of the treatments, rather than synergistic effect (Meissle et al. 2009). Hence, fungi and Bt-hybrids can be used as complementary tactics against rootworms.

Adding a mortality factor, such as entomopathogens to complement a Bt hybrid may delay Bt resistance evolution of herbivores because of associated fitness costs

(Gassmann et al. 2008, Gassmann et al. 2009a,b; Hannon et al. 2010, Gassmann et al.

2012). For instance, resistance to Cry1Ac in the pink bollworm, Pectinophora gossypiella

(Saunders) (Lepidoptera: Gelechiidae) was delayed in treatments with Steinernema 25 riobrave (Gassmann et al. 2006, 2008). Resistance to Cry1Ac may evolve slower if populations are treated with S. riobrave since resistant individuals had higher mortality rates by S. riobrave than Cry1Ac-susceptible individuals (Gassmann et al. 2006, 2009b).

However, this is not the case with WCR Bt resistance and entomopathogens. There were no fitness costs associated with resistance to Cry3Bb1 when WCR were exposed to entomopathogens (Petzold-Maxwell et al. 2012c; Hoffmann et al. 2014). Steinernema carpocapsae and H. bacteriophora had no difference in WCR infection rates for Cry3Bb1 susceptible and resistant individuals (Petzold-Maxwell et al. 2012a). Similarly, S. feltiae,

H. bacteriophora, B. bassiana and M. brunneum did not impose any fitness cost on WCR resistant to Cry3Bb1 (Hoffmann et al. 2014). These entomopathogens increased mortality of both Bt resistant and susceptible individuals when compared to no pathogen treatments but at no resistance-related fitness cost (Hoffmann et al. 2014).

In addition to Bt-entomopathogen interactions, one must also consider the possible interactions of entomopathogens with pesticides (insecticides, fungicides and nematicides). These interactions can be negative, neutral or positive. Negative interactions occur when the pesticides inhibit entomopathogen activity such as reduced reproduction, virulence and altered behavior (Manachini 2002). Positive interactions can create enhanced mortality of the pest in comparison to either product alone (Manachini

2002).

The entomopathogenic nematodes S. feltiae, S. carpocapsae, H. bacteriophora and Heterorhabditis heliothidis for the most part can tolerate pesticides even at higher than recommended concentrations (Rovesti et al. 1988; Rovesti and Deseö 1990, 1991). 26

However, some chemicals were found to have negative effects on movement, infectivity and motility of IJ’s (Rovesti and Deseö 1990, 1991). Alternatively, the insecticide tefluthrin (pyrethroid) in conjunction with S. carpocapsae can be synergistic; together they caused an average of 24% more mortality than the expected additive mortality

(Nishimatsu and Jackson 1998). Hence in this case, EPN’s can be used in conjunction with an insecticide to cause greater mortality to WCR (Nishimatsu and Jackson 1998).

The entomopathogenic fungi (B. bassiana and M. anisopliae) are also subject to interactions with pesticides. These species seem to be compatible with neonicotinoids

(Neves et al. 2001; Batista Filho et al. 2001). Moreover, at the recommended rate some fungicides decreased B. bassiana and M. anisopliae germination, but virulence seemed to be unaffected for most interactions (Shah et al. 2009). All these studies should be taken with caution as the interactions in the field between chemicals and pathogens are certainly different than in vitro interactions. Also, entomopathogenic nematodes and fungi may be protected in their hosts, in which case they may not come in contact with the agrochemicals in the soil.

Interactions between entomopathogens and crop rotation must be investigated to establish entomopathogen persistence in the environment. Strains of the nematodes

S. feltiae and S.carpocapsae were tested to see if they persist in alfalfa-corn and alfalfa- soybean rotations and continue to be infective to the target insect pest, Otiorhynchus ligustici (L.), the alfalfa snout beetle (Shields and Testa 2015a). Data were collected for up to 6 years in some fields and both EPN species were shown to persist in continuous alfalfa as well as in alfalfa-corn and alfalfa-soybean rotations (Shields and Testa 2015). It 27 was inferred, but not tested, that these nematodes infected rootworms in corn years and hence were able to thrive. In some years, Shield and Testa (2015) saw a spike in nematode detection in the non-alfalfa crop rotations, suggesting that insects of corn and soybeans are hosts for both S. feltiae and S. carpocapsae.

Justification

Western corn rootworm mortality (WCR) is a multifactor issue that must be understood in order to establish a sound management program. Multiple interactions among abiotic and biotic factors occur that influence mortality rates in all stages of the

WCR. Nebraska is currently the third largest corn producer in the country and WCR is one of the state’s most important pests. The climate is semiarid in West Central

Nebraska and because of that, continuous corn is primarily grown under center-pivot irrigation systems. There is a strong need for new management options for the WCR as populations of this pest in West Central Nebraska and other areas of the Corn Belt have evolved resistance to many currently used management options. Biological control is an understudied area of WCR management and it is unknown what natural enemy predators and entomopathogens are present in West Central Nebraska.

Research objectives

The broad goal of this dissertation was to understand the biodiversity of natural enemies present in irrigated commercial cornfields in the context of finding alternative and complementary WCR management options. Specific chapter objectives were:

Chapter 2: Identify potential above-ground and epigeal predators of WCR with an

emphasis on determining if Carabidae beetles were consuming WCR in the field. 28

Chapter 3: Document the community of entomogenous fungi from the rhizosphere throughout the field season.

Chapter 4: Determine the impact of native fungal strains isolated from Chapter 3 on

WCR larval mortality in the laboratory.

Chapter 5: Document the diversity of entomogenous nematodes in commercial cornfields and evaluate entomopathogenic nematode applications against WCR in the field.

This study, which is novel for Nebraska, can impact future conservation and inoculative biological control programs in the state. 29

References

Andow, D. A., S. G. Pueppke, A. W. Schaafsma, A. J. Gassmann, T. W. Sappington, L. J. Meinke, P. D. Mitchell, T. M. Hurley, R. L. Hellmich, and R. P. Porter. 2016. Early detection and mitigation of resistance to Bt maize by western corn rootworm (Coleoptera: Chrysomelidae). J. Econ. Entomol. 109: 1–12.

Ball, H.J. and G. T. Weekman. 1963. Differential resistance of corn rootworms to insecticides in Nebraska and adjoining states. J. of Econ. Entomol. 56: 553–555

Batista Filho, A., J. E. M. Almeida, and C. Lamas. 2001. Effect of thiamethoxam on entomopathogenic microorganisms. Neotrop. Entomol. 30: 437–447.

Bledsoe, L.W., and Obermeyer, J.L. 2010. Managing Corn Rootworms. Purdue University Extension. Publication E-49-W

Branson, T. F. 1987. The contribution of prehatch and posthatch development to protandry in the chrysomelid, Diabrotica virgifera virgifera. Entomol. Exp. Appl. 43: 205–208.

Branson, T. F. 1989. Survival of starved neonate larvae of Diabrotica virgifera virgifera LeConte (Coleoptera: Chrysomelidae). J. Kans. Entomol. Soc. 62: 521–523.

Branson, T. F., and G. R. Sutter. 1985. Influence of population density of immatures on size, longevity, and fecundity of adult Diabrotica virgifera virgifera (Coleoptera: Chrysomelidae). Environ. Entomol. 14: 687–690.

Branson, T. F., G. R. Sutter, and J. R. Fisher. 1982. Comparison of a tolerant and a susceptible maize inbred under artificial infestations of Diabrotica virgifera virgifera: yield and adult emergence. Environ. Entomol. 11: 371–372.

Bruck, D. J., and L. C. Lewis. 2001. Adult Diabrotica spp. (Coleoptera: Chrysomelidae) infection at emergence with indigenous Beauveria bassiana (Deuteromycotina: Hyphomycetes). J. Invertebr. Pathol. 77: 288–289.

Campbell, L. A., and L. J. Meinke. 2006. Seasonality and Adult habitat use by four Diabrotica species at prairie-corn interfaces. Environ. Entomol. 35: 922–936.

Chege, P. G., T. L. Clark, and B. E. Hibbard. 2005. Alternate host phenology affects survivorship, growth, and development of western corn rootworm (Coleoptera: Chrysomelidae) Larvae. Environ. Entomol. 34: 1441–1447.

Chu, C.-C., J. L. Spencer, M. J. Curzi, J. A. Zavala, and M. J. Seufferheld. 2013. Gut bacteria facilitate adaptation to crop rotation in the western corn rootworm. Proc. Natl. Acad. Sci. 110: 11917–11922. 30

Clark, T. L., and B. E. Hibbard. 2004. Comparison of nonmaize hosts to support western corn rootworm (Coleoptera: Chrysomelidae) larval biology. Environ. Entomol. 33: 681–689.

DeVries, T. A., and R. J. Wright. 2016. Evaluation of foliar applied insecticides for control of adult corn rootworm in corn, 2015. Arthropod Manag. Tests. 41.

DiFonzo, C. 2017. Handy Bt trait table. Michigan State University.

Eilenberg, J., A. Hajek, and C. Lomer. 2001. Suggestions for unifying the terminology in biological control. BioControl. 46: 387–400.

Eisley, J. B., and R. B. Hammond. 2008. Corn rootworm larval control with soil insecticides. Arthropod Manag. Tests. 33 (1). F19.

Fisher, J. R. 1986. Development and survival of pupae of Diabrotica virgifera virgifera and D. undecimpunctata howardi (Coleoptera: Chrysomelidae) at constant temperatures and humidities. Environ. Entomol. 15: 626–630.

Gassmann, A. J., Y. Carrière, and B. E. Tabashnik. 2009a. Fitness costs of insect resistance to Bacillus thuringiensis. Annu. Rev. Entomol. 54: 147–163.

Gassmann, A. J., J. A. Fabrick, M. S. Sisterson, E. R. Hannon, S. P. Stock, Y. Carrière, and B. E. Tabashnik. 2009b. Effects of pink bollworm resistance to Bacillus thuringiensis on phenoloxidase activity and susceptibility to entomopathogenic nematodes. J. Econ. Entomol. 102: 1224–1232.

Gassmann, A. J., E. R. Hannon, M. S. Sisterson, S. P. Stock, Y. Carrière, and B. E. Tabashnik. 2012. Effects of entomopathogenic nematodes on evolution of pink bollworm resistance to Bacillus thuringiensis Toxin Cry1Ac. J. Econ. Entomol. 105: 994–1005.

Gassmann, A. J., J. L. Petzold-Maxwell, E. H. Clifton, M. W. Dunbar, A. M. Hoffmann, D. A. Ingber, and R. S. Keweshan. 2014. Field-evolved resistance by western corn rootworm to multiple Bacillus thuringiensis toxins in transgenic maize. Proc. Natl. Acad. Sci. 111: 5141–5146.

Gassmann, A. J., J. L. Petzold-Maxwell, R. S. Keweshan, and M. W. Dunbar. 2011. Field- evolved resistance to bt maize by western corn rootworm. PLoS ONE. 6: e22629.

Gassmann, A. J., S. P. Stock, Y. Carrière, and B. E. Tabashnik. 2006. Effect of entomopathogenic nematodes on the fitness cost of resistance to bt toxin Cry1Ac in pink bollworm (Lepidoptera: Gelechiidae). J. Econ. Entomol. 99: 920– 926. 31

Gassmann, A. J., R. B. Shrestha, S. R. K. Jakka, M. W. Dunbar, E. H. Clifton, A. R. Paolino, D. A. Ingber, B. W. French, K. E. Masloski, J. W. Dounda, and C. R. St. Clair. 2016. Evidence of resistance to Cry34/35Ab1 corn by western corn rootworm (Coleoptera: Chrysomelidae): root injury in the field and larval survival in plant-based bioassays. J. Econ. Entomol. 109: 1872-1880.

Gassmann, A. J., S. P. Stock, M. S. Sisterson, Y. Carrière, and B. E. Tabashnik. 2008. Synergism between entomopathogenic nematodes and Bacillus thuringiensis crops: integrating biological control and resistance management. J. Appl. Ecol. 45: 957–966.

Gaugler, R., T. McGuire, and J. Campbell. 1989. Genetic variability among strains of the entomopathogenic nematode Steinernema feltiae. J. Nematol. 21: 247–253.

Godfrey, L. D., L. J. Meinke, and R. J. Wright. 1993. Vegetative and Reproductive biomass accumulation in field com: response to root injury by western com rootworm (Coleoptera: Chrysomelidae). J. Econ. Entomol. 86: 1557–1573.

Godfrey, L. D., L. J. Meinke, R. J. Wright, and G. L. Hein. 1995. Environmental and edaphic effects on western corn rootworm (Coleoptera: Chrysomelidae) overwintering egg survival. J. Econ. Entomol. 88: 1445–1454.

Gray, M. E., G. L. Hein, M. A. Boetel, and D. D. Walgenbach. 1992. Western and northern corn rootworm (Coleoptera: Chrysomelidae) egg densities at three soil depths: implications for future ecological studies. J. Kans. Entomol. Soc. 354– 356.

Gray, M. E., T. W. Sappington, N. J. Miller, J. Moeser, and M. O. Bohn. 2009. Adaptation and invasiveness of western corn rootworm: intensifying research on a worsening pest. Annu. Rev. Entomol. 54: 303–321.

Gray, M. E., and K. L. Steffey. 1998. Corn rootworm (Coleoptera: Chrysomelidae) larval injury and root compensation of 12 maize hybrids: an assessment of the economic injury index. J. Econ. Entomol. 91: 723–740.

Gray, M. E., and J. J. Tollefson. 1988. Influence of tillage systems on egg populations of western and northern corn rootworms (Coleoptera: Chrysomelidae). J. Kans. Entomol. Soc. 61: 186–194.

Grewal, P. S., R. Gaugler, and S. Selvan. 1993. Host recognition by entomopathogenic nematodes: behavioral response to contact with host feces. J. Chem. Ecol. 19: 1219–1231.

Grewal, P. S., E. E. Lewis, R. Gaugler, and J. F. Campbell. 1994. Host finding behaviour as a predictor of foraging strategy in entomopathogenic nematodes. Parasitology. 108: 207–215. 32

Gustin, R. D. 1979. Effect of two moisture and population levels on oviposition of the western corn rootworm. Environ. Entomol. 8: 406–407.

Gustin, R. D. 1981. Soil temperature environment of overwintering western corn rootworm eggs. Environ. Entomol. 10: 483–487.

Hajek, A. E. 2004. Natural enemies: an introduction to biological control. Cambridge University Press.

Hammack, L. 1995. Calling behavior in female western corn rootworm beetles (Coleoptera: Chrysomelidae). Ann. Entomol. Soc. Am. 88: 562–569.

Hannon, E. R., M. S. Sisterson, S. P. Stock, Y. Carrière, B. E. Tabashnik, and A. J. Gassmann. 2010. Effects of four nematode species on fitness costs of pink bollworm resistance to Bacillus thuringiensis toxin Cry1Ac. J. Econ. Entomol. 103: 1821–1831.

Hibbard, B. E., and L. B. Bjostad. 1988. Behavioral responses of western corn rootworm larvae to volatile semiochemicals from corn seedlings. J. Chem. Ecol. 14: 1523– 1539.

Hibbard, B. E., L. N. Meihls, M. R. Ellersieck, and D. W. Onstad. 2010. Density- dependent and density-independent mortality of the western corn rootworm: impact on dose calculations of rootworm-resistant Bt corn. J. Econ. Entomol. 103: 77–84.

Hill, R. E. 1975. Mating, Oviposition patterns, fecundity and longevity of the western corn rootworm. J. Econ. Entomol. 68: 311–315.

Hoback, W. W., T. L. Clark, L. J. Meinke, L. G. Higley, and J. M. Scalzitti. 2002. Immersion survival differs among three Diabrotica species. Entomol. Exp. Appl. 105: 29–34.

Hoffmann, A. M., B. W. French, S. T. Jaronski, and A. J. Gassmann. 2014. Effects of entomopathogens on mortality of western corn rootworm (Coleoptera: Chrysomelidae) and fitness costs of resistance to Cry3Bb1 maize. J. Econ. Entomol. 107: 352–360.

Jackson, J. J. 1996. Field performance of entomopathogenic nematodes for suppression of western corn rootworm (Coleoptera: Chrysomelidae). J. Econ. Entomol. 89: 366–372.

Jackson, J. J., and M. A. Brooks. 1989. Susceptibility and immune response of western corn rootworm larvae (Coleoptera: Chrysomelidae) to the entomogenous nematode, Steinernema feltiae (Rhahditida: Steinernematidae). J. Econ. Entomol. 82: 1073–1077. 33

Jackson, J. J., and M. A. Brooks. 1995. Parasitism of western corn rootworm larvae and pupae by Steinernema carpocapsae. J. Nematol. 27: 15–20.

Journey, A. M., and K. R. Ostlie. 2000. Biological control of the western corn rootworm (Coleoptera: Chrysomelidae) using the entomopathogenic nematode, Steinernema carpocapsae. Environ. Entomol. 29: 822–831.

Kahler, A. L., A. E. Olness, G. R. Sutter, C. D. Dybing, and O. J. Devine. 1985. Root damage by western corn rootworm and nutrient content in maize. Agron. J. 77: 769.

Kaya, H. K., and R. Gaugler. 1993. Entomopathogenic nematodes. Annu. Rev. Entomol. 38: 181–206.

Kirk, V. M. 1979. Drought Cracks as oviposition sites for western and northern corn rootworms (Diabrotica: Coleoptera). J. Kans. Entomol. Soc. 52: 769–776.

Kirk, V. M. 1981. Corn rootworm: population reduction associated with the ant, Lasius neoniger. Environ. Entomol. 10: 966–967.

Kogan, M., and E. F. Ortman. 1978. Antixenosis–a new term proposed to define Painter's “nonpreference” modality of resistance. Bulletin of the ESA, 24(2): 175- 176.

Koppenhöfer, A. M. 2007. Nematodes. In field manual of techniques in invertebrate pathology. pp. 249-264. Springer, Dordrecht.

Krupke, C.H., Obermeyer, J.L., and Bledsoe Larry. 2014. Corn insect control recommendations. Purdue University Extension. Publication E-219-W.

Krysan, J. L. 1978. Diapause, quiescence, and moisture in the egg of the western corn rootworm, Diabrotica virgifera. J. Insect Physiol. 24: 535–540.

Krysan, J. L., J. J. Jackson, and A. C. Lew. 1984. Field termination of egg diapause in diabrotica with new evidence of extended diapause in D. barberi (Coleoptera: Chrysomelidae). Environ. Entomol. 13: 1237–1240.

Kuhlmann, U., and Van der Burgt, A. C. M. 1998. Possibilities for biological control of the western corn rootworm, Diabrotica virgifera virgifera LeConte, in central Europe. Biocontrol News and Information. 19: 59N-68N.

Kuhlmann, U., Toepfer, S., and F. Zhang. 2005. Is classical biological control against western corn rootworm in Europe a potential sustainable management strategy?, pp. 263-284. In Vidal, Stefan, Ulrich Kuhlmann, and C. Richard Edwards, eds. Western corn rootworm: ecology and management. CABI, Oxfordshire, UK. 34

Kurtz, B., I. Hiltpold, T. C. J. Turlings, U. Kuhlmann, and S. Toepfer. 2009. Comparative susceptibility of larval instars and pupae of the western corn rootworm to infection by three entomopathogenic nematodes. BioControl. 54: 255–262.

Landis, D. A., S. D. Wratten, and G. M. Gurr. 2000. Habitat Management to Conserve Natural Enemies of Arthropod Pests in Agriculture. Annu. Rev. Entomol. 45: 175– 201.

Levine, E., and H. Oloumi-Sadeghi. 1996. Western corn rootworm (Coleoptera: Chrysomelidae) larval injury to corn grown for seed production following soybeans grown for seed production. J. Econ. Entomol. 89: 1010–1016.

Levine, E., H. Oloumi-Sadeghi, and C. R. Ellis. 1992. Thermal requirements, hatching patterns, and prolonged diapause in western corn rootworm (Coleoptera: Chrysomelidae) eggs. J. Econ. Entomol. 85: 2425–2432.

Levine, E., J. L. Spencer, S. A. Isard, D. W. Onstad, and M. E. Gray. 2002. Adaptation of the western corn rootworm to crop rotation: evolution of a new strain in response to a management practice. Am. Entomol. 48: 94–107.

Lewis, E. E., J. Campbell, C. Griffin, H. Kaya, and A. Peters. 2006. Behavioral ecology of entomopathogenic nematodes. Biol. Control. 38: 66–79.

Ludwick, D. C., L. N. Meihls, K. R. Ostlie, B. D. Potter, L. French, and B. E. Hibbard. 2017. Minnesota field population of western corn rootworm (Coleoptera: Chrysomelidae) shows incomplete resistance to Cry34Ab1/Cry35Ab1 and Cry3Bb1. J. Appl. Entomol. 141: 28-40.

Lundgren, J. G., M. E. Ellsbury, and D. A. Prischmann. 2009. Analysis of the predator community of a subterranean herbivorous insect based on polymerase chain reaction. Ecol. Appl. 19: 2157–2166.

Lundgren, J. G., and J. K. Fergen. 2014. Predator community structure and trophic linkage strength to a focal prey. Mol. Ecol. 23: 3790–3798.

Lundgren, J. G., T. Haye, S. Toepfer, and U. Kuhlmann. 2009. A multifaceted hemolymph defense against predation in Diabrotica virgifera virgifera larvae. Biocontrol Sci. Technol. 19: 871–880.

Mabry, T. R., and J. L. Spencer. 2003. Survival and oviposition of a western corn rootworm variant feeding on soybean. Entomol. Exp. Appl. 109: 113–121.

Macdonald, P. J., and C. R. Ellis. 1990. Survival time of unfed, first-instar western corn rootworm (Coleoptera: Chrysomelidae) and the effects of soil type, moisture, and compaction on their mobility in soil. Environ. Entomol. 19: 666–671. 35

Manachini, B. 2002. Compatibility of chemical and biological pesticides, pp. 134–137. In Pimentel, D. (ed.), Encycl. Pest Manag. CRC Press.

Maredia, K., and D. Landis. 1993. Corn rootworms: biology, ecology, and management. Ext. Bull. E-2438.

Marquardt, P. T., and C. H. Krupke. 2009. Dispersal and mating behavior of Diabrotica virgifera virgifera (Coleoptera: Chrysomelidae) in Bt cornfields. Environ. Entomol. 38: 176–182.

Meinke, L. J., T. W. Sappington, D. W. Onstad, T. Guillemaud, N. J. Miller, J. Komáromi, N. Levay, L. Furlan, J. Kiss, and F. Toth. 2009. Western corn rootworm (Diabrotica virgifera virgifera LeConte) population dynamics. Agric. For. Entomol. 11: 29–46.

Meinke, L. J., B. D. Siegfried, R. J. Wright, and L. D. Chandler. 1998. Adult susceptibility of Nebraska western corn rootworm (Coleoptera: Chrysomelidae) populations to selected insecticides. J. Econ. Entomol. 91: 594–600.

Meissle, M., C. Pilz, and J. Romeis. 2009. Susceptibility of Diabrotica virgifera virgifera (Coleoptera: Chrysomelidae) to the entomopathogenic fungus Metarhizium anisopliae when feeding on Bacillus thuringiensis Cry3Bb1-expressing maize. Appl. Environ. Microbiol. 75: 3937–3943.

Moeser, J. and B.E. Hibbard. 2005. A synopsis on the nutritional ecology of larvae and adults of Diabrotica virgifera virgifera (LeConte) in the new and old world- nouvelle cuisine for the invasive maize pest Diabrotica virgifera virgifera in Europe? In western corn rootworm: ecology and management. Wallingford, United Kingdom:CABI publishers. p. 41-65.

Mulock, B. S., and L. D. Chandler. 2001. Effect of Beauveria bassiana on the fecundity of western corn rootworm, Diabrotica virgifera virgifera (Coleoptera: Chrysomelidae). Biol. Control. 22: 16–21.

Neves, P. M. O. J., E. Hirose, P. T. Tchujo, and A. Moino Jr. 2001. Compatibility of entomopathogenic fungi with neonicotinoid insecticides. Neotrop. Entomol. 30: 263–268.

Nishimatsu, T., and J. J. Jackson. 1998. Interaction of insecticides, entomopathogenic nematodes, and larvae of the western corn rootworm (Coleoptera: Chrysomelidae). J. Econ. Entomol. 91: 410–418.

O’Neal, M. E., M. E. Gray, S. Ratcliffe, and K. L. Steffey. 2001. Predicting western corn rootworm (Coleoptera: Chrysomelidae) larval injury to rotated corn with Pherocon AM traps in soybeans. J. Econ. Entomol. 94: 98–105. 36

Onstad, D. W., D. W. Crowder, P. D. Mitchell, C. A. Guse, J. L. Spencer, E. Levine, and M. E. Gray. 2003. Economics versus alleles: balancing integrated pest management and insect resistance management for rotation-resistant western corn rootworm (Coleoptera: Chrysomelidae). J. Econ. Entomol. 96: 1872–1885.

Onstad, D. W., B. E. Hibbard, T. L. Clark, D. W. Crowder, and K. G. Carter. 2006. Analysis of density-dependent survival of Diabrotica (Coleoptera: Chrysomelidae) in cornfields. Environ. Entomol. 35: 1272–1278.

Ortman, E. E., T. F. Branson, E. D. Gerloff, and S. Dakota. 1974. Techniques, accomplishments, and future potential of host plant resistance to Diabrotica. Proc. Summer Inst. Biol. Control Plant Insects Dis. Maxwell.

Oyediran, I. O., B. E. Hibbard, and T. L. Clark. 2004. Prairie grasses as hosts of the western corn rootworm (Coleoptera: Chrysomelidae). Environ. Entomol. 33: 740–747.

Painter, R. H. 1951. Insect resistance in crop plants. 520 pp.

Parimi, S., L. J. Meinke, B. Wade French, L. D. Chandler, and B. D. Siegfried. 2006. Stability and persistence of aldrin and methyl-parathion resistance in western corn rootworm populations (Coleoptera: Chrysomelidae). Crop Prot. 25: 269– 274.

Pereira, A. E., H. Wang, S. N. Zukoff, L. J. Meinke, B. W. French, and B. D. Siegfried. 2015. Evidence of field-evolved resistance to bifenthrin in western corn rootworm (Diabrotica virgifera virgifera LeConte) populations in western Nebraska and Kansas. PLoS ONE. 10: e0142299.

Petzold-Maxwell, J. L., X. Cibils-Stewart, B. W. French, and A. J. Gassmann. 2012. Adaptation by western corn rootworm (Coleoptera: Chrysomelidae) to Bt maize: inheritance, fitness costs, and feeding preference. J. Econ. Entomol. 105: 1407– 1418.

Petzold–Maxwell, J. L., S. T. Jaronski, E. H. Clifton, M. W. Dunbar, M. A. Jackson, and A. J. Gassmann. 2013. Interactions among bt maize, entomopathogens, and rootworm species (Coleoptera: Chrysomelidae) in the field: effects on survival, yield, and root injury. J. Econ. Entomol. 106: 622–632.

Petzold-Maxwell, J. L., S. T. Jaronski, and A. J. Gassmann. 2012. Tritrophic interactions among Bt maize, an insect pest and entomopathogens: effects on development and survival of western corn rootworm. Ann. Appl. Biol. 160: 43–55.

Pierce, C. M. F., and M. E. Gray. 2006. Seasonal oviposition of a western corn rootworm, Diabrotica virgifera virgifera LeConte (Coleoptera: Chrysomelidae), 37

variant in east central Illinois commercial maize and soybean fields. Environ. Entomol. 35: 676–683.

Pierce, C. M. F., and M. E. Gray. 2007. Population dynamics of a western corn rootworm (Coleoptera: Chrysomelidae) variant in east central Illinois commercial maize and soybean fields. J. Econ. Entomol. 100: 1104–1115.

Pilz, C., S. Keller, U. Kuhlmann, and S. Toepfer. 2009. Comparative efficacy assessment of fungi, nematodes and insecticides to control western corn rootworm larvae in maize. BioControl. 54: 671–684.

Pilz, C., R. Wegensteiner, and S. Keller. 2007. Selection of entomopathogenic fungi for the control of the western corn rootworm Diabrotica virgifera virgifera. J. Appl. Entomol. 131: 426–431.

Purdue University. 2009. Corn rootworms. Purdue University Field Crops IPM. http://extension.entm.purdue.edu/fieldcropsipm/insects/corn-rootworms.php

Rasmann, S., T. G. Köllner, J. Degenhardt, I. Hiltpold, S. Toepfer, U. Kuhlmann, J. Gershenzon, and T. C. J. Turlings. 2005. Recruitment of entomopathogenic nematodes by insect-damaged maize roots. Nature. 434: 732–737.

Riedell, W. E. 1990. Rootworm and mechanical damage effects on root morphology and water relations in maize. Crop Sci. 30: 628.

Riedell, W. E., and P. D. Evenson. 1993. Rootworm feeding tolerance in single-cross maize hybrids from different eras. Crop Sci. 33: 951–955.

Roth, G.W. 1996. Crop rotations and conservation tillage. Penn State Extension. UC124

Rovesti, L., and K. V. Deseö. 1990. Compatibility of chemical pesticides with the entomopathogenic nematodes, Steinernema carpocapsae Weiser and S. feltiae Filipjev (Nematoda: Steinernematidae). Nematologica. 36: 237–245.

Rovesti, L., and K. V. Deseö. 1991. Compatibility of pesticides with the entomopathogenic nematode, Heterorhabditis heliothidis. Nematologica. 37: 113–116.

Rovesti, L., E. W. Heinzpeter, F. Tagliente, and K. V. Deseö. 1988. Compatibility of pesticides with the entomopathogenic nematode Heterorhabditis bacteriophora Poinar (Nematoda: Heterorhabditidae). Nematologica. 34: 462–476.

Rudeen, M. L., and A. J. Gassmann. 2013. Effects of Cry34/35Ab1 corn on the survival and development of western corn rootworm, Diabrotica virgifera virgifera. Pest Manag. Sci. 69: 709–716. 38

Rudeen, M. L., Jaronski, S. T., Petzold-Maxwell, J. L., and Gassmann, A. J. 2013. Entomopathogenic fungi in cornfields and their potential to manage larval western corn rootworm Diabrotica virgifera virgifera. J. of Invertebrate Pathology. 114(3): 329-332.

Schrader, P. M., R. E. Estes, N. A. Tinsley, A. J. Gassmann, and M. E. Gray. 2016. Evaluation of adult emergence and larval root injury for Cry3Bb1-resistant populations of the western corn rootworm. J. Appl. Entomol. 141: 41-52.

Shah, F. A., M. A. Ansari, J. Watkins, Z. Phelps, J. Cross, and T. M. Butt. 2009. Influence of commercial fungicides on the germination, growth and virulence of four species of entomopathogenic fungi. Biocontrol Sci. Technol. 19: 743–753.

Shields, E. J. 2015. Utilizing persistent entomopathogenic nematodes in a conservation or a more classical biological control approach, pp. 165–184. In Nematode Pathog. Insects pests, sustainability in plant and crop protection. Springer, Cham.

Shields, E. J., and A. M. Testa. 2015. New York Case Study: biological control of Otiorhynchus ligustici with native persistent entomopathogenic nematodes using a more classical approach, pp. 285–307. In Nematode Pathog. Insects Pests, Sustainability in Plant and Crop Protection. Springer, Cham.

Spencer, J. L., B. E. Hibbard, J. Moeser, and D. W. Onstad. 2009. Behaviour and ecology of the western corn rootworm (Diabrotica virgifera virgifera LeConte). Agric. For. Entomol. 11: 9–27.

Stock, S. P., and Goodrich-Blair, H. 2012. Nematode parasites, pathogens and associates of insects and invertebrates of economic importance. In Manual of techniques in invertebrate pathology. 375-425. Academic press, CA.

Strnad, S. P., and M. K. Bergman. 1987. Movement of first-instar western corn rootworms (Coleoptera: Chrysomelidae) in Soil. Environ. Entomol. 16: 975–978.

Toepfer, S., T. Haye, M. Erlandson, M. Goettel, J. G. Lundgren, R. G. Kleespies, D. C. Weber, G. C. Walsh, A. Peters, R.-U. Ehlers, H. Strasser, D. Moore, S. Keller, S. Vidal, and U. Kuhlmann. 2009. A review of the natural enemies of beetles in the subtribe Diabroticina (Coleoptera: Chrysomelidae): implications for sustainable pest management. Biocontrol Sci. Technol. 19: 1–65.

Toepfer, S., A. Peters, R.-U. Ehlers, and U. Kuhlmann. 2008. Comparative assessment of the efficacy of entomopathogenic nematode species at reducing western corn rootworm larvae and root damage in maize. J. Appl. Entomol. 132: 337–348.

Toepfer, S., G. C. Walsh, A. Eben, R. Alvarez-Zagoya, T. Haye, F. Zhang, and U. Kuhlmann. 2008. A critical evaluation of host ranges of parasitoids of the 39

subtribe Diabroticina (Coleoptera: Chrysomelidae: Galerucinae: Luperini) using field and laboratory host records. Biocontrol Sci. Technol. 18: 483–504.

Toepfer, S., F. Zhang, J. Kiss, And U. Kuhlmann. 2005. The invasion of the western corn rootworm, Diabrotica virgifera virgifera, in Europe and potential for classical biological control. In second Int. Symp. Biol. control arthropods.

Turpin, F. T., and D. C. Peters. 1971. Survival of southern and western corn rootworm larvae in relation to soil texture. J. Econ. Entomol. 64: 1448–1451.

Urías-López, M. A., and L. J. Meinke. 2001. Influence of western corn rootworm (Coleoptera: Chrysomelidae) larval injury on yield of different types of maize. J. Econ. Entomol. 94: 106–111.

Urías-López, M. A., L. J. Meinke, L. G. Higley, and F. J. Haile. 2000. Influence of western corn rootworm (Coleoptera: Chrysomelidae) larval injury on photosynthetic rate and vegetative growth of different types of maize. Environ. Entomol. 29: 861– 867.

(USEPA) US Environmental Protection Agency. 2001. Biopesticides registration action document: Bacillus thuringiensis plant-incorporated protectants (10/16/01) (https://www3.epa.gov/pesticides/chem_search/reg_actions/pip/bt_brad.htm) (accessed 2 April 2018),

(USEPA) US Environmental Protection Agency. 2010. Bacillus thuringiensis Cry3Bb1 protein and the genetic material necessary for its production (Vector PV- ZMIR13L) in MON863 corn (OECD Unique Identifier: MON-00863-5) PC Code: 006484 Bacillus thuringiensis Cry3Bb1 protein and the genetic material necessary for its production (Vector PV-ZMIR39) in MON 88017 corn (OECD Unique Identifier: MON-88017-3) PC Code: 006498 (https://www3.epa.gov/ pesticides/chem_search/reg_actions/pip/cry3bb1-brad.pdf) (accessed 2 Apr 2018).

Wangila, D. S., A. J. Gassmann, J. L. Petzold-Maxwell, B. W. French, and L. J. Meinke. 2015. Susceptibility of Nebraska western corn rootworm (Coleoptera: Chrysomelidae) populations to Bt corn events. J. Econ. Entomol. 108: 742-751.

Wangila, D. S., and L. J. Meinke. 2017. Effects of adult emergence timing on susceptibility and fitness of Cry3Bb1-resistant western corn rootworms. J. Appl. Entomol. 141: 372–383.

Weiss, M. J., Z. B. Mayo, and J. P. Newton. 1983. Influence of irrigation practices on the spatial distribution of corn rootworm (Coleoptera: Chrysomelidae) eggs in the soil. Environ. Entomol. 12: 1293–1295. 40

Weiss, M. J., K. P. Seevers, and Z. B. Mayo. 1985. Influence of western corn rootworm larval densities and damage on corn rootworm survival, developmental time, size and sex ratio (Coleoptera: Chrysomelidae). J. Kans. Entomol. Soc. 58: 397–402.

Wilde, G., H. C. Chiang, E. T. Hibbs, and D. E. Lawson. 1972. Variation in egg hatch among western and northern corn rootworms from six midwestern states. J. Kans. Entomol. Soc. 259–263.

Wilde, G. E. 1971. Temperature effect on development of western corn rootworm eggs. J. Kans. Entomol. Soc. 185–187.

Wilson, R. L., and D. C. Peters. 1973. Plant introductions of Zea mays as sources of corn rootworm tolerance. J. Econ. Entomol. 66: 101–104.

Wilson, T. A., and B. E. Hibbard. 2004. Host suitability of nonmaize agroecosystem grasses for the western corn rootworm (Coleoptera: Chrysomelidae). Environ. Entomol. 33: 1102–1108.

Wraight, S.P., G.D. Inglis, and M.S. Goettel. 2007. Overview of pathogen groups-fungi, pp.223-249. In Field manual of techniques in invertebrate pathology Lacey, L. A., and Kaya, H. K. (Eds.). Dordrecht etc.: Kluwer acad. publ. Cop.

Wright, R. J., Witkowski, J. F., Echtenkamp, G., and Georgis, R. 1993. Efficacy and persistence of Steinernema carpocapsae (Rhabditida: Steinemematidae) applied through a center-pivot irrigation system against larval com rootworms (Coleoptera: Chrysomelidae). J. of Economic Entomology. 86(5):1348-1354.

Zukoff, S. N., K. R. Ostlie, B. Potter, L. N. Meihls, A. L. Zukoff, L. French, M. R. Ellersieck, B. W. French, and B. E. Hibbard. 2016. Multiple assays indicate varying levels of cross resistance in Cry3Bb1-selected field populations of the western corn rootworm to mCry3A, eCry3.1Ab, and Cry34/35Ab1. J. Econ. Entomol. 109(3): 1387-1398.

41

CHAPTER 2: SURVEY FOR POTENTIAL ABOVE-GROUND PREDATORS OF THE WESTERN CORN ROOTWORM WITH EMPHASIS ON GROUND BEETLES (COLEOPTERA: CARABIDAE).

Introduction

The western corn rootworm (WCR), Diabrotica virgifera virgifera LeConte

(Coleoptera: Chrysomelidae), is the main root-feeding pest of continuous corn production in the United States and Europe. This pest is present in the field all year-long.

Eggs typically start hatching in late May (Meinke et al. 2009), going through three larval instars and a pupal stage before adult emergence. Adults emerge from the end of June through September, with delayed mean emergence periods occurring in transgenic

Bacillus thuringiensis (Bt) hybrid fields (Hitchon et al. 2015). Oviposition occurs from

July-September (Meinke et al. 2009). Corn is the primary host of the WCR (Clark and

Hibbard 2004) and, therefore, this pest is mainly a problem in continuous corn, except in limited geographic regions where behavioral resistance to rotation has occurred (Gray et al. 2009).

Current management tactics of WCR include crop rotation, seed treatments, soil and foliar insecticides, and transgenic hybrids. Continuous corn production requires constant pest management, and because of the high management pressure and the adaptive nature of this pest (Gray et al. 2009; Miller et al. 2009), populations of the WCR have evolved resistance to a variety of management practices. Resistance or reduced susceptibility has been reported with crop rotation (Levine et al. 2002); chemical insecticides such as organochlorines (Parimi et al. 2006), pyrethroids (Pereira et al.

2015), organophosphates and carbamates (Meinke et al. 1998); and with Bt proteins 42 such as Cry3Bb1 (Gassmann et al. 2011), with cross resistance to mCry3A (Gassmann et al. 2014) and eCry3.1Ab (Zukoff et al. 2016), and most recently Cry34/35 (Gassmann et al. 2016). Management of WCR populations and resistance issues requires a complex integrated pest management (IPM) approach. Due to the frequency and severity of resistance to current management tactics, investigation of WCR biological control is critical and may lead to an additional management option to integrate with current

WCR control practices.

Individual species as well as assemblages of generalist predators have the potential to suppress pest densities and protect yield (Symondson et al., 2002).

However, in the WCR system, no keystone predator has been identified (Lundgren and

Fergen 2014). A variety of arthropod predators have been reported to prey on the different life stages of the WCR in laboratory and/or field conditions, as reviewed by

Kuhlmann et al. (1998) and Toepfer et al. (2009). Taxa reported as predators of one or more life stages of the WCR include ground beetles (Coleoptera: Carabidae), hister beetles (Coleoptera: Histeridae), rove beetles (Coleoptera: Staphylinidae), carpet beetles (Coleoptera: Dermestidae), ants (Hymenoptera: Formicidae), robber flies

(Diptera: Asilidae), crickets (Orthoptera: Gryllidae), katydids (Orthoptera: Tettigoniidae), predatory mites (Mesostigmata: Laelapidae), several families of spiders (Araneae), harvestmen (Opiliones: Phalangiidae), centipedes (Chilopoda) and isopods (Isopoda:

Armadillidiidae).

Diversity, evenness and abundance of the predator community are driving factors in rootworm consumption in the field (Lundgren and Fergen 2014). Western 43 corn rootworm is considered sub-optimal prey in cornfields due to behavioral and physiological defenses (Lundgren and Fergen 2014), including hemolymph defense by the larvae that may deter chewing predators (Lundgren et al. 2009b). In addition, larvae exhibit a cryptic lifestyle in which the early instars tunnel and feed within the root system (Strnad and Bergman 1987b) and are therefore protected from many soil antagonists.

Molecular gut content analyses (MGCA) indicate that several species of

Carabidae can prey on WCR eggs and larvae (Lundgren et al. 2009a, Lundgren and

Fergen 2014). Larvae and adults of Scarites quadriceps Chaudoir as well as adult Poecilus chalcites (Say) had relatively high consumption rates and predation frequencies of WCR immatures as 20.4% and 17.5% of individuals of each species tested positive for WCR

DNA in MGCA (Lundgren et al. 2009a). However, visual observations indicated that

Carabidae are incidental predators of rootworms, with predation dependent upon chance encounters instead of searching for prey (Kirk 1982). Therefore, while some carabids have been confirmed as WCR predators, their role in the field might be minimal. But in conjunction with other pest management strategies in place, predators may provide an additional tool to minimize population pressure and resistance issues.

Nebraska is the third largest corn producer in the U.S.A. (USDA-NASS 2017) and the WCR is one of the state’s biggest insect pests. Predation of any life stage of this pest can aid plant protection in the long term since this is a continuous corn pest. The objectives of this study were to a) characterize the community of arthropod predators 44 and to b) identify potential carabid predators of the WCR via molecular gut content analysis (MGCA).

Materials and Methods

Field Sites. This study was conducted in 2014 in five commercial, irrigated, no-till cornfields located in West Central Nebraska. Four sites had been planted as continuous corn for at least 5 years prior to this study (Fields A-D) and one site had been rotated to winter wheat in 2013 (Field E). Agronomic characteristics of the field sites are listed on

Appendix Table 1.

Collection of Predatory Arthropods and WCR. Sites were surveyed for arthropods and prey availability on seven dates throughout the corn-growing season (June-September).

Four unbaited Pheroconâ AM yellow sticky cards (YSC) (Hein and Tollefson 1985) were placed at canopy height at early vegetative stages and then placed at ear-height for the rest of the season. Yellow sticky cards were left out in the field for seven days and then were placed in the refrigerator until processing. In addition, rootworm densities were also monitored in each field with eight single-plant adult emergence cages (Pierce and

Gray 2007). Emergence cages were checked weekly during the adult emergence period from July.15 to September.26, for a total of 11 collection dates. Sampling units (YSC and emergence cages) were placed 8 to 15 meters from each other and were located between 60-120 meters from the edge of the irrigated field.

Carabidae Collection for Molecular Gut Content Analysis. Five dry pitfall traps were located at each site: four inside the irrigated cornfield and one in the non-irrigated border. Traps were opened for 24-hrs per collection period. On average, traps were 45 opened around noon and closed at noon the next day. A hardware cloth insert (3.2 mm mesh) was placed inside dry pitfall traps (946 ml vol, 11.5 X 14 cm WXH) (Eskelson et al.

2011) to prevent bigger predators from consuming smaller arthropods. Arthropods trapped in the dry pitfall were individually placed into 1.5 ml microcentrifuge tubes

(Fisherbrand™, Pittsburgh, PA, USA) and preserved in 95% EtOH in the field. The samples were then labeled and placed into an Engel 40 portable freezer (Big Frog

Mountain, Chattanooga, TN, USA) in the field, and subsequently frozen at -20 °C until

MGCA. Morphological identification was carried out for each specimen prior to DNA extraction using dichotomous keys found in Arnett & Thomas (2000) and Lindroth

(1961-1969).

Molecular Gut Content Analysis: Gut-content DNA extractions were performed using

QIAGEN DNeasy Blood & Tissue Kits (QIAGEN Inc., Chatsworth, CA, USA). Specimens under 1.0 cm were extracted whole and specimens larger than 1.0 cm had gut dissections performed prior to DNA extraction. Polymerase chain reactions (PCR) were conducted with a T100 Bio-Rad thermocycler (Bio-Rad Laboratories, Inc., Hercules, CA,

USA) with a WCR-specific primer pair (E-F364 and G-R358) targeting the cytochrome c oxidase 1 gene (Peterson 2012). The primer set was tested against eggs (n=5), larvae

(n=2) and adults (n=5) of the WCR to ensure that it could amplify all stages of this pest.

Each PCR reaction contained 17.4 μl of PCR-grade water, 2.5 μl 10X Takara buffer

(Takara Bio Inc., Shiga, Japan), 2.0 μl Takara dNTPs, 0.5 μl of each primer, 0.125 μl

Takara Taq Polymerase, and 2.0 μl of DNA template per sample. PCR cycling protocol included an initial denaturation step at 94 °C for 1 min; then 45 cycles of denaturation at 46

94 °C for 45 s, annealing at 66 °C for 45 s, and extension at 72 °C for 30 s; and final extension at 72 °C for 10 min. Gel electrophoresis amplification verification was performed on 2% agarose gels in 1× Tris-acetate-EDTA (TAE) buffer stained with GelRed

Ô Nucleic Acid Gel Stain (Biotium, Hayward, CA, USA). Visualization of gels was carried out on GelDocä XR+ Gel Documentation system (Bio-Rad Laboratories Inc.).

Data Analysis. Western corn rootworm beetle densities (emergence cage and YSC) were analyzed on SAS (SAS Institute Inc., version 9.4, Cary, NC). Data were fitted to negative- binomial distributions (Tripathi 2006) and analyzed with PROC GLIMMIX. One-way

ANOVAs were analyzed to determine field differences on the mean cumulative count of beetles (between beginning and end collection dates) per emergence cage or per YSC.

Mean estimates and multiple-mean comparisons were obtained with LSMEANS and t- grouping differences were obtained with the LINES option. Mean differences were considered significant at the P < 0.05 level.

Results

Arthropod survey. A total of 866 predators were collected using yellow sticky cards.

Predators were all adults identified as thirteen taxa in five orders: Araneae, Coleoptera,

Hemiptera, Hymenoptera and Neuroptera (Table 2.1). The most abundant taxa were

Chrysopidae, followed by Formicidae, Orius insidiosus, and Araneae. Chrysopidae were the most reported taxa from each field, although they were most abundant in Fields A and D. Formicidae counts were primarily driven by Field D and and O. insidiosus counts driven by Field A. The greatest abundance of predators was found at Field A, followed by Field D, Field B, Field E and lastly, Field C (Table 2.1). 47

WCR density. There was a significant field effect on mean WCR densities per emergence cage (F4, 35 = 20.63, Pr > F < 0.0001) and per YSC (F4,15 = 45.29, Pr > F < 0.0001). Mean cumulative WCR per emergence cage was higher for field A (48.9 ± 12.8) and field C

(33.2 ± 8.8), followed by field B (15.4 ± 4.2), then field D (4.2 ± 1.3), and lastly field E (1.4

± 0.5) (Fig. 2.1). Mean of cumulative WCR beetles per YSC was higher for field A (358.7 ±

83.4) and field C (233.0 ± 54.4), followed by field B (31.7 ± 7.8) and field D (20.7 ± 5.3), and lastly field E (2.5 ± 1.0) (Fig. 2.2).

Carabidae and MGCA. A total of 235 adult carabids were collected, identified, and screened using MGCA to detect WCR DNA (Table 2.3). Thirty-six species were identified belonging to sixteen genera. The most diverse genus was Harpalus with ten species, followed by Anisodactylus, Bembidion and Agonum each with four species. The most abundant species were Anisodactylus sanctaecrucis (Fabricius), Elaphropus anceps

LeConte, and Bembidion quadrimaculatum L.; respectively making up 27.7%, 21.7% and

16.6 % of all specimens encountered (Figure 2.3.). All 235 predator extractions were screened with general COI primers that amplify DNA from members of Arthropoda

(LCO-1490 and HCO-700dy; Folmer et al. 1994) to ensure that DNA extraction had been completed successfully and eliminate any potential false negative results. The preliminary PCR primer test with WCR eggs (n=5), larvae (n=2) and adults (n=5) all yielded strong positive results, indicating successful prey DNA amplification by this primer set. However, none of the Carabidae specimens tested positive for WCR DNA in our gut-content analysis.

48

Discussion

The current study provides new knowledge on the above-ground arthropod predator community and epigeal carabids of no-till, continuous, irrigated corn in West

Central Nebraska. The role of predators found on the YSC (Table 2.1) in rootworm control in the field is likely to be minimal. However, they may be useful for management of other important pests in Nebraska corn production such as the western bean cutworm (WBC), Striacosta albicosta (Smith) (Archibald 2017). Most taxa found on YSC are polyphagous predators but have not been reported in the literature to consume

WCR (Kuhlmann et al. 1998, Toepfer et al. 2009). Moreover, three predators commonly found in the present survey, Coccinella septempunctata L., Hippodamia convergens

Guérin-Méneville, and Orius insidiosus (Say) were positive for WBC DNA in a MGCA and

Coleomegilla maculata was visually confirmed as a predator of WBC eggs in the field

(Archibald 2017). The results presented here along with Archibald (2017) show the potential of exploiting natural enemy communities for insect pest suppression in continuous corn.

It is also important to notice that all of the field sites in this study expressed Bt traits (below-ground and/or above-ground) (Appendix 1). Perkins County has documented Cry3Bb1 WCR resistance (Wangila et al. 2015), and Field A (Keith Co.) and

Field C (Perkins Co.) fields had high densities of WCR (Figures 2.1 and 2.2). Western corn rootworm resistance in the area that we conducted this study stresses even more the need for complementary WCR controls in the region and hence the need for studies like this. All fields had high densities of predators (Table 2.1 and 2.2) which suggests that 49 predator trends were not adversely affected by Bt traits expressed in the field sites. This finding is supported by analyses that found that Bt fields allow abundant communities of naturally occurring predators to exist because of the reduction in insecticide applications (de la Poza et al. 2005; Ahmad et al. 2006; Marvier et al. 2007; Lu et al.

2012; Svobodová et al. 2017). Increasing the understanding of biological control services in commercial fields may provide complementary pest management tactics as biological control and Bt traits are generally regarded as compatible (Romeis et al. 2006).

Pitfall traps revealed a diverse community of epigeal carabids (Table 2.2). The most abundant species were A. sanctaecrucis, B. quadrimaculatum and E. anceps that collectively made up 66% (155/235) of all Carabidae described (Figure 2.3). Harpalus species were also quite abundant and constituted 15% (35/235) of the specimens found

(Figure 2.3). The Carabidae data reported herein add to our knowledge of ground beetles in agroecosystems throughout the state. Few studies have focused on investigating the community of ground beetles present in Nebraska. Unique to corn,

Hariharan (1988) found seven species in Clay County (south-central Nebraska) that were also found in our study: Agonum placidum (Say), B. quadrimaculatum, B. rapidum

(LeConte), E. anceps, Harpalus pensylvanicus (DeGeer), P. chalcites (syn. Pterostichus chalcites), and Poecilus lucublandus (Say). In addition to the seven species listed above, sugar beet plots in Scotts Bluff County (western Nebraska) revealed an additional six species that were also present in the current study: Anisodactylus carbonarius (Say),

Harpalus amputatus Say, Harpalus erraticus Say, Harpalus herbivagus Say, Harpalus somnulentus Dejean, and Stenolophus comma F. The present study together with 50

Hariharan (1988) and Pretorius et al. (2017) show that ground beetles have a wide geographical distribution in the state and are common members of agroecosystems.

In general, ground beetles have extremely diverse diets which can include insect and weed pests (Holland and Luff 2000). Contrary to our study, the species A. placidum,

B. quadrimaculatum, B. rapidum, H. pensylvanicus, P. chalcites and S. comma have all tested positive for WCR DNA in other studies (Lundgren et al. 2009a; Lundgren and

Fergen 2011, 2014). One explanation may be the smaller sample size of our study which had a total of 235 ground beetles analyzed in comparison to 432 in South Dakota

(Lundgren et al. 2009a). Similarly to our study, Lundgren et al. (2009a) also used pitfall traps to collect their predators but their traps were time-sorting pitfalls in which contents were segregated every 3h (Lundgren et al. 2009b). When comparing Lundgren and Fergen (2014) to our study we find methodology differences that may account for differences in results. Our study used natural levels of infestations in continuous corn, whereas they artificially infested first year corn. It is possible that artificially infested eggs were more readily available at the surface for those predators on the top soil layers. Moreover, Lundgren and Fergen (2014) used 10 cm soil cores for predator sampling, the same depth where the majority of WCR eggs are found (Gray et al. 1992), while our dry pitfalls selected only for surface-dwelling predators. Collecting predators throughout the soil core instead of focusing in the surface-dwellers probably increased the chances of finding predators positive for WCR DNA. However, dry pitfalls were the appropriate method to select for a high abundance of predators to be collected in a cost-efficient and timely manner. Due to time constraints, we decided to focus on 51

Carabidae taxa only because of their potential as WCR predators; however, the other surface-dwelling arthropod predators that were captured such as spiders are currently being analyzed for their potential as WBC and WCR predators.

Another explanation for the lack of WCR-DNA detection in carabids is the fact that only adult ground beetles were analyzed herein, while some of the beetles that consumed WCR such as Scarites quadriceps (not found in this study) did so in their larval stage (Lundgren et al. 2009a). It may also be possible that ground beetles collected were deterred by the sticky hemolymph defense that WCR larvae exhibit when facing predation (Lundgren et al. 2010). However, ground beetles likely also encountered eggs, pupae and adult beetles that do not have the sticky hemolymph defense, and therefore, this defense alone does not explain our negative results. Still, we should consider the possibility of lack of predation. For instance, WCR can be protected from predators while feeding within the roots as larvae (Strnad and Bergman 1987) or as pupae in their earthen pupal cell (Chiang 1973). Absence of WCR prey was expected in Field E (first year cornfield) where 9% of carabids the samples came from (Table 2.2), but it was expected that we would detect some predation in fields A and C, the high rootworm pressure fields (Figures 2.2. and 2.3). Western corn rootworms are also considered sub- optimal prey items for predators as its consumption is correlated with the increase in predator community abundance (Lundgren and Fergen 2014). Hence, it is possible that the abundance of Carabidae species in our fields is low and therefore predators are not being forced into eating non-preferred items. However, the most likely explanation is that there may have been an abundance of other, more preferred prey items or food 52 items available. As previously stated, carabid beetles have varied diets and many are polyphagous, opportunistic feeders (Holland and Luff 2000). Reviewing the most abundant species in our traps, we notice that A. sanctaecrucis is primarily a granivore although it does have a polyphagous lifestyle (Hagley et al. 1982; Lundgren and

Rosentrater 2007), and E. anceps and B. quadrimaculatum are predators, but the latter feeds on plant tissue as well (Brousseau et al. 2018, Fox et al. 2005, Kamenova et al.

2015). Anisodactylus sanctaecrucis density was primarily driven from Field B (Table 2.3) as 63 out of 65 specimens were found in that site. Field B had a high density of weeds across the field, so it is possible that weed seeds supported A. sanctaecrucis densities.

The different feeding habits of these main species reinforces the notion that

WCR are not their primary prey choice. Kirk (1982) anecdotal observations suggest carabids prey on rootworms only on chance encounters. This agrees with the other papers that find a small percentage of Carabidae specimens testing positive for WCR

DNA (Lundgren et al. 2009a, Lundgren and Fergen 2014). The negative results from the present study along with other studies support the conclusion that carabids uncommonly use WCR life stages as food in cornfields. This study is the first of this kind in the state of Nebraska and is the first effort to better understand the trophic interactions between WCR and carabids in this region. Future studies should focus on other soil predators with a focus on understanding egg predation as this life stage is present in the field soil for the majority of the year. Documentation of the availability and efficacy of native WCR predators will aid in the development of sound conservation biological control programs in the state of Nebraska. 53

The trophic web of WCR is still poorly understood. Many predator-prey reports are laboratory experiments, including no-choice experiments or are weak trophic interactions based on MCGA or personal observations. There is a need to describe natural enemies that are effective mortality agents of the WCR that can be used for biological control and IPM programs. Entomopathogens, primarily fungi and nematodes, have shown great promise for WCR control and will be the topic of the next chapters.

54

References

Archibald, W. R. 2017. Conservation biological control of western bean cutworm: Molecular gut content analysis of arthropod predators, feeding trials for key predators and agricultural surveys for integrated pest management. M.S. Thesis. University of Nebraska – Lincoln.

Arnett, R. H. JR. and Thomas, M. C. 2000. American Beetles. Archostemata, Myxophaga, Adephaga, Polyphaga: Staphiliniformia. CRC Press LLC. Boca Raton, FL. 1: 32-132.

Ahmad, A., G. E. Wilde, R. J. Whitworth, and G. Zolnerowich. 2006. Effect of corn hybrids expressing the coleopteran-specific Cry3Bb1 protein for corn rootworm control on aboveground insect predators. J. Econ. Entomol. 99: 1085–1095.

Brousseau, P-M., Gravel, D., Handa, I.T., 2018. Trait matching and phylogeny as predictors of predator–prey interactions involving ground beetles. Functional Ecol. 32, 192-202. DOI: 10.1111/1365-2435.12943

Chiang, H. C. 1973. Bionomics of the northern and western corn rootworms. Annu. Rev. Entomol. 18: 47–72.

Clark, T. L., and B. E. Hibbard. 2004. Comparison of nonmaize hosts to support western corn rootworm (Coleoptera: Chrysomelidae) larval biology. Environ. Entomol. 33: 681–689. de la Poza, M., Pons, X., Farinós, G.P., López, C., Ortego, F., Eizaguirre, M., Castañera, P., Albajes, R., 2005. Impact of farm-scale Bt maize on abundance of predatory arthropods in Spain. Crop Protec. 24, 677–684. https://doi.org/10.1016/j.cropro.2004.12.003

Duan, J.J., Paradise, M.S., Lundgren, J.G., Bookout, J.T., Jiang, C., Wiedenmann, R.N., 2006. Assessing nontarget impacts of Bt corn resistant to corn rootworms: Tier-1 testing with larvae of Poecilus chalcites (Coleoptera: Carabidae). Environ. Entomol. 35, 135–142. https://doi.org/10.1603/0046-225X-35.1.135

Eskelson, M.J., Chapman, E.G., Archbold, D.D., Obrycki, J.J., Hardwood, J.D. 2011. Molecular identification of predation by carabid beetles on exotic and native slugs in a strawberry agroecosystem. Bio. Control. 56: 245-253.

Folmer, O., Black, M., Hoeh, W., Lutz, R., Vrijenhoek, R., 1994. DNA primers for amplification of mitochondrial cytochrome c oxidase subunit I from diverse metazoan invertebrates. Mol. Mar. Biol. and Biotech. 3(5), 294-299. 55

Fox, T.B., Landis, D.A., Cardoso, F.F., DiFonzo, C.D., 2005. Impact of predation on establishment of the soybean aphid, Aphis glycines in soybean, Glycine max. BioControl 50: 545-563.

Gassmann, A. J., J. L. Petzold-Maxwell, E. H. Clifton, M. W. Dunbar, A. M. Hoffmann, D. A. Ingber, and R. S. Keweshan. 2014. Field-evolved resistance by western corn rootworm to multiple Bacillus thuringiensis toxins in transgenic maize. Proc. Natl. Acad. Sci. 111: 5141–5146.

Gassmann, A. J., J. L. Petzold-Maxwell, R. S. Keweshan, and M. W. Dunbar. 2011. Field- Evolved resistance to Bt maize by western corn rootworm. PLoS ONE. 6: e22629.

Gassmann, A. J., R. B. Shrestha, S. R. K. Jakka, M. W. Dunbar, E. H. Clifton, A. R. Paolino, D. A. Ingber, B. W. French, K. E. Masloski, J. W. Dounda, and C. R. S. Clair. 2016. Evidence of resistance to Cry34/35Ab1 corn by western corn rootworm (Coleoptera: Chrysomelidae): root injury in the field and larval survival in plant-based bioassays. J. Econ. Entomol. 109(4): 1872-1880.

Gray, M. E., G. L. Hein, M. A. Boetel, and D. D. Walgenbach. 1992. Western and northern corn rootworm (Coleoptera: Chrysomelidae) egg densities at three soil depths: implications for future ecological studies. J. Kans. Entomol. Soc. 354– 356.

Gray, M. E., T. W. Sappington, N. J. Miller, J. Moeser, and M. O. Bohn. 2009. Adaptation and invasiveness of western corn rootworm: intensifying research on a worsening pest. Annu. Rev. Entomol. 54: 303–321.

Hariharan, M.J. 1988. Effect of chlorpyrifos on selected ground-dwelling beetles (Carabidae) in a cornfield. M.S. Thesis. University of Nebraska-Lincoln.

Hagley, E. a. C., N. J. Holliday, and D. R. Barber. 1982. Laboratory studies of the food preferences of some orchard carabids (Coleoptera: Carabidae). Can. Entomol. 114: 431–437.

Hein, G. L., and J. J. Tollefson. 1985. Use of the Pherocon AM trap as a scouting tool for predicting damage by corn rootworm (Coleoptera: Chrysomelidae) larvae. J. Econ. Entomol. 78: 200–203.

Hitchon, A. J., J. L. Smith, B. W. French, and A. W. Schaafsma. 2015. Impact of the Bt corn proteins Cry34/35Ab1 and Cry3Bb1, alone or pyramided, on western corn rootworm (Coleoptera: Chrysomelidae) beetle emergence in the field. J. Econ. Entomol. 108: 1986–1993.

Holland, J. M., and M. L. Luff. 2000. The effects of agricultural practices on carabidae in temperate agroecosystems. Integr. Pest Manag. Rev. 5: 109–129. 56

Kamenova, S., K. Tougeron, M. Cateine, A. Marie, and M. Plantegenest. 2015. Behaviour-driven micro-scale niche differentiation in carabid beetles. Entomol. Exp. Appl. 155: 39–46.

Kirk, V. M. 1982. Carabids: minimal role in pest management of corn rootworms. Environ. Entomol. 11: 5–8.

Kuhlmann, U., and W. A. C. M. Van der Burgt. 1998. Possibilities for biological control of the western corn rootworm, Diabrotica virgifera virgifera LeConte, in central Europe. Biocontrol News Inf. 19: 59N–68N.

Levine, E., J. L. Spencer, S. A. Isard, D. W. Onstad, and M. E. Gray. 2002. Adaptation of the western corn rootworm to crop rotation: evolution of a new strain in response to a management practice. Am. Entomol. 48: 94–107.

Lindroth, C.H., 1961. The ground-beetles (Carabidae, excluding Cicindelinae) of Canada and Alaska, parts 1-6. Opuscula Entomologica Supplementa XX, XXIV, XXIX, XXXIII, XXXV.

Lu, Y., K. Wu, Y. Jiang, Y. Guo, and N. Desneux. 2012. Widespread adoption of Bt cotton and insecticide decrease promotes biocontrol services. Nature. 487: 362–365.

Lundgren, J. G., M. E. Ellsbury, and D. A. Prischmann. 2009a. Analysis of the predator community of a subterranean herbivorous insect based on polymerase chain reaction. Ecol. Appl. 19: 2157–2166.

Lundgren, J. G., and J. K. Fergen. 2011. DNA primers for amplification of mitochondrial cytochrome c oxidase subunit I from diverse metazoan invertebrates. Mol. Mar. Biol. and Biotech. 3(5): 294-299.

Lundgren, J. G., and J. K. Fergen. 2014. Predator community structure and trophic linkage strength to a focal prey. Mol. Ecol. 23: 3790–3798.

Lundgren, J. G., S. Nichols, D. A. Prischmann, and M. M. Ellsbury. 2009b. Seasonal and diel activity patterns of generalist predators associated with Diabrotica virgifera immatures (Coleoptera: Chrysomelidae). Biocontrol Sci. Technol. 19: 327–333.

Lundgren, J. G., and K. A. Rosentrater. 2007. The strength of seeds and their destruction by granivorous insects. Arthropod-Plant Interact. 1: 93–99.

Lundgren, J. G., S. Toepfer, T. Haye, and U. Kuhlmann. 2010. Haemolymph defence of an invasive herbivore: its breadth of effectiveness against predators. J. Appl. Entomol. 134: 439–448.

Marvier, M., C. McCreedy, J. Regetz, and P. Kareiva. 2007. A meta-analysis of effects of Bt cotton and maize on nontarget invertebrates. Science. 316: 1475–1477. 57

Meinke, L. J., B. D. Siegfried, R. J. Wright, and L. D. Chandler. 1998. Adult susceptibility of Nebraska western corn rootworm (Coleoptera: Chrysomelidae) populations to selected insecticides. J. Econ. Entomol. 91: 594–600.

Miller, N. J., T. Guillemaud, R. Giordano, B. D. Siegfried, M. E. Gray, L. J. Meinke, and T. W. Sappington. 2009. Genes, gene flow and adaptation of Diabrotica virgifera virgifera. Agric. For. Entomol. 11: 47–60.

Parimi, S., L. J. Meinke, B. Wade French, L. D. Chandler, and B. D. Siegfried. 2006. Stability and persistence of aldrin and methyl-parathion resistance in western corn rootworm populations (Coleoptera: Chrysomelidae). Crop Prot. 25: 269– 274.

Pereira, A. E., H. Wang, S. N. Zukoff, L. J. Meinke, B. W. French, and B. D. Siegfried. 2015. Evidence of field-evolved resistance to bifenthrin in western corn rootworm (Diabrotica virgifera virgifera LeConte) populations in western Nebraska and Kansas. PLoS ONE. 10: e0142299.

Peterson, J. A. 2012. Delineating the influence of genetically modified crops and non- prey food resources on generalist predator food webs. Ph.D. Dissertation. University of Kentucky.

Pierce, C. M. F., and M. E. Gray. 2007. Population dynamics of a western corn rootworm (Coleoptera: Chrysomelidae) variant in east central Illinois commercial maize and soybean fields. J. Econ. Entomol. 100: 1104–1115.

Pretorius, R.J., Hein, G.L., Blankenship, E.E., Purrington, F.F., Bradshaw, J.D., 2017. Response of Pemphigus betae (Hemiptera: Aphididae) and beneficial epigeal arthropod communities to sugarbeet plant density and seed-applied insecticide in western Nebraska. Environ. Entomol. 46: 107–117. https://doi.org/10.1093/ee/nvw157 de la Poza, M., X. Pons, G. P. Farinós, C. López, F. Ortego, M. Eizaguirre, P. Castañera, and R. Albajes. 2005. Impact of farm-scale Bt maize on abundance of predatory arthropods in Spain. Crop Prot. 24: 677–684.

Romeis, J., M. Meissle, and F. Bigler. 2006. Transgenic crops expressing Bacillus thuringiensis toxins and biological control. Nat. Biotechnol. 24: 63–71.

Strnad, S. P., and M. K. Bergman. 1987. Distribution and orientation of western corn rootworm (Coleoptera: Chrysomelidae) larvae in corn roots. Environ. Entomol. 16: 1193–1198.

Svobodová, Z., Y. Shu, O. S. Habuštová, J. Romeis, and M. Meissle. 2017. Stacked Bt maize and arthropod predators: exposure to insecticidal Cry proteins and potential hazards. Proc R Soc B. 284: 20170440. 58

Symondson, W. O. C., Sunderland, K. D. and M. H. Greenstone. 2002. Can generalist predators be effective biocontrol agents? Annu. Rev. Entomol. 47: 561–594.

Toepfer, S., T. Haye, M. Erlandson, M. Goettel, J. G. Lundgren, R. G. Kleespies, D. C. Weber, G. C. Walsh, A. Peters, R.-U. Ehlers, H. Strasser, D. Moore, S. Keller, S. Vidal, and U. Kuhlmann. 2009. A review of the natural enemies of beetles in the subtribe Diabroticina (Coleoptera: Chrysomelidae): implications for sustainable pest management. Biocontrol Sci. Technol. 19: 1–65.

Tripathi, R. C. 2006. Negative binomial distribution. Encycl. Stat. Sci., Major Reference Works.

USDA-NASS. Crop Production 2017 Summary. 2017. https://www.nass.usda.gov/Publications/Todays_Reports/reports/crop1117.pdf Accessed 4.April.2018

Wangila D.S., Gassmann A.J., Petzold-Maxwell J.L., French B.W., Meinke L.J. 2015. Susceptibility of Nebraska western corn rootworm (Coleoptera: Chrysomelidae) populations to Bt corn events. J Econ Entomol 108:742–751.

Zukoff, S. N., K. R. Ostlie, B. Potter, L. N. Meihls, A. L. Zukoff, L. French, M. R. Ellersieck, B. W. French, and B. E. Hibbard. 2016. Multiple assays indicate varying levels of cross resistance in Cry3Bb1-selected field populations of the western corn rootworm to mCry3A, eCry3.1Ab, and Cry34/35Ab1. J. Econ. Entomol. 109(3): 1387-1398. 59

Tables

Table 2.1. Total abundance of predator taxa found on yellow sticky cards per field.

Field Total Identity A B C D E Arachnida Araneae 31 13 7 13 23 87 Insecta Coleoptera 9 15 20 6 8 58 Carabidae 1 1 0 0 1 3 Staphylinidae 15 13 8 11 12 59 Coccinellidae 1 1 0 1 0 3 Coleomegilla maculata 7 5 3 4 1 20 Hippodamia convergens 15 3 2 6 5 31 Coccinella septempunctata 2 1 0 3 7 13 Hemiptera Anthocoridae Orius insidiosus 55 13 15 2 5 90 Geocoridae Geocoris spp. 2 1 0 1 3 7 Hymenoptera Formicidae 4 18 0 86 20 128 Neuroptera Chrysopidae 90 67 55 89 48 349 Hemerobiidae 0 1 4 2 1 8 Others 2 1 5 1 1 10 Total per Field 234 153 119 225 135 866

60

Table 2.2. Carabidae species found in dry pitfalls per field.

Field Species A B C D E Total Agonum extensicolle (Say) 1 0 0 0 0 1 Agonum lutulentum Leconte 0 0 0 3 0 3 Agonum placidum (Say) 1 0 0 1 0 2 Agonum sp. Bonelli 0 2 0 2 0 4 Anisodactylus carbonarius (Say) 0 2 0 0 0 2 Anisodactylus laetus Dejean 1 1 0 0 0 2 Anisodactylus merula (Germar) 0 1 0 0 0 1 Anisodactylus sanctaecrucis (Fabricius) 0 63 1 1 0 65 Athrostictus punctatulus (Putzeys) 1 2 0 0 0 3 Bembidion constrictum LeConte 1 0 0 0 0 1 Bembidion mundum (LeConte) 0 0 0 0 1 1 Bembidion quadrimaculatum Say 6 2 23 5 3 39 Bembidion rapidum (LeConte) 0 0 0 1 1 2 Calathus opaculus LeConte 0 0 0 2 0 2 Callisthenes affinis Chaudoir 0 0 0 1 0 1 Carabus serratus Say 0 0 1 1 0 2 Chlaenius tricolor Dejean 1 0 0 0 0 1 Cicindela punctulata Olivier 0 0 0 0 4 4 Elaphropus anceps LeConte 5 7 12 20 7 51 Harpalus affinis (Schrank) 1 4 0 0 1 6 Harpalus amputatus Say 2 0 0 0 0 2 Harpalus compar LeConte 0 0 0 0 1 1 Harpalus erraticus Say 0 1 0 0 0 1 Harpalus erythropus Dejean 2 0 0 0 0 2 Harpalus faunus Say 0 1 0 0 0 1 Harpalus herbivagus Say 0 3 1 4 0 8 Harpalus pensylvanicus (DeGeer) 0 6 0 0 1 7 Harpalus seclusus Casey 1 2 0 0 0 3 Harpalus somnulentus Dejean 1 2 0 0 1 4 Lachnocrepis parallela (Say) 1 1 0 0 0 2 Lebia viridis Say 0 0 0 1 0 1 Poecilus chalcites (Say) 0 0 1 0 0 1 Poecilus lucublandus (Say) 0 1 0 0 0 1 Stenolophus comma (F.) 0 3 1 1 0 5 Stenolophus fuliginosus Dejean 1 0 0 0 0 1 Tachys scitulus (LeConte) 0 0 0 0 2 2 Total 26 104 40 43 22 235 Percentage 11.1% 44.2% 17% 18.3% 9.4% 61

Figures

60 A 50 a B C 40 D a E 30

20 b 10 Mean cumulative WCR per cage WCR Mean cumulative c d 0 15 22 29 7 11 19 26 2 9 16 26 July August September Sampling Date

Figure 2.1. Mean cumulative western corn rootworm (WCR) adults per emergence cage. Upper-case letters represent cornfields A-E. One-way ANOVA from cumulative values on September.26, revealed significant field effects (F4, 35 = 20.63, Pr > F < 0.0001) on mean cumulative WCR beetles per emergence cage (n = 8). Means with different lower-case letters are significantly different (P <0.05). 62

a

a

b b c

Figure 2.2. Mean cumulative WCR adults per yellow sticky card. Upper-case letters represent cornfields A-E. One-way ANOVA from cumulative counts at September.9 revealed significant field effects (F4,15 = 45.29, Pr > F < 0.0001) on mean WCR beetles per yellow sticky card (n = 4). Means with different lower-case letters are significantly different (P <0.05).

63

16.6% Anisodactylus sanctaecrucis 27.7% Elaphropus anceps Others 14.8% Harpalus spp. Bembidion quadrimaculatum

21.7% 19.2%

Figure 2.3. Most common Carabidae taxa in dry pitfalls.

64

CHAPTER 3: CHARACTERIZATION OF POTENTIAL FUNGAL ENTOMOPATHOGENS FROM COMMERCIAL, IRRIGATED, CONTINUOUS CORNFIELDS.

Introduction

The United States is the leading producer of corn (Zea mays L.) in the world

(USDA-FAS, 2017). Corn production in the U.S. Corn Belt is a high input system, and in some areas, corn is produced as corn-on-corn in a continuous production system involving growing the crop for two or more consecutive years. Continuous crop production can lead to increased disease and insect pressure in the following crop seasons (Tilman et al. 2002). In many areas of the U.S. Corn Belt, the key soil pest of continuous corn is the western corn rootworm (WCR), Diabrotica virgifera virgifera

LeConte (Chrysomelidae: Coleoptera) (Gray et al. 2009). The soil also hosts secondary pests that feed on corn seedlings such as white grubs (Phyllophaga spp., Cyclocephala spp., and Popillia japonica Newman; Scarabaeidae: Coleoptera) and wireworms

(Melanotus spp., Agriotes spp., and Limonius dubitans LeConte; Elateridae: Coleoptera).

Soil pests are very difficult to control because their cryptic feeding and associated damage is difficult to predict (Jackson et al. 2000). Common options for control of soil pests include seed treatments and soil insecticides, however there are often efficacy inconsistencies and environmental issues associated with those methods (Harris 1972;

Tooker et al. 2017).

The soil is an important source of natural enemies of insects (Klingen and

Haukeland 2006). Fungi that are associated with insects are called entomogenous fungi

(EF) and can have important relationships with insects such as commensalism,

65 parasitism and pathogenesis (Roberts and Humber 1981). The last relationship is of utmost importance for insect pest management. There are many entomopathogenic fungi (EPF) species that have been described and commercialized as biopesticides

(Humber 2000; Lacey et al. 2015). Entomopathogenic fungi have evolved complex and diverse life histories (Wang and Wang 2017). Estimates suggest there are 750-1,000 EPF species in 100 genera (Vega et al. 2012). EPF are present in many high fungal taxa but

Entomophthoromycota (Entomophthorales) and Ascomycota (Hypocreales) hold the majority of species (Humber 2016). Entomophthorales are responsible for many large- scale epizootics but many species have limited host ranges and are difficult to cultivate in vitro because they are obligate pathogens (Roy et al. 2006). On the other hand,

Hypocreales have large host ranges, are easily produced in vitro and are highly explored as biological control agents (BCAs) (Shah and Pell 2003; Vega et al. 2012).

Knowledge of native strains of EPF in the soil can lead to the discovery of novel

BCAs and is also a necessary first step in the development of conservation and classical biological control programs (Hajek et al. 2000; Meyling and Eilenberg 2007; Solter et al.

2017). The hypocrealeans Beauveria bassiana (Balsamo) Vuillemin, Metarhizium anisopliae (Metschn.) Sorokin, Lecanicillium and Isaria are commonly produced as commercial strains worldwide (Shah and Pell 2003; de Faria and Wraight 2007; Humber

2016). Entomopathogenic fungi have many environmental benefits in comparison to chemical control such as safety to non-target organisms, including humans, and reduced pesticide residue (Lacey et al. 2001). However, they also possess drawbacks like low persistence, slow killing speed, and higher cost (Humber 2016, Lacey et al. 2001). In

66 combination with other pest management practices, entomopathogens have the potential to reduce pest population densities including a potential role in delaying or mitigating resistance evolution under certain conditions (Shah and Pell 2003; Meissle et al. 2009; Hannon et al. 2010; Petzold-Maxwell et al. 2012; Gassmann et al. 2012;

Hoffmann et al. 2014). Studying the EPF community in continuous, irrigated cornfields can help identify BCA’s that are adapted to this system and that can be further explored for a wide range of insect pests. Therefore, the objectives of this study were to (i) document the biodiversity of insect-associated fungal pathogens from the corn rhizosphere in irrigated cornfields of western Nebraska, and (ii) identify entomopathogens to be further explored as pest management tools.

Materials and Methods

Study Sites. All five study sites were commercial, irrigated, no-till cornfields located in

West Central Nebraska. Details on western corn rootworm (WCR) density, location, soil type, rotational history, hybrids, transgenic traits, planting date, and insecticide and fungicide use are listed in Appendix 1 and Fig. 3.1a. All fields were maintained under standard agronomic practices for the region, including fertilization, irrigation, and weed management.

Soil Sampling. In 2014, soil sampling from each of the five sites was conducted on seven dates between June and September. In each field, a total of ten randomly chosen soil samples were taken: eight within the irrigated area and two from dryland corners adjacent to the irrigated field. Randomization was performed using a random number generator with a range of 8 to 15. Then along the pivot tire track (Fig. 3.1b) samples

67 were taken 8 to 15 meters apart. In 2015, soil sampling from each of the five sites was conducted on five dates between June and August. In each field, a total of eight soil samples were taken from fixed locations that were randomly assigned at the beginning of the season within the irrigated area with no dryland corner sampling (Fig. 3.1b). Soil samples were taken with a hand trowel from the corn-root zone to a depth of approximately 10 cm and 946 ml volume. Tools were sterilized with 90% Ethanol and then flamed between samples.

Baiting Assays. In the laboratory, each soil sample was homogenized manually and approximately 200 ml of soil was dispensed into a 13.6 X 11.4 X 5.1 cm (LxWxH) clear plastic dish (Dartâ, Mason, Michigan, USA). In 2014, five Galleria mellonella L.

(Lepidoptera: Pyralidae) larvae were added to each container (Zimmermann 1986). In an attempt to isolate a higher diversity of entomopathogens and to isolate for Coleoptera- specific pathogens, in 2015, Tenebrio molitor L. (Coleoptera: Tenebrionidae) larvae were also utilized in baiting assays (Pilz et al. 2008). Soil samples were divided into two dishes with approximately 200 ml of soil in each; one received three G. mellonella and the other received three T. molitor larvae. Both G. mellonella and T. molitor were obtained from Speedy Worm, Alexandria, MN. Baiting dishes were kept at 20-22°C in the dark and soil was kept moist with double distilled millipore (ddH2O) water throughout the duration of the baiting period. Larval mortality was assessed after 5, 7 and 10 days.

Cadavers were surface-sterilized with 70% ethanol, ddH2O, and 1% sodium hypochlorite then blot-dried on filter paper (Lacey and Solter 2012). Cadavers were then placed into humid chambers made with petri dishes (Fisherbrand™, Pittsburgh, PA) and moist filter

68 paper (Fisherbrand™) to allow for sporulation. Condition of sporulation was checked 3-5 days after larvae were placed in humid chambers. Infection status was recorded and specimens that were not consumed by saprophytic fungi were then stored at 4°C for fungal isolation and identification procedures. Larval cadavers with abundant sporulation were not surface sterilized but were saved for fungal isolation procedures.

Isolation and Identification of Fungi. Fungal material from G. mellonella and T. molitor cadavers were transferred to agar plates via direct transfer of spores with sterile wooden toothpicks or by transferring infected small larval pieces (approximately 2 to 3 mm2) that were cut with sterile scalpels. Cadaver pieces were rinsed with distilled water to remove any organic debris, dipped in 95% ethanol for 5 s, rinsed again in sterile distilled water, and allowed to drain on a sterile paper towel. The fungal material were placed on CTC medium containing potato dextrose agar and yeast extract supplemented with chloramphenicol, thiabendazole and cycloheximide (2014 only) and potato dextrose agar amended with 0.01% tetracycline (PDAt) (2014 and 2015) (Fernandes et al. 2010; Adesemoye et al. 2014). In 2015, CTC medium was not used because it was determined in 2014 that PDAt allowed for a greater diversity of potential entomopathogens to grow. Inoculated culture plates were incubated at 25°C and serially transferred until pure cultures were obtained. Fungal isolates were allowed to sporulate for morphological identification using a high-resolution compound microscope mounted with a single-lens reflex (SLR) camera.

Genomic DNA from fungal isolates was extracted using DNeasyâ Plant Mini Kit

(QIAGENã Inc., Chatsworth, CA, USA). Polymerase chain reaction (PCR) was conducted

69 using internal transcribed spacer (ITS), ITS4 and ITS5 (White et al. 1990) for the ITS1,

ITS2, and 5.8S regions of ribosomal DNA. Additionally, Bt2a and Bt2b (Glass and

Donaldson 1995) was used for the β-tubulin gene. Each PCR reaction contained 12.5 μl of GoTaq Green Master Mix (Promega Corp., Fitchburg, WI, USA), 9.3 μl of PCR-grade water, 0.6 μl of 10 μM each primer, and 2 μl of DNA template. The PCR reaction protocol for both primer sets included an initial preheat at 94°C for 2 min; 35 cycles of denaturation at 94°C for 15 s, annealing at 58°C for 15 s, and extension at 72°C for 45 s; and final extension at 72°C for 5 min (Adesemoye et al. 2014). PCR products were verified via gel electrophoresis in 1% agarose gels in 1× Tris-acetate-EDTA (TAE) buffer and photographed with GelDocä XR+ Gel Documentation system. PCR products were purified using QIAquick PCR Purification kit (QIAGENã Inc.) and quality-checked with

NanoVue Plus UV/Visible Spectrophotometer (GE Healthcare Life Sciences,

Marlborough, MA, USA). Sequencing was done at the University of California, Riverside

Gencore. Sequences were edited with MEGA 7 and analyzed with DNASTAR Lasergene version 14 software (DNASTAR, Madison, WI, USA) and BLAST search was done on the

National Center for Biotechnology Information, United States National Institutes of

Health (NCBI) website and sequences were submitted to GenBank.

Data Analysis. All statistical analyses were performed using generalized linear mixed models with PROC GLIMMIX (SAS software SAS, v. 9.4; SAS Institute Inc.). All data were analyzed as proportion data fitted to a beta-binomial distribution prior to analyses

(Ferrari and Cribari-Neto 2004; Stroup 2015). Two-way ANOVAs of main effects of field and sampling date and their interaction were analyzed for 2014 and 2015. One-way

70

ANOVA with PROC GLIMMIX was run to determine the effect of insect species (G. mellonella and T. molitor) on the mean proportion of entomogenous fungi detection across fields and dates. The same procedure was repeated specifying Metarhizium spp. detection. The LSMEANS function was used with ILINK option to convert mean estimates, standard errors and confidence limits to the data scale prior to fitting beta- distribution (Schabenberger 2005). Simple effects and interactions at P< 0.05 were considered significant.

Results

Frequency of Detection of Entomogenous Fungi. Entomogenous fungi were detected in

40.6% (138/340) of all soil samples in 2014 and in 34.0% (67/197) in 2015. In 2014,

15.24% (250/1640) of G. mellonella larvae were infected. Mean percentage of cadavers infected per arena in 2014 had a significant field by date interaction (F23, 306 = 1.61, Pr > F

= 0.0394). Least square means comparisons showed varying levels of mean cadaver infection per each interaction of field by date (Fig. 3.2). Field B had higher mean cadavers infected on Sept.3 when compared to all other sampling dates, except June.24 and July.22. All other fields did not show any significant seasonality pattern in mean cadaver infection (Fig 3.2). In 2015, 10.26% of all G. mellonella and T. molitor larvae

(121/1179) were infected. In 2015, there was no significant effect of insect species used for baiting on frequency of fungal detection: 10.7% (63/588) of G. mellonella and 9.81%

(58/591) of T. molitor were infected (F1,48 = 0.46, Pr > F = 0.50). There was no significant effect of field by date interaction (F16, 175 = 0.32, Pr > F=0.9944) or sampling date effect

(F4, 175 = 1.13, Pr > F = 0.3462) on mean cadaver infection per arena (Fig. 3.3). However,

71

there was an impact of field (F4, 175 =4.43, Pr > F = 0.0020) on cadaver infection, with

Field B being significantly higher than the others (Fig. 3.3).

Fungal Identification. In total, 254 pure fungal cultures were obtained from infected cadavers in 2014 and 119 in 2015. From the pure cultures, 132 samples were selected for molecular identification: 64 (25.6% of all infected cadavers) from 2014, which included 18 samples from the non-irrigated borders, and 68 (56.2%) from 2015. Isolate selection was made to represent all morphological groups in the collection.

The isolated fungi were identified as belonging to 11 families, 14 genera and 29 species (Table 3.1). The most diverse family was Trichomaceae: Eurotiales with four genera and nine species, followed by Nectriceae: Hypocreales with four Fusarium species, then Myxotrichaceae with three Pseudogymnoascus species. Chaetomiaceae:

Sordariales, Cladosporiceae: Capnodiales, and Hypocreaceae: Hypocreales all had two species in each family. Clavicipitaceae: Hypocreales was represented by the genus

Metarhizium. Bionectriaceae, Cordycipitaceae, and Ophiocordycipitaceae (all

Hypocreales) were all represented by one species.

Metarhizium was the most prevalent genus in both years with 45.3% (29/64) detection of the identified fungi in 2014 and 69.1% (47/68) in 2015. Three species were identified; sixty-six isolates of M. robertsii J.F. Bisch., Rehner & Humber, seven of M. anisopliae and three remained under Metarhizium spp. In 2015, Metarhizium spp. detection was not statistically different due to insect species used for baiting: 63.8%

(30/47) of G. mellonella and 48.9% (23/47) T. molitor were positive (F1,48 = 0.66, Pr > F =

72

0.42). However, some species were identified uniquely in T. molitor or G. mellonella

(Table 3.1). For instance: B. bassiana, Penicillium spp. and Trichoderma spp. were only baited from G. mellonella, while Clonostachys spp. were only baited from T. molitor. The genus Penicillium was identified down to the greatest number of species (n=4); followed by Pseudogymnoascus, Talaromyces, (both with 3 species); Aspergillus, Cladosporium,

Trichoderma (all with 2 species); and Beauveria, Chaetomium, Clonostachys,

Geotrichum, Neosartorya, Purpureocillium, Taifanglania all with one species each.

Sequences obtained for the ITS and BT regions were deposited in the GenBank database

(Table 3.1).

Fungal Ecological Classification. The concept of classifying baited fungi into ecological functions (Table 3.2) was developed from Sun et al. (2008) and Oliveira et al. (2011).

While all strains were recovered from insect cadavers, it doesn’t mean they were the causal agents of mortality. Therefore, classification of species was based on available literature, with many species expressing multiple ecological roles. Species described at the genus level were classified on available reports of one or more species.

Entomopathogenic status was confirmed for 62.1% (82/132) of strains with Metarhizium spp., B. bassiana and P. lilacinum being the species most commonly described as insect pathogens. Antagonists of plant pathogens were found in 7.6% (10/132) that may be explored for biological control of plant diseases. Antagonists included Clonostachys sp.,

Chaetomium sp., Trichoderma gamsii, T. virens and P. lilacinum. Strains classified as entomogenous/insect antagonists belong to species or genera that have been previously reported from insects or that kill insects via toxins or endophytic properties.

73

Those placed under this category make up 10.6% (14/132) and are Clonostachys sp.,

Chaetomium sp., Fusarium acuminatum, F. oxysporum, F. solani, and A. flavus. Potential plant pathogens are those that have been reported with features harmful to plants and were found in 17.6% (24/132) of identified samples (i.e., Chaetomium sp., Cladosporium spp., Fusarium spp., Aspergillus spp. and Penicillium spp). Species classified under

“Others” made up 25.7% (34/132) of samples and included Taifanglania sp.,

Cladosporium spp., Geotrichum candidum, Pseudogymnoascus spp., Aspergillus spp.,

Neosartorya sp., Penicillium spp., and Talaromyces spp. These fungi exhibit saprophytic and/or opportunistic features.

Discussion

The present work is the first to characterize entomogenous fungi and identify potential EPF from commercial cornfields in Nebraska. Around the world many studies have evaluated the distribution and abundance of EPF species with insect baits in a variety of cropping systems (Vänninen 1996; Chandler et al. 1997; Ali-Shtayeh et al.

2002; Meyling and Eilenberg 2006a; Quesada-Moraga et al. 2007; Sun et al. 2008;

Oliveira et al. 2011; Wakil et al. 2013; Rudeen et al. 2013; Clifton et al. 2015). Soil baiting assay studies are often restricted to searches for EPF genera such as Beauveria,

Metarhizium and Purpureocillium (Paecilomyces) because of their potential as BCAs.

However, we found a wide range of fungi in addition to these known entomopathogens that can be isolated with the insect baiting assay method. The current study is similar to studies from Portugal (Oliveira et al. 2011) and China (Sun and Liu 2008; Sun et al. 2008)

74 that found secondary colonizers, opportunistic fungi, plant pathogens, and some insect and phytopathogenic antagonistic species from insect cadavers.

There was a significant interaction between field and sampling date (Fig. 3.2) on the mean cadaver infection in 2014 and a field effect in 2015 (Fig. 3.3). This variation between fields and sampling dates is common in agroecosystems and has been reported by other authors (Chandler et al. 1997; Meyling and Eilenberg 2006a; Quesada-Moraga et al. 2007; Pilz et al. 2008; Rudeen et al. 2013; Clifton et al. 2015). Fungal populations are influenced by a variety of dynamic interactions between abiotic (e.g. pH, soil properties, organic matter, temperature, moisture, agronomical practices) and biotic

(e.g. resource/host availability, environmental persistence, competition, natural enemies) conditions that can influence their distribution in an area (Quesada-Moraga et al. 2007; Wraight et al. 2007; Meyling and Hajek 2010). For instance, Field B was a particularly weedy field in both years. Weeds can have positive interactions with soil fungi that enables them to compete with crop plants in agroecosystems (Massenssini et al. 2014) but also can have negative interactions with fungi responsible for seed-decay

(Müller-Stöver et al. 2016). Field B contained 60% (6/10) of Fusarium spp. isolates (plant pathogens) but also contained 100% (n=3) of the Trichoderma spp. isolates (antagonist of phytopathogens) (Table 3.1). The impact of biotic and abiotic factors on fungal frequency should be studied in the context of creating pest management recommendations for this system.

Insect host species (G. mellonella or T. molitor) used in baiting assays did not impact the abundance of the total EF community or Metarhizium strains isolated during

75

2015. However, Clonostachys spp. were associated only with T. molitor; and B. bassiana,

Penicillium spp. and Trichoderma spp. only with G. mellonella (Table 3.1). The effect of insect species on the detection of EPF has varied between published studies. Pilz et al.

(2008) found significantly more T. molitor infected with EPF (16.6%) than G. mellonella

(1.7%), but Rudeen et al. (2013) found more EPF-related mortality on G. mellonella, followed by T. molitor and then D. v. virgifera. Moreover, Tolypocladium cylindrosporum

W. Gams (Hypocreales: Ophiocordycipitaceae) was isolated only from Delia floralis

(Fallen) (Diptera: Anthomyiidae), but G. mellonella was better at isolating M. anisopliae and B. bassiana (Klingen et al. 2002). As demonstrated by these studies, the use of multiple insect hosts for fungal baiting can lead to detection of a more diverse community of fungi and therefore can help in the search for taxon-specific strains that can be explored as BCAs against different insect groups.

Two of the most commonly reported genera of EPF (Beauveria and Metarhizium) were found in this study, even though the frequency of occurrence was highly variable

(Table 3.1). Beauveria bassiana and Metarhizium have a cosmopolitan distribution

(Humber 2000; Roberts and St Leger 2004) and have been isolated from cornfields before (Pilz et al. 2008; Rudeen et al. 2013; Clifton et al. 2015). Metarhizium was the dominant genus in our survey with 76 identified isolates from all five field sites in both years; while B. bassiana had only two isolates, both from the same soil sample from field C in 2015. Both EPF species are found in natural habitats as well as agricultural habitats (Meyling and Eilenberg 2007). However, M. anisopliae, the most prominent and widespread member of the genus is regarded as an agricultural species due to its high

76 prevalence in cultivated/disturbed habitats in comparison to non-disturbed habitats

(Vänninen 1996; Bidochka et al. 1998; Quesada-Moraga et al. 2007; Meyling and

Eilenberg 2007; Schneider et al. 2012). The generalist Metarhizium anisopliae Sorokin was historically considered to have multiple lineages (or varieties) (Driver et al. 2000) but it was later recognized as a species complex of four individual species also known as the PARB clade: M. pingshaense Q.T. Chen & H.L. Guo, M. anisopliae, M. robertsii and

M. brunneum Petch (Bischoff et al. 2009). Kepler et al. (2014) added an additional six species as part of the M. anisopliae species complex: M. globosum J.F. Bisch., Rehner &

Humber; M. acridum and M. lepidiotae (Driver & Milner) J.F. Bisch., Rehner & Humber;

M. guizhouense Q.T. Chen & H.L. Guo; M. majus (J.R. Johnst.) J.F. Bisch., Rehner &

Humber; and M. indigoticum (Kobayasi & Shimizu). Most recently, the M. anisopliae species complex received its eleventh species: M. alvesii Lopes, Faria, Montalva &

Humber (Lopes et al. 2017).

The Metarhizium isolates found in the current study, M. robertsii and M. anisopliae, belong to the PARB clade. Metarhizium robertsii was the predominant species in this survey, accounting for 86.8% of all Metarhizium isolates. This species has a cosmopolitan distribution and it is morphologically identical to M. anisopliae, being only distinguishable with molecular markers (Bischoff et al. 2009; Nishi et al. 2011;

Kepler et al. 2014). It is also known that M. robertsii is rhizosphere competent and it provides plant protection by acting as an insect pathogen and as a beneficial endophyte that stimulates root development (Barelli et al. 2011; Sasan and Bidochka 2012; Pava-

Ripoll 2013). Metarhizium robertsii is also the most prevalent strain in other cropping

77 systems (Bidochka et al. 2001; Wyrebek et al. 2011; Kepler et al. 2015) and it is thought to be the predominant species from the PARB clade in the Holoartic region (Rehner and

Kepler 2017). However, studies indicate that Metarhizium spp. community structures seem to vary among sampled agricultural locations (Steinwender et al. 2014). In other studies, M. anisopliae or M. brunneum (Fisher et al. 2011; Steinwender et al. 2014,

2015; Rezende et al. 2015; Brunner-Mendoza et al. 2017) were the most reported species from the PARB clade in the soil. Metarhizium species community structures are highly governed by site-specific associations such as insect host distribution, plant root associations, temperature activity profile and soil characteristics (Quesada-Moraga et al.

2007; Schneider et al. 2012; Brunner-Mendoza et al. 2017). The factors governing

Metarhizium structure in our study system, in the context of increasing pest management services, should be a topic of further studies.

Finding only two strains of B. bassiana during this study is not surprising or unusual because the species has been more commonly found in non-disturbed habitats than in agricultural soils (Bidochka et al. 1998; Quesada-Moraga et al. 2007; Meyling and

Eilenberg 2007). In Finland, the likelihood for isolating B. bassiana declined by 41.5-

92.4% from un-managed ecosystems to agricultural fields (Vänninen 1996). Beauveria bassiana is regarded as more sensitive to environmental stressors than M. anisopliae

(Bidochka et al. 1998, Quesada-Moraga et al. 2007). This finding is consistent with

Bidochka et al. (2002) who found that only select genetic groups of B. bassiana can survive in agricultural soils. In Iowa, a neighboring state to Nebraska, B. bassiana was rarely recovered using G. mellonella baiting in corn and soybean, Glycine max (L.), fields

78

(Clifton et al. 2015). Moreover, B. bassiana is commonly found above-ground, while M. anisopliae is more predominant belowground (Meyling and Eilenberg 2006b; Meyling et al. 2011). The bulk of the soil samples in this report came from inside cornfields and all were taken belowground, hence, it may be possible that the sampling method chosen restricted the chances of finding B. bassiana strains. However, using similar methodology, but including corn root pieces in the baiting assays with G. mellonella, T. molitor and D. v. virgifera, Rudeen et al. (2013) found B. bassiana in 60% of soil samples from corn root masses in Iowa. Endophytic colonization of corn by B. bassiana strains may be responsible for the differences between Rudeen et al. (2013) and Clifton et al.

2015 and the present study.

Metarhizium and Beauveria species are widely used in classical (Hajek and

Delalibera 2010) and inundative (de Faria and Wraight 2007; Li et al. 2010) biological control programs worldwide, but not in conservation biological control (Mulock and

Chandler 2000a, 2001; Meyling and Eilenberg 2007; Pell et al. 2010). In the irrigated, continuous corn system sampled in this study, a suitable strategy to use the isolated strains would be an inundative approach; however, extensive efficacy and host range tests would need to be performed before large scale field application. In Nebraska, the main pest of continuous corn is the western corn rootworm (WCR) which is also a host for Metarhizium and Beauveria isolates. Studies show that M. anisopliae, M. brunneum and Beauveria bassiana individually or in conjunction with other entomopathogens or

Bt hybrids are able to reduce larval and adult WCR populations (Bruck and Lewis 2001;

Mulock and Chandler 2001; Pilz et al. 2009; Meissle et al. 2009; Petzold-Maxwell et al.

79

2012; Rudeen et al. 2013; Hoffmann et al. 2014). Hence our isolates hold promising potential for developing inundative biological control techniques in this system.

Strains targeted against insects may also work against phytopathogenic organisms and vice-versa. Purpureocillium lilacinum strains have been developed into commercial products to control the eggs cyst and root-knot nematodes. But, P. lilacinum also has topical and endophytic entomopathogenic properties against some insects

(Fiedler and Sosnowska 2007; Lopez et al. 2014). Clonostachys species have been tested as mycoparasites of fungal phytopathogens and against species in Orthoptera,

Lepidoptera and Hemiptera (Jensen et al. 2000; Toledo et al. 2006; Sun and Liu 2008;

Sönmez et al. 2016). Soil EPF make up only a fraction of all fungi and other microorganisms that can be found in the rhizosphere (Jackson et al. 2000). Although our main goal was to find entomopathogens against key insect pests of corn, the isolates from this study have many ecological functions beyond those associated with insects.

Some of the baited fungi are known plant pathogens (Aspergillus spp., Cladosporium spp., Fusarium spp., and Penicillium spp.) or are common saprophytic or opportunistic fungi (G. candidum, Pseudogymnoascus spp., Taifanglia sp. and Talaromyces spp.)

(Table 3). Other strains can have antagonistic properties against insects and/or plant diseases through the production of mycotoxins or endophytic colonization (A. flavus,

Chaetomium sp., Cladosporium sp.) (Table 3.2). This study also identified Trichoderma sp., a BCA genus widely used against phytopathogens worldwide (Howell 2003).

Trichoderma gamsii and T. virens species are known antagonists of Fusarium isolates, including F. oxysporum and F. solani, which were also identified here and are the causal

80 agents of plant wilts and rots in many crops (Rinu et al. 2014). The strains found here should be explored in entomological and plant pathology studies in order to understand factors governing plant protection in this corn system. However, trans-kingdom fungi such as Aspergillus flavus, Chaetomium sp., Cladosporium sp., Fusarium acuminatum,

Fusarium oxysporum, Fusarium solani and P. lilacinum (Table 3.2) should only be explored for use as biological control agents with extreme caution due to the potential risk to crop plants or human health (De Lucca 2007; Luangsa-ard et al. 2011).

This study found an abundant community of entomogenous fungi from commercial fields in Nebraska. Microbial communities and their associated soil processes can have a direct impact on plant health (Garbeva et al. 2004). Beneficial fungi described here may aid in disease and pest suppression in agroecosystems.

Entomopathogenic species made up the majority of fungi isolated from the baiting insects but other fungal ecological roles, such as antagonists of phytopathogens, were described. Understanding the soil fungal community can contribute to the exploration of sustainable agriculture practices. In some regions of the US Corn Belt, such as

Nebraska, corn production is a high-input system where corn is grown without rotation to other crops for many years, if not decades. The use of EPF in this system should be explored alongside current and future insect management practices such as Bt hybrids, insecticides, tillage and rotation. Pests like the WCR require a multi-tactic approach to successfully reduce populations. The next steps will be to test our isolates against the

WCR and other soil-dwelling corn pests to identify strong biological control agents. If a prominent isolate is identified, it could provide an additional or complementary

81 management practice against this pest that has evolved resistance to so many existing control tactics (Meinke et al. 1998; Levine et al. 2002; Parimi et al. 2006; Gassmann et al. 2011, 2014, 2016; Wangila et al. 2015; Zukoff et al. 2016; Ludwick et al. 2017).

82

References

Amaike S, Keller NP (2011) Aspergillus flavus. Annu Rev Phytopathol 49:107–133.

Adesemoye AO, Mayorquin JS, Wang DH, Twizeyimana M, Lynch SC, Eskalen A (2014) Identification of species of Botryosphaeriaceae causing bot gummosis in citrus in California. Plant Dis 98:55–61

Ali-Shtayeh MS, Mara’i A-BBM, Jamous RM (2002) Distribution, occurrence and characterization of entomopathogenic fungi in agricultural soil in the Palestinian area. Mycopathologia 156:235–244.

Barelli L, Padilla-Guerrero IE, Bidochka MJ (2011) Differential expression of insect and plant specific adhesin genes, Mad1 and Mad2, in Metarhizium robertsii. Fungal Biol 115:1174–1185.

Bensch K, Groenewald JZ, Braun U, Dijksterhuis, J, de Jesús Yáñez-Morales M, Crous PW (2015) Common but different: The expanding realm of Cladosporium. Stud Mycol 82:23–74.

Bidochka MJ, Kamp AM, Lavender TM, Dekoning J, De Croos JA (2001) Habitat Association in two genetic groups of the insect-pathogenic fungus Metarhizium anisopliae: uncovering cryptic species? Appl Environ Microbiol 67:1335–1342.

Bidochka MJ, Kasperski JE, Wild GA (1998) Occurrence of the entomopathogenic fungi Metarhizium anisopliae and Beauveria bassiana in soils from temperate and near-northern habitats. Can J Bot 76:1198–1204.

Bidochka MJ, Menzies FV, Kamp AM (2002) Genetic groups of the insect-pathogenic fungus Beauveria bassiana are associated with habitat and thermal growth preferences. Arch Microbiol 178:531–537.

Bischoff JF, Rehner SA, Humber RA (2009) A multilocus phylogeny of the Metarhizium anisopliae lineage. Mycologia 101:512–530

Blehert DS, Hicks AC, Behr M, Meteyer CU, Berlowski-Zier BM, Buckles EL, Coleman JT, Darling SR, Gargas A, Niver R, Okoniewski JC (2009) Bat white-nose syndrome: an emerging fungal pathogen? Science 323:227–227.

Bruck DJ, Lewis LC (2001) Adult Diabrotica spp.(Coleoptera: Chrysomelidae) infection at emergence with indigenous Beauveria bassiana (Deuteromycotina: Hyphomycetes). J Invertebr Pathol 77:288–289

83

Brunner-Mendoza C, Moonjely S, Reyes-Montes M del R, Toriello C, Bidochka M (2017) Physiological and phylogenetic variability of Mexican Metarhizium strains. BioControl 62:779–791.

Carmichael JW (1957) Geotrichum candidum. Mycologia 49:820–830

Chandler D, Hay D, Reid AP (1997) Sampling and occurrence of entomopathogenic fungi and nematodes in UK soils. Appl Soil Ecol 5:133–141.

Chibucos MC, Crabtree J, Nagaraj S, Chaturvedi S, Chaturvedi V (2013) Draft genome sequences of human pathogenic fungus Geomyces pannorum Sensu Lato and bat white nose syndrome pathogen Geomyces (Pseudogymnoascus) destructans. Genome Announc. 1(6): e01045-13. doi:10.1128/genomeA.01045-13.

Clifton EH, Jaronski ST, Hodgson EW, Gassmann AJ (2015) Abundance of soil-borne entomopathogenic fungi in organic and conventional fields in the midwestern USA with an emphasis on the effect of herbicides and fungicides on fungal persistence. PLoS ONE 10(7), e0133613. de Faria MR, Wraight SP (2007) Mycoinsecticides and mycoacaricides: a comprehensive list with worldwide coverage and international classification of formulation types. Biol Control 43:237–256

De Lucca AJ (2007) Harmful fungi in both agriculture and medicine. Rev Iberoam Micol 24:3

Driver F, Milner RJ, Trueman JWH (2000) A taxonomic revision of Metarhizium based on a phylogenetic analysis of rDNA sequence data. Mycol Res 104:134–150

Drummond J, Pinnock DE (1990) Aflatoxin production by entomopathogenic isolates of Aspergillus parasiticus and Aspergillus flavus. J. Invertebr Pathol 55:332–336.

Fernandes ÉKK, Keyser CA, Rangel DEN, Foster RN, Roberts DW (2010) CTC medium: A novel dodine-free selective medium for isolating entomopathogenic fungi, especially Metarhizium acridum, from soil. Biol Control 54:197–205.

Ferrari S, Cribari-Neto F (2004) Beta regression for modelling rates and proportions. J Appl Stat 31:799–815

Fiedler Ż, Sosnowska D (2007) Nematophagous fungus Paecilomyces lilacinus (Thom) Samson is also a biological agent for control of greenhouse insects and mite pests. BioControl 52:547–558.

84

Fisher JJ, Rehner SA, Bruck DJ (2011) Diversity of rhizosphere associated entomopathogenic fungi of perennial herbs, shrubs and coniferous trees. J Invertebr Pathol 106: 289–295

Garbeva P, van Veen JA, van Elsas JD (2004) Microbial Diversity in soil: selection of microbial populations by plant and soil type and implications for disease suppressiveness. Annu Rev Phytopathol 42:243–270.

Gange AC, Eschen R, Wearn JA, Thawer A, Sutton BC (2011) Differential effects of foliar endophytic fungi on insect herbivores attacking a herbaceous plant. Oecologia 168:1023–1031.

Gassmann AJ, Hannon ER, Sisterson MS, Stock SP, Carrière Y, Tabashnik BE (2012) Effects of entomopathogenic nematodes on evolution of pink bollworm resistance to Bacillus thuringiensis Toxin Cry1Ac. J Econ Entomol 105:994–1005.

Gassmann AJ, Petzold-Maxwell JL, Clifton EH, Stock SP, Carrière Y, Tabashnik BE (2014) Field-evolved resistance by western corn rootworm to multiple Bacillus thuringiensis toxins in transgenic maize. Proc Natl Acad Sci 111:5141–5146.

Gassmann AJ, Petzold-Maxwell JL, Keweshan RS, Dunbar MW (2011) Field-evolved resistance to bt maize by western corn rootworm. PLoS ONE 6:e22629.

Gassmann AJ, Shrestha RB, Jakka SRK, Dunbar MW, Clifton EH, Paolino AR, Ingber DA, French BW, Masloski KE, Dounda JW, St. Clair CR (2016) Evidence of resistance to Cry34/35Ab1 corn by western corn rootworm (Coleoptera: Chrysomelidae): root injury in the field and larval survival in plant-based bioassays. J Econ Entomol 109(4):1872-80.

Glass NL, Donaldson GC (1995) Development of primer sets designed for use with the PCR to amplify conserved genes from filamentous ascomycetes. Appl Environ Microbiol 61:1323–1330.

Gray, M. E., T. W. Sappington, N. J. Miller, J. Moeser, and M. O. Bohn. 2009. Adaptation and invasiveness of western corn rootworm: intensifying research on a worsening pest. Annu. Rev. Entomol. 54: 303–321.

Guo JW, Cheng JS, Yang LF, Zhou YL, Wang HF, Chen JY, Abdala Mohamad O, Tian XJ, Liu YH, Liu YH, Li WJ (2016). First report of a leaf spot disease caused by Chaetomium globosum on pomegranate from Yunnan, China. Plant Disease. 100(1), pp 223.

Hajek AE, Delalibera I (2010) Fungal pathogens as classical biological control agents against arthropods. BioControl 55:147–158.

85

Hajek AE, Delalibera I, Mcmanus ML (2000) Introduction of exotic pathogens and documentation of their establishment and impact. in: Field manual of techniques in invertebrate pathology, 2nd edn. Springer, Dordrecht, pp 339–369

Hannon ER, Sisterson MS, Stock SP, Carrière Y, Tabashnik BE, Gassmann AJ (2010) Effects of Four nematode species on fitness costs of pink bollworm resistance to Bacillus thuringiensis toxin CrylAc. J Econ Entomol 103:1821–1831.

Harris CR (1972) Factors influencing the effectiveness of soil insecticides. Annu Rev Entomol 17:177–198

Hoffmann AM, French BW, Jaronski ST, Gassmann AJ (2014) Effects of entomopathogens on mortality of western corn rootworm (Coleoptera: Chrysomelidae) and fitness costs of resistance to Cry3Bb1 maize. J Econ Entomol 107:352–360

Howell CR (2003) Mechanisms employed by Trichoderma species in the biological control of plant diseases: the history and evolution of current concepts. Plant Dis 87:4–10.

Humber RA (2000) Fungal pathogens and parasites of insects. in: Applied microbial systematics. Springer, Dordrecht, pp 203–230

Humber RA (2016) Seeking stability for research and applied uses of entomopathogenic fungi as biological control agents. J. Asia-Pac Entomol 19:1019–1025.

Jackson TA, Alves SB, Pereira RM (2000) Success in biological control of soil-dwelling insects by pathogens and nematodes. In: Gurr G, Wratten S (eds) Biological control: measures of success. Springer Netherlands, pp 271–296

Janevska S, Tudzynski B (2018) Secondary metabolism in Fusarium fujikuroi: strategies to unravel the function of biosynthetic pathways. Appl Microbiol Biotechnol 102:615–630.

Jensen B, Knudsen IM, Jensen DF (2000) Biological seed treatment of cereals with fresh and long-term stored formulations of Clonostachys rosea: biocontrol efficacy against Fusarium culmorum. Eur J Plant Pathol 106:233–242

Kepler RM, Humber RA, Bischoff JF, Rehner SA (2014) Clarification of generic and species boundaries for Metarhizium and related fungi through multigene phylogenetics. Mycologia 106:811–829.

Kepler RM, Ugine TA, Maul JE, Cavigelli MA, Rehner SA (2015) Community composition and population genetics of insect pathogenic fungi in the genus Metarhizium

86

from soils of a long-term agricultural research system. Environ Microbiol 17:2791–2804.

Klingen I, Eilenberg J, Meadow R (2002) Effects of farming system, field margins and bait insect on the occurrence of insect pathogenic fungi in soils. Agric Ecosyst Environ 91:191–198

Klingen, I., Haukeland, S. (2006) The soil as a reservoir for natural enemies of pest insects and mites with emphasis on fungi and nematodes. In: An ecological and societal approach to biological control. Series: Progress in biological control, Vol. 2 Spr. 145-211.

Kozakiewicz Z (1989) Aspergillus species on stored products. Mycol Papers.

Kucey RMN (1983) Phosphate-solubilizing bacteria and fungi in various cultivated and virgin Alberta soils. Can J Soil Sci 63:671–678

Lacey LA, Frutos R, Kaya HK, Vail P (2001) Insect pathogens as biological control agents: do they have a future? Biol Control 21:230–248.

Lacey LA, Grzywacz D, Shapiro-Ilan DI, Frutos R, Brownbridge M, Goettel MS (2015) Insect pathogens as biological control agents: Back to the future. J Invertebr Pathol 132:1–41.

Lacey, LA, Solter L F (2012) Initial handling and diagnosis of diseased invertebrates. In: Manual of techniques in invertebrate pathology, 2nd edn. Elsevier, London pp. 1-14.

Leushkin EV, Logacheva MD, Penin AA, Sutormin RA, Gerasimov ES, Kochkina GA, Ivanushkina NE, Vasilenko OV, Kondrashov AS, Ozerskaya SM (2015) Comparative genome analysis of Pseudogymnoascus spp. reveals primarily clonal evolution with small genome fragments exchanged between lineages. BMC Genomics 16:400.

Levine E, Spencer JL, Isard SA, Onstad DW, Gray ME (2002) adaptation of the western corn rootworm to crop rotation: evolution of a new strain in response to a management practice. Am Entomol 48:94–107.

Li Z, Alves SB, Roberts DW, Fan M, Delalibera Jr I, Tang J, Lopes RB, Faria M, Rangel DE (2010) Biological control of insects in Brazil and China: history, current programs and reasons for their successes using entomopathogenic fungi. Biocontrol Sci Technol 20:117–136.

Lopes RB, Souza DA, Rocha LF, Montalva C, Luz C, Humber RA, Faria M. (2017) Metarhizium alvesii sp. nov.: A new member of the Metarhizium anisopliae species complex. J Invertebr Pathol 151:165-168

87

Lopez DC, Zhu-Salzman K, Ek-Ramos MJ, Sword GA (2014) The entomopathogenic fungal endophytes Purpureocillium lilacinum (formerly Paecilomyces lilacinus) and Beauveria bassiana negatively affect cotton aphid reproduction under both greenhouse and field conditions. PloS One 9:e103891

Luangsa-ard J, Houbraken J, van Doorn T, Hong SB, Borman AM, Hywel-Jones NL, Samson RA (2011) Purpureocillium, a new genus for the medically important Paecilomyces lilacinus. FEMS Microbiol Lett 321:141–149.

Ludwick DC, Meihls LN, Ostlie KR, Potter BD, French L, Hibbard BE (2017) Minnesota field population of western corn rootworm (Coleoptera: Chrysomelidae) shows incomplete resistance to Cry34Ab1/Cry35Ab1 and Cry3Bb1. J Appl Entomol 141:28–40.

Massenssini AM, Bonduki VHA, Melo CA, Totola MR, Ferreira FA, Costa MD (2014) Soil microorganisms and their role in the interactions between weeds and crops. Planta Daninha 32:873–884.

Meinke LJ, Siegfried BD, Wright RJ, Chandler LD (1998) Adult susceptibility of Nebraska western corn rootworm (Coleoptera: Chrysomelidae) populations to selected insecticides. J Econ Entomol 91:594–600

Meissle M, Pilz C, Romeis J (2009) Susceptibility of Diabrotica virgifera virgifera (Coleoptera: Chrysomelidae) to the entomopathogenic fungus Metarhizium anisopliae when feeding on Bacillus thuringiensis Cry3Bb1-Expressing Maize. Appl Environ Microbiol 75:3937–3943.

Meyling NV, Eilenberg J (2007) Ecology of the entomopathogenic fungi Beauveria bassiana and Metarhizium anisopliae in temperate agroecosystems: Potential for conservation biological control. Biol Control 43:145–155.

Meyling NV, Eilenberg J (2006a) Occurrence and distribution of soil borne entomopathogenic fungi within a single organic agroecosystem. Agric Ecosyst Environ 113:336–341

Meyling NV, Eilenberg J (2006b) Isolation and characterisation of Beauveria bassiana isolates from phylloplanes of hedgerow vegetation. Mycol Res 110:188–195

Meyling NV, Hajek AE (2010) Principles from community and metapopulation ecology: application to fungal entomopathogens. BioControl 55:39–54.

Meyling NV, Thorup-Kristensen K, Eilenberg J (2011) Below-and aboveground abundance and distribution of fungal entomopathogens in experimental conventional and organic cropping systems. Biol Control 59:180–186

88

Müller-Stöver D, Nybroe O, Baraibar B, Loddo D, Eizenberg H, French K, Sønderskov M, Neve P, Peltzer DA, Maczey N, Christensen S (2016) Contribution of the seed microbiome to weed management. Weed Res 56:335–339

Mulock B, Chandler L (2000) Field-cage studies of Beauveria bassiana (Hyphomycetes: Moniliaceae) for the suppression of adult western corn rootworm, Diabrotica virgifera virgifera (Coleoptera: Chrysomelidae). Biocontrol Sci Technol 10:51–60.

Mulock BS, Chandler LD (2001) Effect of Beauveria bassiana on the fecundity of western corn rootworm, Diabrotica virgifera virgifera (Coleoptera: Chrysomelidae). Biol Control 22:16–21.

Navarro-Velasco, G.Y., Prados-Rosales, R.C., Ortíz-Urquiza, A., Quesada-Moraga, E., Di Pietro, A. (2011) Galleria mellonella as model host for the trans-kingdom pathogen Fusarium oxysporum. Fungal Gen. and Biol. 48: 1124-1129 Nishi O, Hasegawa K, Iiyama K, Yasunaga-Aoki C, Shimizu S (2011) Phylogenetic analysis of Metarhizium spp. isolated from soil in Japan. Appl Entomol Zool 46:301–309

Oliveira I, Pereira JA, Lino-Neto T, Bento A, Baptista P (2011) Fungal diversity associated to the olive moth, Prays oleae Bernard: A survey for potential entomopathogenic fungi. Microb Ecol 63:964–974.

Parimi S, Meinke LJ, French WB, Chandler LD, Siegfried BD (2006) Stability and persistence of aldrin and methyl-parathion resistance in western corn rootworm populations (Coleoptera: Chrysomelidae). Crop Prot 25:269–274.

Pava-Ripoll M (2013) Metarhizium robertsii, a rhizosphere-competent insect pathogen. In: Bruijn FJ de (ed) Molecular Microbial Ecology of the Rhizosphere. John Wiley & Sons, Inc., pp 149–159

Pell JK, Hannam JJ, Steinkraus DC (2010) Conservation biological control using fungal entomopathogens. BioControl 55:187–198.

Petzold-Maxwell JL, Jaronski ST, Gassmann AJ (2012) Tritrophic interactions among Bt maize, an insect pest and entomopathogens: effects on development and survival of western corn rootworm. Ann Appl Biol 160:43–55.

Pilz C, Keller S, Kuhlmann U, Toepfer S (2009) Comparative efficacy assessment of fungi, nematodes and insecticides to control western corn rootworm larvae in maize. BioControl 54:671–684.

Pilz C, Wegensteiner R, Keller S (2008) Natural occurrence of insect pathogenic fungi and insect parasitic nematodes in Diabrotica virgifera virgifera populations. BioControl 53:353–359

89

Pitt JI (2002) Biology and ecology of toxigenic Penicillium species. In: Mycotoxins and Food Safety. Springer, Boston, MA, pp 29–41

Quesada-Moraga E, Navas-Cortés JA, Maranhao EAA, Ortiz-Urquiza A, Santiago- Álvarez C (2007) Factors affecting the occurrence and distribution of entomopathogenic fungi in natural and cultivated soils. Mycol Res 111:947–966.

Rehner SA, Kepler RM (2017) Species limits, phylogeography and reproductive mode in the Metarhizium anisopliae complex. J Invertebr Pathol 148:60–66

Rezende JM, Zanardo ABR, Lopes M da S, Delalibera I, Rehner SA (2015) Phylogenetic diversity of Brazilian Metarhizium associated with sugarcane agriculture. BioControl 60:495–505.

Rinu K, Sati P, Pandey A (2014) Trichoderma gamsii (NFCCI 2177): A newly isolated endophytic, psychrotolerant, plant growth promoting, and antagonistic fungal strain. J Basic Microbiol 54:408–417.

Roberts DW, Humber RA (1981) Entomogeneous fungi. In: Biology of Conidial Fungi. Elsevier

Roberts DW, St Leger RJ (2004) Metarhizium spp., cosmopolitan insect-pathogenic fungi: mycological aspects. Adv Appl Microbiol 54:1–70

Roy HE, Steinkraus DC, Eilenberg J, Hajek AE, Pell JK (2006) Bizarre interactions and endgames: entomopathogenic fungi and their arthropod hosts. Annu Rev Entomol 51:331–357

Rudeen ML, Jaronski ST, Petzold-Maxwell JL, Gassmann AJ (2013) Entomopathogenic fungi in cornfields and their potential to manage larval western corn rootworm Diabrotica virgifera virgifera. J Invertebr Pathol 114:329–332.

Sasan RK, Bidochka MJ (2012) The insect-pathogenic fungus Metarhizium robertsii (Clavicipitaceae) is also an endophyte that stimulates plant root development. Am J Bot 99:101–107. doi: 10.3732/ajb.1100136

Schabenberger O (2005) Introducing the GLIMMIX procedure for generalized linear mixed models. SUGI 30 Proc. 196.

Schneider S, Widmer F, Jacot K, Kölliker R, Enkerli J (2012) Spatial distribution of Metarhizium clade 1 in agricultural landscapes with arable land and different semi-natural habitats. Appl Soil Ecol 52:20–28

Shah PA, Pell JK (2003) Entomopathogenic fungi as biological control agents. Appl Microbiol Biotechnol 61:413–423. doi: 10.1007/s00253-003-1240-8

90

Singh S, Pandey RK, Goswami BK (2013) Bio-control activity of Purpureocillium lilacinum strains in managing root-knot disease of tomato caused by Meloidogyne incognita. Biocontrol Sci Technol 23:1469–1489.

Solter LF, Hajek AE, Lacey LA (2017) Exploration for entomopathogens. In: Microbial control of insect and mite pests. Academic Press, pp 13–23

Sönmez E, Sevim A, Demirbağ Z, Demir İ (2016) Isolation, characterization and virulence of entomopathogenic fungi from Gryllotalpa gryllotalpa (Orthoptera: Gryllotalpidae). Appl Entomol Zool 51:213–223.

Soytong K (1992) Biological control of tomato wilt caused by Fusarium oxysporum f. sp. lycopersici using Chaetomium cupreum. Kasetsart JNat Sci 26:310–313

Steinwender BM, Enkerli J, Widmer F, Eilenberg J, Thorup-Kristensen K, Meyling NV (2014) Molecular diversity of the entomopathogenic fungal Metarhizium community within an agroecosystem. J Invertebr Pathol 123:6–12

Stroup WW (2015) Rethinking the analysis of non-normal data in plant and soil science. Agron J 107:811–827

Sun B-D, Liu X-Z (2008) Occurrence and diversity of insect-associated fungi in natural soils in China. Appl Soil Ecol 39:100–108

Sun B-D, Yu H, Chen AJ, Liu X-Z (2008) Insect-associated fungi in soils of field crops and orchards. Crop Prot 27:1421–1426.

Tilman D, Cassman KG, Matson PA, Naylor R, Polasky S (2002) Agricultural sustainability and intensive production practices. Nature 418:671–677

Toledo AV, Virla E, Humber RA, Paradell SL, Lastra CL (2006) First record of Clonostachys rosea (Ascomycota: Hypocreales) as an entomopathogenic fungus of Oncometopia tucumana and Sonesimia grossa (Hemiptera: Cicadellidae) in Argentina. J Invertebr Pathol 92:7–10. doi: 10.1016/j.jip.2005.10.005

Tooker JF, Douglas MR, Krupke CH (2017) Neonicotinoid Seed Treatments: Limitations and Compatibility with Integrated Pest Management. Agric Environ Lett 2:. doi: 10.2134/ael2017.08.0026

USDA-FAS. Grain: World Market and Trade. 2017. http://usda.mannlib.cornell.edu/usda/fas/grain-market//2010s/2017/grain-market-12- 12-2017.pdf. Accessed 4.April.2018

91

Vänninen I (1996) Distribution and occurrence of four entomopathogenic fungi in Finland: effect of geographical location, habitat type and soil type. Mycol Res 100:93–101.

Varga J, Tóth B, Kocsubé S, Farkas B, Szakács G, Téren J, Kozakiewicz Z (2005) Evolutionary Relationships Among Aspergillus terreus Isolates and their Relatives. Antonie Van Leeuwenhoek 88: 141–150. doi: 10.1007/s10482-005- 3870-6

Vega FE, Meyling NV, Luangsa-ard JJ, Blackwell M (2012) Fungal entomopathogens. In: Kaya FEVK (ed) Insect pathology (Second Edition). Academic Press, San Diego, pp 171–220

Vojvodic S, Jensen AB, James RR, Boomsma JJ, Eilenberg J (2011) Temperature dependent virulence of obligate and facultative fungal pathogens of honeybee brood. Vet Microbiol 149:200–205

Wakil W, Ghazanfar MU, Riasat T, Kwon YJ, Qayyum MA, Yasin M (2013) Occurrence and diversity of entomopathogenic fungi in cultivated and uncultivated soils in Pakistan. Entomol Res 43:70–78.

Wang C, Wang S (2017) Insect pathogenic fungi: genomics, molecular interactions, and genetic improvements. Annu Rev Entomol 62:73–90.

Wangila DS, Gassmann AJ, Petzold-Maxwell JL, French BW, Meinke LJ (2015) Susceptibility of Nebraska western corn rootworm (Coleoptera: Chrysomelidae) populations to Bt corn events. J Econ Entomol 108:742–751.

Wenda-Piesik A, Sun Z, Grey WE, Weaver DK, Morrill WL (2009) Mycoses of wheat stem sawfly (Hymenoptera: Cephidae) larvae by Fusarium spp. isolates. Environ Entomol 38:387–394.

White TJ, Bruns T, Lee S, Taylor JL (1990) Amplification and direct sequencing of fungal ribosomal RNA genes for phylogenetics. PCR Protoc Guide Methods Appl 18:315–322

Wicklow DT, Dowd PF, Gloer JB (1999) Chaetomium mycotoxins with antiinsectan or antifungal activity. Mycotoxins 1999:267–271.

Wraight, S.P., G.D. Inglis, and M.S. Goettel (2007) Fungi. Lacey, L. A., and Kaya, H. K. (Eds.). Field manual of techniques in invertebrate pathology. Dordrecht etc.: Kluwer acad. publ. Cop. Pp.223-249

92

Wyrebek M, Huber C, Sasan RK, Bidochka MJ (2011) Three sympatrically occurring species of Metarhizium show plant rhizosphere specificity. Microbiology 157:2904–2911

Yilmaz N, Visagie CM, Houbraken J, Frisvad JC, Samson RA (2014) Polyphasic taxonomy of the genus Talaromyces. Stud Mycol 78:175–341. doi: 10.1016/j.simyco.2014.08.001

Zimmermann G (1986) The ‘Galleria bait method’ for detection of entomopathogenic fungi in soil. J Appl Entomol 102:213–215.

Zhang Y, Liu F, Wu W, Cai L (2015) A phylogenetic assessment and taxonomic revision of the thermotolerant hyphomycete genera Acrophialophora and Taifanglania. Mycologia 107:768–779.

Zukoff SN, Ostlie KR, Potter B, Meihls LN, Zukoff AL, French L, Ellersieck MR, Wade French B, Hibbard BE (2016) Multiple assays indicate varying levels of cross resistance in Cry3Bb1-selected field populations of the western corn rootworm to mCry3A, eCry3.1Ab, and Cry34/35Ab1. J Econ Entomol 109(3):1387-98

Tables

Table 3.1 Fungal isolates from irrigated cornfields identified by the internal transcriber spacer (ITS) and beta-tubulin (BT) regions. Strains were isolated via baiting assays of G. mellonella (G.M.) (2014 and 2015) and T. molitor (T.M.) (2015 only). Gen Bank Accession Number Insect No. Isolate Locality Year Host ITS BT Species ID 1 E998 A 2015 G.M. MF681598 - Aspergillus flavus 2 E311 B 2014 G.M. MF681539 MH193388 Aspergillus terreus 3 E1040 C 2015 G.M. MF681618 - Beauveria bassiana 4 E1041 C 2015 G.M. MF681619 - Beauveria bassiana 5 E312 A 2014 G.M. MF681540 - Chaetomium sp. 6 E126 C 2014 G.M. MF681516 - Cladosporium halotolerans 7 E1060 B 2015 T.M. MF681624 - Cladosporium sp. 8 E651 B 2015 T.M. MF681583 - Clonostachys sp. 9 E1039 C 2015 T.M. MF681617 - Clonostachys sp. 10 E649 B 2015 T.M. MG654677 - Fusarium acuminatum 11 E1010 B 2015 G.M. MG654680 - Fusarium acuminatum 12 E163 B 2014 G.M. MG654673 - Fusarium fujikuroi 13 E171 B 2014 G.M. MG654674 - Fusarium oxysporum 14 E648 B 2015 T.M. MG654676 - Fusarium oxysporum 15 E641 A 2015 T.M. MG654675 - Fusarium solani 16 E656 E 2015 G.M. MG654678 - Fusarium solani 17 E999 A 2015 T.M. MG654679 - Fusarium solani 18 E1049 B 2015 T.M. MG654681 - Fusarium solani 19 E1072 D 2015 G.M. MG654682 - Fusarium solani 20 E320 A 2015 G.M. MF681547 - Geotrichum candidum 21 E127 B 2014 G.M. MF681517 - Metarhizium robertsii 93

Table 3.1 (Continued) Gen Bank Accession Number Insect No. Isolate Locality Year Host ITS BT Species ID 22 E136 B 2014 G.M. MF681518 MH048528 Metarhizium robertsii 23 E137 D 2014 G.M. MF681519 MH048529 Metarhizium robertsii 24 E138 B 2014 G.M. MF681520 - Metarhizium robertsii 25 E156 D 2014 G.M. MF681521 MH048530 Metarhizium robertsii 26 E157 E 2014 G.M. MF681522 MH048531 Metarhizium robertsii 27 E160 E 2014 G.M. MF681523 - Metarhizium robertsii 28 E161 C 2014 G.M. MF681524 - Metarhizium robertsii 29 E169 C 2014 G.M. MF681526 - Metarhizium robertsii 30 E211 E 2014 G.M. MF681528 MH048532 Metarhizium robertsii 31 E213 A 2014 G.M. MF681530 MH048533 Metarhizium anisopliae 32 E214 C 2014 G.M. MF681531 MH048534 Metarhizium robertsii 33 E215 C 2014 G.M. MF681532 MH048535 Metarhizium robertsii 34 E217 C 2014 G.M. MF681533 MH048536 Metarhizium robertsii 35 E275 B 2014 G.M. MF681534 MH048537 Metarhizium robertsii 36 E276 E 2014 G.M. MF681535 MH048538 Metarhizium robertsii 37 E277 A 2014 G.M. MF681536 MH048539 Metarhizium robertsii 38 E278 E 2014 G.M. MF681537 MH048540 Metarhizium anisopliae 39 E322 E 2014 G.M. MF681548 MH048541 Metarhizium robertsii 40 E323 E 2014 G.M. MF681549 MH048542 Metarhizium robertsii 41 E324 E 2014 G.M. MF681550 MH048543 Metarhizium robertsii 42 E328 B 2014 G.M. MF681552 MH048544 Metarhizium robertsii 43 E335 C 2014 G.M. MF681556 MH048545 Metarhizium robertsii 44 E367 B 2014 G.M. MF681557 MH048546 Metarhizium robertsii

94

Table 3.1 (Continued) Gen Bank Accession Number Insect No. Isolate Locality Year Host ITS BT Species ID 45 E369 B 2014 G.M. MF681559 MH048547 Metarhizium robertsii 46 E371 E 2014 G.M. MF681561 MH048548 Metarhizium robertsii 47 E374 B 2014 G.M. MF681564 MH048549 Metarhizium robertsii 48 E377 C 2014 G.M. MF681566 MH048550 Metarhizium robertsii 49 E380 C 2014 G.M. MF681568 MH048551 Metarhizium robertsii 50 E642 A 2015 T.M. MF681577 MH048552 Metarhizium robertsii 51 E645 D 2015 T.M. MF681579 MH048553 Metarhizium robertsii 52 E647 B 2015 T.M. MF681581 MH048554 Metarhizium robertsii 53 E650 B 2015 T.M. MF681582 MH048555 Metarhizium robertsii 54 E652 B 2015 T.M. MF681584 MH048556 Metarhizium robertsii 55 E653 B 2015 T.M. MF681585 MH048557 Metarhizium robertsii 56 E654 E 2015 T.M. MF681586 MH048558 Metarhizium robertsii 57 E658 C 2015 G.M. MF681587 MH048559 Metarhizium robertsii 58 E982 B 2015 G.M. MF681588 - Metarhizium robertsii 59 E983 B 2015 G.M. MF681589 MH048560 Metarhizium robertsii 60 E985 B 2015 G.M. MF681590 MH048561 Metarhizium robertsii 61 E987 B 2015 T.M. MF681591 MH048562 Metarhizium robertsii 62 E989 B 2015 G.M. MF681592 MH048563 Metarhizium robertsii 63 E991 B 2015 T.M. MF681593 MH048564 Metarhizium robertsii 64 E992 B 2015 G.M. MF681594 MH048565 Metarhizium robertsii 65 E994 E 2015 G.M. MF681595 MH048566 Metarhizium robertsii 66 E996 A 2015 T.M. MF681596 - Metarhizium robertsii 67 E997 A 2015 T.M. MF681597 - Metarhizium robertsii 68 E1000 E 2015 G.M. MF681599 MH048567 Metarhizium robertsii 95

Table 3.1 (Continued) Gen Bank Accession Number Insect No. Isolate Locality Year Host ITS BT Species ID 69 E1001 E 2015 G.M. MF681600 MH048568 Metarhizium robertsii 70 E1005 E 2015 G.M. MF681601 MH048569 Metarhizium robertsii 71 E1012 B 2015 G.M. MF681603 MH048570 Metarhizium robertsii 72 E1016 A 2015 T.M. MF681604 MH048571 Metarhizium robertsii 73 E1020 B 2015 G.M. MF681605 MH048572 Metarhizium robertsii 74 E1022 A 2015 G.M. MF681606 MH048573 Metarhizium robertsii 75 E1023 C 2015 G.M. MF681607 MH048574 Metarhizium robertsii 76 E1024 C 2015 T.M. MF681608 MH048575 Metarhizium robertsii 77 E1026 C 2015 T.M. MF681609 MH048576 Metarhizium robertsii 78 E1030 D 2015 T.M. MF681610 MH048577 Metarhizium robertsii 79 E1033 B 2015 G.M. MF681612 MH048578 Metarhizium anisopliae 80 E1034 B 2015 G.M. MF681613 - Metarhizium anisopliae 81 E1037 A 2015 T.M. MF681616 - Metarhizium anisopliae 82 E1045 B 2015 T.M. MF681620 MH048579 Metarhizium robertsii 83 E1053 B 2015 T.M. MF681621 MH048580 Metarhizium robertsii 84 E1054 E 2015 T.M. MF681622 MH048581 Metarhizium robertsii 85 E1056 E 2015 G.M. MF681623 MH048582 Metarhizium robertsii 86 E1061 B 2015 G.M. MF681625 MH048583 Metarhizium sp. 87 E1066 E 2015 G.M. MF681627 - Metarhizium sp. 88 E1068 E 2015 G.M. MF681628 - Metarhizium sp. 89 E1074 E 2015 G.M. MF681629 - Metarhizium robertsii 90 E1080 B 2015 T.M. MF681630 MH048584 Metarhizium robertsii 91 E1081 B 2015 T.M. MF681631 MH048585 Metarhizium anisopliae

96

Table 3.1 (Continued) Gen Bank Accession Number Insect No. Isolate Locality Year Host ITS BT Species ID 92 E1084 B 2015 G.M. MF681632 MH048586 Metarhizium robertsii 93 E1089 A 2015 G.M. MF681633 MH048587 Metarhizium anisopliae 94 E1090 E 2015 G.M. MF681634 MH048588 Metarhizium robertsii 95 E1092 B 2015 G.M. MF681635 MH048589 Metarhizium robertsii 96 E1093 B 2015 G.M. MF681636 MH048590 Metarhizium robertsii 97 E314 C 2014 G.M. MF681541 - Neosartorya sp. 98 E368 D 2014 G.M. MF681558 MH193396 Penicillium bilaiae 99 E1036 E 2015 G.M. MF681615 MH193407 Penicillium bilaiae 100 E212 E 2014 G.M. MF681529 - Penicillium griseofulvum 101 E166 A 2014 G.M. MF681525 MH193386 Penicillium griseofulvum 102 E172 E 2014 G.M. MF681527 - Penicillium janthinellum 103 E317 B 2014 G.M. MF681544 MH193391 Penicillium janthinellum 104 E334 C 2014 G.M. MF681555 MH193395 Penicillium janthinellum 105 E1035 B 2015 G.M. MF681614 MH193406 Penicillium janthinellum. 106 E124 E 2014 G.M. MF681515 - Penicillium raistrickii 107 E370 A 2014 G.M. MF681560 MH193397 Pseudogymnoascus destructans 108 E376 B 2014 G.M. MF681565 - Pseudogymnoascus destructans 109 E395 A 2014 G.M. MF681575 MH193403 Pseudogymnoascus destructans 110 E393 A 2014 G.M. MF681573 - Pseudogymnoascus pannorum

97

Table 3.1 (Continued) Gen Bank Accession Number Insect No. Isolate Locality Year Host ITS BT Species ID 111 E372 E 2014 G.M. MF681562 MH193398 Pseudogymnoascus sp. 112 E392 A 2014 G.M. MF681572 - Pseudogymnoascus sp. 113 E394 C 2014 G.M. MF681574 - Pseudogymnoascus sp. 114 E397 D 2014 G.M. MF681576 MH193404 Pseudogymnoascus sp. 115 E279 A 2014 G.M. MF681538 MH193387 Purpureocillium lilacinum 116 E318 A 2014 G.M. MF681545 MH193392 Purpureocillium lilacinum 117 E319 D 2014 G.M. MF681546 - Purpureocillium lilacinum 118 E378 A 2014 G.M. MF681567 - Purpureocillium lilacinum 119 E325 B 2014 G.M. MF681551 - Taifanglania sp. 120 E646 B 2015 T.M MF681580 MH193405 Talaromyces pinophilus 121 E390 C 2014 G.M. MF681571 MH193402 Talaromyces trachyspermus 122 E315 B 2014 G.M. MF681542 MH193389 Talaromyces trachyspermus 123 E316 D 2014 G.M. MF681543 MH193390 Talaromyces trachyspermus 124 E331 E 2014 G.M. MF681553 MH193393 Talaromyces trachyspermus 125 E332 C 2014 G.M. MF681554 MH193394 Talaromyces trachyspermus 126 E373 D 2014 G.M. MF681563 MH193399 Talaromyces trachyspermus 127 E381 A 2014 G.M. MF681569 MH193400 Talaromyces trachyspermus 128 E389 B 2014 G.M. MF681570 MH193401 Talaromyces trachyspermus 129 E644 A 2015 T.M. MF681578 - Talaromyces trachyspermus 130 E1032 B 2015 G.M. MF681611 - Trichoderma gamsii 131 E1064 B 2015 G.M. MF681626 - Trichoderma gamsii 132 E1007 B 2015 G.M. MF681602 - Trichoderma virens 98

Table 3.2 Ecological functions associated with baited fungi. The concept for this table was developed from Sun et al. 2008 and Oliveira et al. 2011. Percentage column is the frequency of each species from the 132 samples sent for molecular identification. Fungal Identification N % EPF Plant Path. Entomogenous Potential Others References Antagonist / Insect Plant Antagonists Pathogens

Bionectriaceae

Clonostachys sp. 2 1.52 X X Toledo et al. 2006, Jensen et al. 2000 Chaetomiacea

Chaetomium sp. 1 0.76 X X X Gange et al 2011, Wicklow et al. 1999, Soytong et al. 1992, Guo et al. 2016 Taifanglania sp. 1 0.76 X Zhang et al. 2015

Cladosporiaceae

Cladosporium halotolerans Zalar 1 0.76 X X Bensch et al. 2015

Cladosporium sp. 1 0.76 X X X Gange et al 2011

Clavicipitaceae Metarhizium anisopliae (Metschn.) 7 5.30 X Shah and Pell 2003 Sorokin Metarhizium robertsii J.F. Bisch., 66 50 X Sasan and. Rehner & Humber Bidochka 2012 Metarhizium sp. 3 2.27 X Shah and Pell 2003

Cordycipitaceae

Beauveria bassiana (Bals.-Criv.)Vuill. 2 1.52 X Shah and Pell 2003

99

Table 3.2 (Continued) Fungal Identification N % EPF Plant Path. Entomogenous Potential Others References Antagonist / Insect Plant Antagonists Pathogens

Dipodascaceae

Geotrichum candidum Link 1 0.76 X Carmichael 1957

Hypocreaceae

Trichoderma gamsii Samuels & 2 1.52 X Rinu et al. 2014 Druzhin. Trichoderma virens (J.H. Mill., 1 0.76 X Howell 2003 Giddens & A.A. Foster) Arx

Myxotrichaceae Pseudogymnoascus destructans 3 2.27 X Blehert et al 2009 (Blehert & Gargas) Minnis & D.L. Lindner Pseudogymnoascus pannorum 1 0.76 X Chibucos et al. 2013 (Link) Minnis & D.L. Lindner Pseudogymnoascus sp. 4 3.03 X Leushkin et al. 2015

Nectriaceae

Fusarium acuminatum Ellis & 2 1.52 X X Wenda-Piesik et al. Everh. 2009 Fusarium fujikuroi Nirenberg 1 0.76 X Janevska and Tudzynski 2018 Fusarium oxysporum 2 1.52 X X Navarro-Velasco et al. Schlechtendal 2011 Fusarium solani (Mart.) Sacc. 5 3.79 X X Sun and Liu 2008

100

Table 3.2 (Continued) Fungal Identification N % EPF Plant Path. Entomogenous Potential Others References Antagonist / Insect Plant Antagonists Pathogens

Ophiocordycipitaceae

Purpureocillium lilacinum (Thom) 4 3.03 X X Lopez 2014, Singh et al 2013 Luangsa-ard, Houbraken, Hywel- Jones & Samson

Trichocomaceae Amaike and Keller 2011, Aspergillus flavus Link 1 0.76 X X X Vojvodic et al. 2011, Drummond and Pinnock 1990

Aspergillus terreus Thom 1 0.76 X X Varga et al. 2005 Neosartorya sp. 1 0.76 X Kozakiewicz 1989 Penicillium bilaiae Chalabuda 2 1.52 X X Kucey 1983

Penicillium griseofulvum Dierckx 2 1.52 X X Pitt 2002 Penicillium janthinellum Biourge 4 3.03 X X Pitt 2002 Penicillium raistrickii G. Smith 1 0.76 X X Pitt 2002 Talaromyces pinophilus (Hedgc.) 1 0.76 X Yilmaz et al. 2014 Samson, N. Yilmaz, Frisvad & Seifert Talaromyces trachyspermus (Shear) 9 6.82 X Yilmaz et al. 2014 Stolk & Samson

101

102

Figures

(a)

(b)

Fig. 3.1. Locations sampled in Keith and Perkins Counties in Nebraska in 2014 and 2015. a) Fields represented by letters, field characteristics can be found in Appendix 1. All fields in Perkins Co. were at least 5 miles apart, fields “a” and “e” were 20 miles apart. and b) Sampling diagram of the soil collection zone from inside each center-pivot irrigated cornfield. Cross-markings represent soil samples along the second and third pivot tire track (60-120 meters from field edge).

a

ab

abc abc

abc abc abc abc bc abc abc abc bc abc abc abc bc

bc bc bc bc bc bc bc bc bc bc c c *

Fig.3.2. Mean percent of cadavers infected with entomogenous fungi per arena in 2014. Borders and irrigated field combined for a total of 10 arenas per sample date by field combination. Field by date interaction was significant (F23,306 =1.61, Pr>F=0.0394). Simple effect comparisons of field x date least square means adjusted with Tukey’s adjustment. Letters represent means significantly different at P< 0.05. Field C had no collection date on 22. July because of pesticide sprays.

103

25%

a 20%

15% b b b b 10%

Mean cadaver infection per arena infection Mean cadaver 5%

0% A B C D E Field

Fig.3.3. Mean cadaver infection with entomogenous fungi per arena in 2015. Total of 8 arenas per sample date by field combination. Field by date interaction (F16, 175 =0.32, Pr > F = 0.9944) and sampling date effect (F4, 175 =1.13, Pr >F = 0.3462) were not significant. Field simple effect was significant (F4, 175 = 4.43, Pr > F = 0.0020). Letters represent means significantly different at P< 0.05 between fields.

104

105

CHAPTER 4: SCREENING OF INSECT-ASSOCIATED FUNGI FROM NEBRASKA AGAINST WESTERN CORN ROOTWORM, DIABROTICA VIRGIFERA VIRGIFERA LECONTE.

Introduction

The western corn rootworm (WCR), Diabrotica virgifera virgifera LeConte

(Coleoptera: Chrysomelidae), is the most damaging belowground pest of continuous corn (Zea mays L.) in North America (Gray et al. 2009) with annual cost estimates of over US $1 billion associated with control and yield loss (Sappington et al. 2006, Gray et al. 2009, Dun et al. 2010, Tinsley et al. 2012, 2015, Andow et al. 2016). Management strategies are mostly focused on controlling the larval stage of the WCR, although adult control can be employed as well to reduce silk clipping or reduce egg laying, which subsequently decreases larval injury in next season’s corn crop (Chandler 2003). Tactics used to control WCR larvae include soil insecticide applications, seed treatments, corn hybrids expressing Bacillus thuringiensis (Bt), and crop rotation. However, there has been a high adaptive rate of the WCR to these tactics when they are used repeatedly in the same location (Gray et al. 2009; Miller et al. 2009). Although these strategies are still effective in many regions of the Corn Belt, some WCR populations have evolved resistance to one or more of the management practices above (Meinke et al. 1998;

Levine et al. 2002; Parimi et al. 2006; Gassmann et al. 2011, 2014, 2016; Wangila et al.

2015; Zukoff et al. 2016; Ludwick et al. 2017).

The increasing incidence of field-evolved resistance to various tactics highlights the need to develop new management strategies against the WCR. New management practices can complement existing tools to mitigate resistance problems and to prolong durability of existing technologies within an integrated pest management framework. In

106 addition, there is also a need to provide alternatives for low input sustainable corn production, as well as popcorn, white corn, seed corn, and organic production, where Bt hybrids and certain insecticides are not an option. The use of entomopathogens as biological control agents can be a sustainable and selective alternative to pesticides

(Lacey et al. 2001; Glare et al. 2012).

Mycoinsecticides, primarily those originating from Beauveria and Metarhizium strains, have been explored in the U.S. and abroad for a wide range of pests (de Faria and Wraight 2007). Entomopathogenic fungi infect through the host’s cuticle, unlike other entomopathogens that need to be ingested to infect the host, and thus, could be considered contact agents. Spores germinate on and penetrate through the insect host’s cuticle, becoming established in the hemolymph, and eventually causing death (Wraight et al. 2007). The fungi Beauveria bassiana (Balsamo) Vuillemin and Metarhizium anisopliae (Metschn.) Sorokin infect all stages of the WCR (Toepfer et al. 2009). The potential for WCR control by EPF alone or in conjunction with other control methods has been demonstrated in lab (Pilz et al. 2007; Meissle et al. 2009; Petzold-Maxwell et al.

2012; Rudeen et al. 2013) and field studies (Krueger and Roberts 1997; Mulock and

Chandler 2000, 2001; Bruck and Lewis 2001, 2002; Pilz et al. 2009; Petzold–Maxwell et al. 2013).

A wide range of entomogenous, or insect-associated, fungi were recovered from

Galleria mellonella L. and Tenebrio molitor L. baiting assays in a survey of five cornfields in Western Nebraska (see Chapter 3). The fungi recovered included the known entomopathogenic genera Beauveria, Metarhizium and Purpureocillium (Paecilomyces),

107 as well as 11 other genera, some of which include species that have exhibited antagonistic properties against insects, e.g., Aspergillus, Chaetomium, Cladosporium, and Fusarium (Hajek et al. 1993; Wicklow et al. 1999; Lage et al. 2001; Gange et al.

2011). Therefore, the objective of this study was to screen representatives of the native entomogenous fungal community described in Chapter 3 for their pathogenicity against

WCR in soil and immersion-exposure assays.

Materials and Methods

Insect sources and rearing. Western corn rootworm eggs or third-instar larvae were obtained from non-diapausing colonies maintained at French Agricultural Research Inc.

(Lamberton, MN) or Crop Characteristics, Inc. (Farmington, MN). Eggs were received in plastic petri dishes (100 X 15 mm, Fisherbrand™, Pittsburgh, PA) containing pre-sifted, autoclaved soil. Petri dish contents were checked for moisture, then sealed with

Parafilm M (Bemis Company, Inc., Neenah, WI) and kept at 25°C until egg hatch.

Neonates were moved to non-transgenic corn seedling mats (Reid’s Yellow Dent) with a soft hair paint brush and allowed to develop to third instar. WCR larvae were then recovered from the soil manually or by placing the seedling mats in Berlese funnels employing 40W, 120V bulbs (Philips Lighting Company, Worcester, MA) for 3 hours.

Larvae were collected in clear glass jars (Solo Cup Company, Lake Forest, IL) attached to the Berlese funnels that contained moist paper towels and corn seedlings. Larvae were immediately transferred to the laboratory and third instars utilized in experiments within 24 hours.

108

Fungal sources and inoculum preparation. The 48 native fungal strains used in this experiment were isolated from soil samples collected in Nebraska cornfields via G. mellonella and T. molitor baiting assays (see Chapter 3). Strains were selected to represent the taxa diversity and origin locations from Chapter 3. In addition,

BotaniGardâ 22WP, being B. bassiana strain GHA (Arbico Organics, Oro Valley, AZ) was included as a commercial comparison product in each assay. Native fungal strains were surface-cultured on full strength potato dextrose agar media supplemented with 1 g L-1 yeast extract (PDAY) media, with pH adjusted to 6.7-6.8 prior to autoclaving (Rangel et al. 2004). PDA media was made with fresh homemade potato broth from Russet potatoes. Spore viability was checked on PDAY 1-2 days prior to bioassays. Viability plates were incubated for 16-18 hours at 26.3 ± 0.5°C and then squares of the agar excised, transferred to a microscope slide and stained with lactophenol cotton blue.

Spore germination was checked under a phase-contrast microscope at 400X magnification. Spores were considered viable if germ tube length was ³ 2x the spore diameter (Inglis et al. 2012). Conidia from 14-day-old cultures were gently scraped with cell scrapers into a small volume of 0.1% Tween 80 (Sigma-AldrichÒ, St. Louis, MO) and the suspensions were then filtered with Miracloth (22-25µL pore size) (Sigma-Aldrich).

Conidia were counted in a hemocytometer and concentrations were adjusted to 1 x 107 viable spores g-1 for soil assays or 1 x 107 viable spores ml-1 for immersion-exposure assays. Sporulation and/or viability was poor for many strains; therefore, it was not always possible to obtain 1 x 107 viable spores ml-1 or g-1. For those strains, the maximum final concentration of viable spores was used. Spore stock suspensions were

109 made up to 24-hours prior to bioassays due to time constraints, but inoculum was prepared the same day as inoculation.

Pathogenicity bioassays against WCR larvae. All 48 strains, BotaniGard and negative controls were initially tested in soil assays against 30 WCR third-instar larvae per treatment. Spore suspensions were mixed thoroughly into 100 grams of autoclaved, pre-sifted (60-mesh) silty clay loam soil at 25% water holding capacity (WHC). This level of soil moisture was chosen to enable good larval survival and good spore germination

(Macdonald and Ellis 1990; Jaronski 2007, Hoffmann et al. 2014). The inoculated soil was dispensed into 59 ml plastic soufflé cups containing three 3-day-old corn seedlings (from planting). Ten third instar WCR larvae were then placed into each cup. The cups were subsequently covered with a fine polyester mesh (No-see-um mesh, Quest Outfitters,

Sarasota, FL) and vented lid to prevent larval escape. This procedure was replicated three times for a total of 30 larvae per strain. Bioassay cups were placed in between 46 x 36 cm cafeteria-trays (Carlisle, Scottsdale, AZ) lined with moistened paper towels and then all were covered with large trash bags to retain original moisture (100% RH)

(Hoffmann et al. 2014). Bioassays were conducted in an incubator at 26.3±0.5°C for 9 days, with 1 ml of water added to each cup on day 5. The number of dead larvae and larvae showing sporulation were recorded at the end of the 9-day bioassay period. In order to confirm mycosis from the tested strains, dead larvae that did not already show external sporulation were placed in a humid chamber at 26.3±0.5°C for another 3-6 days to confirm sporulation. Soil assays were conducted in four batches on separate dates: the first and second each with 15 strains, the third with 5 strains and the fourth

110 with 13 strains. All batches contained BotaniGard (1 x 107 viable spores g-1) as the commercial standard and a negative control (0.1% Tween 80).

Because of the variability in fungal concentrations of the inocula used in the soil assays and in order to confirm pathogenesis, 14 out of the 48 strains were evaluated using immersion-exposure assays with a constant concentration of 1X107 viable spores ml-1 (Pilz et al. 2007). Ten third- instar larvae were placed onto fine polyester mesh cloth

(No-see-um mesh) and then dipped into 5 ml spore suspension for 5 seconds (Pilz et al.

2007). Control larvae were dipped in 0.1% Tween 80. Then, larvae were transferred to a

59-ml cup containing three 3-day old corn seedlings and pre-sifted, autoclaved soil moistened to 25% WHC. This was replicated three times with new inoculum for a total of 30 larvae per strain. Cups were sealed as previously described to prevent larval escape and bioassays were terminated at 7 days.

Data analysis. Proportional mortality was determined as the number of dead larvae per replicate/10 at the end of the bioassay. Larvae were considered dead if they did not move in response to prodding by a toothpick. Fungal growth was considered as positive if at least one infected cadaver showed external fungal growth consistent with gross morphology of fungal strain. All proportion data were fitted to a beta-binomial distribution prior to statistical analysis (Ferrari and Cribari-Neto 2004; Stroup 2015). In the soil assays, a preliminary two-way ANOVA of main effects treatment (fungal strain) and batch and their interaction were analyzed using PROC GLIMMIX in SAS (SAS Institute

Inc., version 9.4, Cary, NC) to evaluate if there was a significant effect of batch on mortality of larvae from the negative control and BotaniGard treatments (n = 120 larvae

111

per treatment). Because there was not a significant batch by treatment interaction (F1, 18

= 0.01, Pr > F = 0.9183), or a significant main effect of batch (F1, 18 = 0.51, Pr > F = 0.4828) or treatment (BotaniGard and Control only) (F1, 18 = 2.71, Pr > F = 0.1579) data were pooled across batches and a one-way ANOVA with PROC GLIMMIX was run to determine the effect of treatment on larval mortality. The immersion-exposure assay was also analyzed with a one-way ANOVA in PROC GLIMMIX with treatment as a fixed factor. For both projects, Dunnett’s multiple mean means comparison was used to test the control and the commercial comparison (BotaniGard) against each strain. Means were obtained using the LSMEANS function with the ILINK option to provide mean estimates, standard errors and confidence limits on the probability scale before the beta-distribution

(Schabenberger 2005). Comparisons were obtained via the DIFF option and adjusted using DUNNETT adjustment for multiple comparisons. Treatment effects and interactions and mean comparisons at P < 0.05 were considered significant.

Results

Soil assays. A significant treatment effect was detected from the one-way ANOVA of soil assays (F49, 118 = 3.75, Pr > F < 0.0001). Fourteen strains caused mortality that was significantly higher than the negative control (E1089 through E1016, Table 4.1). These strains were identified as M. anisopliae (n=2), Metarhizium robertsii J.F. Bisch., Rehner &

Humber (n=11), and Pseudogymnoascus sp. (n=1). Only one strain (E1089, M. anisopliae) caused mortality significantly higher than the commercial standard,

BotaniGard (Table 4.1). External sporulation on larval cadavers was present in 92%

(23/25) of Metarhizium spp. strains. Larval cadavers also showed external sporulation

112 from BotaniGard, E1060 (Cladosporium sp.), E212 (Penicillium griseofulvum Dierckx),

E1035 (Penicillium sp.), E378 (Purpureocilium lilacinum), and E315 (Talaromyces trachyspermus (Shear) Stolk & Samson).

Immersion-exposure assays. The one-way ANOVA also revealed a significant treatment effect in this assay (F15, 32 = 2.78, Pr > F= 0.0074, Table 4.1). Eight strains, including

BotaniGard, caused mortality that was significantly higher than the negative control.

However, the seven field-collected strains were significantly higher than BotaniGard.

These strains were identified as M. anisopliae (n=1), M. robertsii (n=5), and Metarhizium sp. (n=1). BotaniGard (B. bassiana) and 61.5% (8/13) of Metarhizium spp. tested via immersion-exposure assay exhibited sporulation of cadavers (Table 4.1). A trend of higher mean larval mortality, including controls, was observed for the soil assays (41 ±

10%) in comparison to immersion-exposure assays (19 ± 7%). In both experiments, no fungal growth was detected in larval cadavers from the controls.

Discussion

The ability of entomopathogenic fungi to infect the WCR has been tested before

(Krueger and Roberts 1997; Mulock and Chandler 2000, 2001; Bruck and Lewis 2001,

2002; Pilz et al. 2007, 2009; Meissle et al. 2009; Rudeen et al. 2013); however, these studies were restricted to studying pathogenesis and virulence of only Metarhizium and

Beauveria spp. The present study is novel in that it tested a wide range of insect- associated fungi from the soil of cornfields (see Chapter 3) against WCR larvae. One strain of M. anisopliae (E213) and three strains of M. robertsii (E1030, E1056, and

E1016) had mean mortality statistically higher than the control for both assay types.

113

Sporulation of fungi on cadavers confirmed the pathogenic status of 100% and 57% of the Metarhizium strains that caused mortality greater than the control in soil and immersion assays, respectively. Sporulation also indicated that Cladosporium sp.

(E1060), P. griseofulvum (E212), Penicillium sp. (E1035) and T. trachyspermus (E315) are capable of infecting larvae but are weak pathogens since mortality from these strains was comparable to the control.

Mortality significantly higher than the control was found in 52% (13/25) of the

Metarhizium strains tested (Table 4.1). Tested concentrations varied between 1.4 x 105 spores gram-1 to 1 x 107 spores gram-1, but some of the highest mortality (E1000-E1016,

Table 4.1) was obtained from strains with concentrations below 107. High mortality from low-concentrations infers superior virulence of those strains compared to high- concentration strains with the same mortality rates. However, a dose-response bioassay would be needed to confirm this hypothesis. Overall, Metarhizium strains tested in the soil assays caused higher mortality than reported in other studies using the same small- cup methodology (Rudeen et al. 2013; Hoffmann et al. 2014). Two native strains of M. anisopliae from Iowa caused around 10 - 20% corrected mortality at 6.1 x 105 conidia gram-1 and around 30% mortality at 6.1 x 106 conidia gram -1 (Rudeen et al. 2013).

Moreover, mean mortality of WCR larvae was 9% from M. brunneum (F52 strain) inoculations at 104, 105, 106 and 107 spores gram -1 (Hoffmann et al. 2014). It is possible that the isolates collected in Nebraska have superior virulence against WCR larvae, however, variance in soil properties among studies probably played a large role in the mortality differences. Both Rudeen et al. (2013) and Hoffmann et al (2014) used field

114 collected soil while the soil used here was sterile. Sterilization changes the chemical composition and the microbial community within the soil, which can in turn allow rapid colonization of the fungal inoculum (Wilson et al. 1988; Inglis et al. 2012). In this study, it was unknown whether any strains would cause WCR mortality. Therefore, soil sterilization allowed us to isolate the effect of individual fungal strains on WCR larvae.

This is the first report to describe M. robertsii infecting WCR in the literature.

Metarhizium robertsii is part of the M. anisopliae species complex and was just recently described as a new species (Bischoff et al. 2009). The two species are morphologically identical but differ genetically, hence it is possible that other studies have tested M. robertsii strains against the WCR but reported it under M. anisopliae s.l. Metarhizium robertsii is a great target for WCR control because of its multifunctional lifestyle (Barelli et al. 2015). It is rhizosphere competent which means it can survive antimicrobial root exudates and live saprophytically in the absence of a host (Pava-Ripoll 2013).

Metarhizium robertsii also promotes plant health by acting as an entomopathogen, and by colonizing the plant endophytically, it also improves root development and aids in translocating insect-derived nitrogen to roots (Sasan and Bidochka 2012; Behie et al.

2012; Barelli et al. 2016). Moreover, M. robertsii, like other Metarhizium species, is adapted to disturbed environments and is compatible with agroecosystems (Bidochka et al. 2001; Meyling and Eilenberg 2007; Wyrebek et al. 2011; Kepler et al. 2015). An evaluation of the M. robertsii strains from this study in the cornfields may not only benefit WCR management, but also promote plant health.

115

The native B. bassiana (E1040) and BotaniGard (B. bassiana GHA strain) in the soil assay each had low mortality rates that did not differ from control mortality, but

BotaniGard mortality was significantly higher than the control in the immersion exposure assay (Table 4.1). Despite low mortality in the soil assay, BotaniGard infection was confirmed on sporulating cadavers in both assay types meaning that the fungus is able to infect WCR. The WCR mortality from E1040 (21%) is similar to other lab studies that found £ 11% corrected mortality from B. bassiana in laboratory assays (Pilz et al.

2007; Rudeen et al. 2013; Hoffmann et al. 2014). The low larval susceptibility in the lab is consistent with a field study that showed 0 - 3.2% B. bassiana infection at adult emergence (Bruck and Lewis 2001). Moreover, field applications of B. bassiana against adult WCR resulted in inconsistent levels of beetle infection in the field (Bruck and Lewis

2002). The results from this study, together with the above lab and field studies suggest that B. bassiana is not a good mortality agent of WCR. However, B. bassiana can engage in endophytic colonization of plant roots which can increase insect mortality but also promote plant growth (Lopez et al. 2014, Lopez and Sword 2015). Entomopathogenic fungi that are also endophytes have been linked to a variety of plant health roles including disease protection, nutrient acquisition and increased tolerance to abiotic stresses (Bamisile et al. 2018). Testing EPF strains from this paper for endophytic colonization could give insights into their role in the cornfield.

Pseudogymnoascus sp. (E376) caused mortality significantly higher than the control in the soil assay, but they did not show cadaver sporulation. Pseudogymnoascus spp. are widely found in soils, but their relationship to insects has not been studied

116

(Leushkin et al. 2015). Fungi that are not regarded as entomopathogenic in the literature can act as insect antagonists via toxin production or endophytic colonization

(Chapter 3 and references therein). Beyond direct pathogenicity, Pseudogymnoascus sp. can be explored as sources of novel insect resistant genes or other traits that can be beneficial for insect management (Lacey et al. 2015).

Although we cannot make direct mortality comparisons between the assay types, there was a trend of higher mortality in the soil assay versus immersion-exposure assays. It is important to note that WCR larvae in the dipping assay were exposed to the inoculum only for 5 seconds with the bioassay being terminated at 7 days, while the

WCR larvae in the soil assay were exposed to inoculated soil for 9 days. Soil assays are thought to simulate field conditions as larvae are exposed to the fungi in the soil

(Hoffmann et al. 2014).

The definition of pathogenicity is “the potential ability to produce disease” with disease meaning a “departure from the state of health or normality” (Onstad et al.

2006). If the negative control larvae are considered normal, then 17 strains out of the 48 strains are pathogenic to the WCR. But biological control agents require other characteristics such as virulence, environmental competence and persistence, and host specificity to be successful in reducing pest populations (Glazer 1996, Kaya and

Koppenhöfer 1996). Although we cannot directly compare the strains tested in the soil assays because they had varying spore concentrations, the data can provide insight into the potential for using these strains for rootworm control. Under the conditions of this study, M. robertsii, M. anisopliae, Metarhizium sp., Pseudogymnoascus sp., and

117

BotaniGard (commercial comparison) were effective mortality agents of the WCR. These species were also distributed in commercial cornfields in Nebraska (see Chapter 3) but their role in WCR mortality in the field is still unknown. Further studies should be conducted to explore the suitability of strains tested here as biological control agents in the field. Sustainable alternatives for WCR pest management could greatly minimize pest-caused yield losses, management costs, insecticide exposure to the environment and growers, and enhance profitability of corn production in the long term.

118

References:

Andow, D. A., S. G. Pueppke, A. W. Schaafsma, A. J. Gassmann, T. W. Sappington, L. J. Meinke, P. D. Mitchell, T. M. Hurley, R. L. Hellmich, and R. P. Porter. 2016. Early detection and mitigation of resistance to bt maize by western corn rootworm (Coleoptera: Chrysomelidae). J. Econ. Entomol. 109: 1–12.

Bamisile, B. S., C. K. Dash, K. S. Akutse, R. Keppanan, and L. Wang. 2018. Fungal endophytes: beyond herbivore management. Front. Microbiol. 9.

Barelli, L., S. Moonjely, S. W. Behie, and M. J. Bidochka. 2015. Fungi with multifunctional lifestyles: endophytic insect pathogenic fungi. Plant Mol. Biol. 1– 8.

Barelli, L., S. Moonjely, S. W. Behie, and M. J. Bidochka. 2016. Fungi with multifunctional lifestyles: endophytic insect pathogenic fungi. Plant Mol. Biol. 90: 657–664.

Behie, S. W., P. M. Zelisko, and M. J. Bidochka. 2012. Endophytic insect-parasitic fungi translocate nitrogen directly from insects to plants. Science. 336: 1576–1577.

Bidochka, M. J., A. M. Kamp, T. M. Lavender, J. Dekoning, and J. N. A. D. Croos. 2001. Habitat association in two genetic groups of the insect-pathogenic fungus Metarhizium anisopliae: uncovering cryptic species? Appl. Environ. Microbiol. 67: 1335–1342.

Bischoff, J. F., S. A. Rehner, and R. A. Humber. 2009. A multilocus phylogeny of the Metarhizium anisopliae lineage. Mycologia. 101: 512–530.

Bruck, D. J., and L. C. Lewis. 2001. Adult Diabrotica spp. (Coleoptera: Chrysomelidae) infection at emergence with indigenous Beauveria bassiana (Deuteromycotina: Hyphomycetes). J. Invertebr. Pathol. 77: 288–289.

Bruck, D. J., and L. C. Lewis. 2002. Whorl and pollen-shed stage application of Beauveria bassiana for suppression of adult western corn rootworm. Entomol. Exp. Appl. 103: 161–169.

Chandler, L. D. 2003. Corn rootworm areawide management program: United States Department of Agriculture–Agricultural Research Service. Pest Manag. Sci. 59: 605–608.

Dun, Z., P. D. Mitchell, and M. Agosti. 2010. Estimating Diabrotica virgifera virgifera damage functions with field trial data: applying an unbalanced nested error component model. J. Appl. Entomol. 134: 409–419.

119 de Faria, M. R., and S. P. Wraight. 2007. Mycoinsecticides and mycoacaricides: a comprehensive list with worldwide coverage and international classification of formulation types. Biol. Control. 43: 237–256.

Ferrari, S., and F. Cribari-Neto. 2004. Beta regression for modelling rates and proportions. J. Appl. Stat. 31: 799–815.

Gange, A. C., R. Eschen, J. A. Wearn, A. Thawer, and B. C. Sutton. 2011. Differential effects of foliar endophytic fungi on insect herbivores attacking a herbaceous plant. Oecologia. 168: 1023–1031.

Gassmann, A. J., J. L. Petzold-Maxwell, E. H. Clifton, M. W. Dunbar, A. M. Hoffmann, D. A. Ingber, and R. S. Keweshan. 2014. Field-evolved resistance by western corn rootworm to multiple Bacillus thuringiensis toxins in transgenic maize. Proc. Natl. Acad. Sci. 111: 5141–5146.

Gassmann, A. J., J. L. Petzold-Maxwell, R. S. Keweshan, and M. W. Dunbar. 2011. Field- Evolved resistance to Bt maize by western corn rootworm. PLoS ONE. 6: e22629.

Gassmann, A. J., R. B. Shrestha, S. R. K. Jakka, M. W. Dunbar, E. H. Clifton, A. R. Paolino, D. A. Ingber, B. W. French, K. E. Masloski, J. W. Dounda, and C. R. S. Clair. 2016. Evidence of resistance to Cry34/35Ab1 corn by western corn rootworm (Coleoptera: Chrysomelidae): root injury in the field and larval survival in plant-based bioassays. J. Econ. Entomol. 109: 1872-1880

Glare, T., J. Caradus, W. Gelernter, T. Jackson, N. Keyhani, J. Köhl, P. Marrone, L. Morin, and A. Stewart. 2012. Have biopesticides come of age? Trends Biotechnol. 30: 250–258.

Glazer, I. 1996. Survival mechanisms of entomopathogenic nematodes. Biocontrol Sci. Technol. 6: 373–378.

Gray, M. E., T. W. Sappington, N. J. Miller, J. Moeser, and M. O. Bohn. 2009. Adaptation and invasiveness of western corn rootworm: intensifying research on a worsening pest. Annu. Rev. Entomol. 54: 303–321.

Hajek, A. E., P. E. Nelson, R. A. Humber, and J. L. Perry. 1993. Two Fusarium species pathogenic to gypsy moth, Lymantria dispar. Mycologia. 85: 937–940.

Hoffmann, A. M., B. W. French, S. T. Jaronski, and A. J. Gassmann. 2014. Effects of entomopathogens on mortality of western corn rootworm (Coleoptera: Chrysomelidae) and fitness costs of resistance to Cry3Bb1 maize. J. Econ. Entomol. 107: 352–360.

120

Inglis, G. D., J. Enkerli, and M. S. Goettel. 2012. Laboratory techniques used for entomopathogenic fungi: Hypocreales, pp. 189–253. In Man. Tech. Invertebr. Pathol. Second Ed. Elsevier.

Jaronski, S.T. 2007. Soil Ecology of the entomopathogenic Ascomycetes: A critical examination of what we (think) we know. In: Use of entomopathogenic fungi in biologicla pest management, eds: Sunday Ekesi and Nguya K. Maniania.

Kaya, H. K., and A. M. Koppenhöfer. 1996. Effects of microbial and other antagonistic organism and competition on entomopathogenic nematodes. Biocontrol Sci. Technol. 6: 357–372.

Kepler, R. M., T. A. Ugine, J. E. Maul, M. A. Cavigelli, and S. A. Rehner. 2015. Community composition and population genetics of insect pathogenic fungi in the genus Metarhizium from soils of a long-term agricultural research system. Environ. Microbiol. 17: 2791–2804.

Krueger, S. R., and D. W. Roberts. 1997. Soil Treatment with entomopathogenic fungi for corn rootworm (Diabrotica spp.) larval control. Biol. Control. 9: 67–74.

Lacey, L. A., R. Frutos, H. K. Kaya, and P. Vail. 2001. Insect pathogens as biological control agents: do they have a future? Biol. Control. 21: 230–248.

Lacey, L. A., D. Grzywacz, D. I. Shapiro-Ilan, R. Frutos, M. Brownbridge, and M. S. Goettel. 2015. Insect pathogens as biological control agents: Back to the future. J. Invertebr. Pathol. 132: 1–41.

Lage, A. M., G. L. Moraes, and M. Z. Costa. 2001. The entomopathogenic potential of Aspergillus spp. in mosquitoes vectors of tropical diseases. J Basic Microbiol. 41: 45–49.

Leushkin, E. V., M. D. Logacheva, A. A. Penin, R. A. Sutormin, E. S. Gerasimov, G. A. Kochkina, N. E. Ivanushkina, O. V. Vasilenko, A. S. Kondrashov, and S. M. Ozerskaya. 2015. Comparative genome analysis of Pseudogymnoascus spp. reveals primarily clonal evolution with small genome fragments exchanged between lineages. BMC Genomics. 16: 400.

Levine, E., J. L. Spencer, S. A. Isard, D. W. Onstad, and M. E. Gray. 2002. Adaptation of the western corn rootworm to crop rotation: evolution of a new strain in response to a management practice. Am. Entomol. 48: 94–107.

Lopez, D. C., and G. A. Sword. 2015. The endophytic fungal entomopathogens Beauveria bassiana and Purpureocillium lilacinum enhance the growth of cultivated cotton (Gossypium hirsutum) and negatively affect survival of the cotton bollworm (Helicoverpa zea). Biol. Control. 89: 53-60.

121

Lopez, D. C., K. Zhu-Salzman, M. J. Ek-Ramos, and G. A. Sword. 2014. The entomopathogenic fungal endophytes Purpureocillium lilacinum (formerly Paecilomyces lilacinus) and Beauveria bassiana negatively affect cotton aphid reproduction under both greenhouse and field conditions. PloS One. 9: e103891.

Ludwick, D. C., L. N. Meihls, K. R. Ostlie, B. D. Potter, L. French, and B. E. Hibbard. 2017. Minnesota field population of western corn rootworm (Coleoptera: Chrysomelidae) shows incomplete resistance to Cry34Ab1/Cry35Ab1 and Cry3Bb1. J. Appl. Entomol. 141: 28–40.

Macdonald, P. J., and C. R. Ellis. 1990. Survival time of unfed, first-instar western corn rootworm (Coleoptera: Chrysomelidae) and the Effects of soil type, moisture, and compaction on their mobility in soil. Environ. Entomol. 19: 666–671.

Meinke, L. J., B. D. Siegfried, R. J. Wright, and L. D. Chandler. 1998. Adult susceptibility of Nebraska western corn rootworm (Coleoptera: Chrysomelidae) populations to selected insecticides. J. Econ. Entomol. 91: 594–600.

Meissle, M., C. Pilz, and J. Romeis. 2009. Susceptibility of Diabrotica virgifera virgifera (Coleoptera: Chrysomelidae) to the entomopathogenic fungus Metarhizium anisopliae when feeding on Bacillus thuringiensis Cry3Bb1-expressing maize. Appl. Environ. Microbiol. 75: 3937–3943.

Meyling, N. V., and J. Eilenberg. 2007. Ecology of the entomopathogenic fungi Beauveria bassiana and Metarhizium anisopliae in temperate agroecosystems: Potential for conservation biological control. Biol. Control. 43: 145–155.

Miller, N. J., T. Guillemaud, R. Giordano, B. D. Siegfried, M. E. Gray, L. J. Meinke, and T. W. Sappington. 2009. Genes, gene flow and adaptation of Diabrotica virgifera virgifera. Agric. For. Entomol. 11: 47–60.

Mulock, B., and L. Chandler. 2000. Field-cage studies of Beauveria bassiana (Hyphomycetes: Moniliaceae) for the suppression of adult western corn rootworm, Diabrotica virgifera virgifera (Coleoptera: Chrysomelidae). Biocontrol Sci. Technol. 10: 51–60.

Mulock, B. S., and L. D. Chandler. 2001. Effect of Beauveria bassiana on the fecundity of western corn rootworm, Diabrotica virgifera virgifera (Coleoptera: Chrysomelidae). Biol. Control. 22: 16–21.

Onstad, D. W., J. R. Fuxa, R. A. Humber, J. Oestergaard, D. I. Shapiro-Ilan, V. V. Gouli, R. S. Anderson, T. G. Andreadis, and L. A. Lacey. 2006. Abridged glossary of terms used in invertebrate pathology, 3rd edn. Society for Invertebrate Pathology.

122

Parimi, S., L. J. Meinke, B. Wade French, L. D. Chandler, and B. D. Siegfried. 2006. Stability and persistence of aldrin and methyl-parathion resistance in western corn rootworm populations (Coleoptera: Chrysomelidae). Crop Prot. 25: 269– 274.

Pava-Ripoll, M. 2013. Metarhizium robertsii, a rhizosphere-competent insect pathogen, pp. 149–159. In Bruijn, F.J. de (ed.), Mol. Microb. Ecol. Rhizosphere. John Wiley & Sons, Inc.

Petzold–Maxwell, J. L., S. T. Jaronski, E. H. Clifton, M. W. Dunbar, M. A. Jackson, and A. J. Gassmann. 2013. Interactions among Bt Maize, entomopathogens, and rootworm species (Coleoptera: Chrysomelidae) in the field: effects on survival, yield, and root injury. J. Econ. Entomol. 106: 622–632.

Petzold-Maxwell, J. L., S. T. Jaronski, and A. J. Gassmann. 2012. Tritrophic interactions among Bt maize, an insect pest and entomopathogens: effects on development and survival of western corn rootworm. Ann. Appl. Biol. 160: 43–55.

Pilz, C., S. Keller, U. Kuhlmann, and S. Toepfer. 2009. Comparative efficacy assessment of fungi, nematodes and insecticides to control western corn rootworm larvae in maize. BioControl. 54: 671–684.

Pilz, C., R. Wegensteiner, and S. Keller. 2007. Selection of entomopathogenic fungi for the control of the western corn rootworm Diabrotica virgifera virgifera. J. Appl. Entomol. 131: 426–431.

Rangel, D. E. N., G. U. L. Braga, S. D. Flint, A. J. Anderson, and D. W. Roberts. 2004. Variations in UV-B tolerance and germination speed of Metarhizium anisopliae conidia produced on insects and artificial substrates. J. Invertebr. Pathol. 87: 77– 83.

Rudeen, M. L., S. T. Jaronski, J. L. Petzold-Maxwell, and A. J. Gassmann. 2013. Entomopathogenic fungi in cornfields and their potential to manage larval western corn rootworm Diabrotica virgifera virgifera. J. Invertebr. Pathol. 114: 329–332.

Sappington, T. W., B. D. Siegfried, and T. Guillemaud. 2006. Coordinated Diabrotica genetics research: accelerating progress on an urgent insect pest problem. Am. Entomol. 52: 90.

Sasan, R. K., and M. J. Bidochka. 2012. The insect-pathogenic fungus Metarhizium robertsii (Clavicipitaceae) is also an endophyte that stimulates plant root development. Am. J. Bot. 99: 101–107.

Schabenberger, O. 2005. Introducing the GLIMMIX procedure for generalized linear mixed models. SUGI 30 Proc. 196.

123

Stroup, W. W. 2015. Rethinking the analysis of non-normal data in plant and soil science. Agron. J. 107: 811–827.

Tinsley N. A., Estes R. E., and Gray M. E. 2012. Validation of a nested error component model to estimate damage caused by corn rootworm larvae. J. Appl. Entomol. 137: 161–169.

Tinsley, N. A., P. D. Mitchell, R. J. Wright, L. J. Meinke, R. E. Estes, and M. E. Gray. 2015. Estimation of efficacy functions for products used to manage corn rootworm larval injury. J. Appl. Entomol. 140: 414–425.

Toepfer, S., T. Haye, M. Erlandson, M. Goettel, J. G. Lundgren, R. G. Kleespies, D. C. Weber, G. C. Walsh, A. Peters, R.-U. Ehlers, H. Strasser, D. Moore, S. Keller, S. Vidal, and U. Kuhlmann. 2009. A review of the natural enemies of beetles in the subtribe Diabroticina (Coleoptera: Chrysomelidae): implications for sustainable pest management. Biocontrol Sci. Technol. 19: 1–65.

Wangila, D. S., A. J. Gassmann, J. L. Petzold-Maxwell, B. W. French, and L. J. Meinke. 2015. Susceptibility of Nebraska western corn rootworm (Coleoptera: Chrysomelidae) populations to Bt corn events. J. Econ. Entomol. 108: 742–751.

Wicklow, D. T., P. F. Dowd, and J. B. Gloer. 1999. Chaetomium mycotoxins with antiinsectan or antifungal Activity. Mycotoxins. 1999: 267–271.

Wilson, G. W. T., B. A. D. Hetrick, and D. G. Kitt. 1988. Suppression of mycorrhizal growth response of big bluestem by non-sterile soil. Mycologia. 80: 338–343.

Wraight, S.P., G.D. Inglis, and M.S. Goettel. 2007. Fungi. Lacey, L. A., and Kaya, H. K. (Eds.). Field manual of techniques in invertebrate pathology. Dordrecht etc.: Kluwer acad. publ. Cop. Pp.223-249

Wyrebek, M., C. Huber, R. K. Sasan, and M. J. Bidochka. 2011. Three sympatrically occurring species of Metarhizium show plant rhizosphere specificity. Microbiology. 157: 2904–2911.

Zukoff, S. N., K. R. Ostlie, B. Potter, L. N. Meihls, A. L. Zukoff, L. French, M. R. Ellersieck, B. W. French, and B. E. Hibbard. 2016. Multiple assays indicate varying levels of cross resistance in Cry3Bb1-selected field populations of the western corn rootworm to mCry3A, eCry3.1Ab, and Cry34/35Ab1. J. Econ. Entomol. 109: 1387- 1398.

Table

Table 4.1 Mean mortality of western corn rootworm larvae in soil and immersion-exposure assays with entomopathogenic fungi collected from cornfields in Western Nebraska.

a b Soil Assay Immersion-exposure Assay Mortality ± Fungal Mortality ± Strain Species Spores gram -1 SEM (%) growth SEM (%) Fungal growth E1089 Metarhizium anisopliae 1.0 x 107 75 ± 9** Y 7 ± 4 Y E1000 Metarhizium robertsii 5.8 x 105 70 ± 10* Y 18 ± 7 N E645 Metarhizium robertsii 2.3 x 106 68 ± 10* Y - - E1026 Metarhizium robertsii 1.1 x 106 65 ± 10* Y 9 ± 5 Y E653 Metarhizium robertsii 8.7 x 105 61 ± 11* Y - - E138 Metarhizium robertsii 7.1 x 106 61 ± 11* Y - - E1022 Metarhizium robertsii 8.3 x 106 60 ± 11* Y 7 ± 4 Y E1030 Metarhizium robertsii 1.9 x 106 59 ± 11* Y 30 ± 9* Y E380 Metarhizium robertsii 4.2 x 106 58 ± 11 * Y 18 ±7 Y E328 Metarhizium robertsii 5.6 x 106 56 ± 11 * Y 7 ± 4 N E376 Pseudogymnoascus sp. 1.0 x 107 56 ± 11* N 7 ± 4 N E1056 Metarhizium robertsii 2.7 x 106 53 ± 11* Y 22 ± 8* N E213 Metarhizium anisopliae 3.3 x 106 50 ± 11* Y 26 ± 9* N E1016 Metarhizium robertsii 2.5 x 106 50 ± 11* Y 38 ± 10* Y E161 Metarhizium robertsii 8.3 x 106 47 ± 11 Y 25 ± 8* Y E1038 Metarhizium sp. 4.4 x 106 46 ± 11 Y 30 ± 9* Y E312 Chaetomium sp. 2.7 x 105 43 ± 11 N - - E1093 Metarhizium robertsii 1.4 x 105 43 ± 11 Y 28 ± 9* N

124

Table 4.1 (Continued)

a b Soil Assay Immersion-exposure Assay

Mortality ± Fungal Mortality ± Strain Species Spores gram -1 SEM (%) growth SEM (%) Fungal growth E136 Metarhizium robertsii 6.6 x 106 41 ± 11 Y - - E1033 Metarhizium anisopliae 7.5 x 106 36 ± 11 Y - - E1095 Metarhizium sp. 7.1 x 105 35 ± 10 Y - - E378 Purpureocillium lilacinum 1.0 x 107 35 ± 10 Y - - E648 Fusarium oxysporum 1.0 x 107 30 ± 10 N - - E211 Metarhizium robertsii 4.7 x 106 30 ± 10 N - - E172 Penicillium janthinellum 9.6 x 105 30 ± 10 N - - BotaniGard Beauveria bassiana 1.0 x 107 29 ± 5 Y 32 ± 10* Y E1090 Metarhizium robertsii 4.8 x 106 28 ± 10 Y - - E212 Penicillium griseofulvum 1.0 x 107 28 ± 10 Y - - E166 Penicillium sp. 1.0 x 107 28 ± 10 N - - E393 Pseudogymnoascus pannorum 1.0 x 107 27 ± 9 N - - E1035 Penicillium sp. 1.0 x 107 26 ± 9 Y - - E1005 Metarhizium robertsii 1.0 x 107 24 ± 9 Y - - E314 Neosartorya sp. 1.0 x 107 24 ± 9 N - - E1040 Beauveria bassiana 1.0 x 107 21 ± 8 N - - E322 Metarhizium robertsii 5.8 x 106 21 ± 8 N - - E374 Metarhizium robertsii 1.0 x 107 21 ± 8 Y - - E646 Talaromyces pinophilus 1.0 x 107 21 ± 8 N - - E1060 Cladosporium sp. 6.7 x 106 20 ± 8 Y - - E368 Penicillium bilaiae 1.0 x 107 20 ± 8 N - -

125

Table 4.1 (Continued)

a b Soil Assay Immersion-exposure Assay

Mortality ± Fungal Mortality ± Strain Species Spores gram -1 SEM (%) growth SEM (%) Fungal growth E651 Clonostachys sp. 9.6 x 106 19 ± 8 N - - E315 Talaromyces trachyspermus 5.8 x 105 19 ± 8 Y - - E331 Talaromyces trachyspermus 7.7 x 106 16 ± 7 - - - Control 0.1 % Tween 80 - 14 ± 3 - 2 ± 1 - E390 Talaromyces sp. 1.0 x 107 8 ± 4 N - - E1034 Metarhizium anisopliae 2.7 x 106 8 ± 4 Y - - E999 Fusarium solani 6.9 x 106 7 ± 4 N - - Pseudogymnoascus E370 destructans 1.2 x 106 7 ± 4 N - - E126 Cladosporium halotolerans 4.2 x 106 7 ± 4 N - - E998 Aspergillus flavus 1.0 x 107 6 ± 3 - - - E325 Taifanglania sp. 1.0 x 107 6 ± 3 - - - a: Means with (*) are significantly different from the control and means with (**) are significantly different than the commercial standard (BotaniGard) at P< 0.05 with Dunnet’s adjustment for multiple pairwise comparisons. b: Immersion- exposure assays: All conducted with 1.0 x 107 spores ml -1.

126

127

CHAPTER 5: IDENTITY AND SEASONAL PATTERNS OF NEMATODES ASSOCIATED WITH INSECT CADAVERS FROM CORNFIELDS IN NEBRASKA.

Introduction

Entomopathogenic nematodes (EPN) are common components of insects’ trophic interactions and can function as biocontrol agents of soil insects (Strong et al.

1999; Jackson et al. 2000). These nematodes are distributed globally and show great biological control potential for insect pests (Hominick et al. 1996; Adams et al. 2006).

Members of the Steinernematidae Travassos and Heterorhabditidae Poinar families are obligate insect pathogens whose third-stage infective juveniles (IJs) kill insect hosts through the release of symbiotic bacteria: Photorhabdus for Heterorhabditidae and

Xenorhabdus for Steinernematidae (Koppenhöfer 2007). Steinernematidae contains 2 genera: Neosteinernema with one species and Steinernema with 70 species;

Heterorhabditidae contains a single genus: Heterorhabditis with 20 species (Stock and

Goodrich-Blair 2012).

Nematodes are identified primarily via morphological, biochemical and molecular tools (Seesao et al. 2017). Steinernema and Heterorhabditis species identification based on traditional morphological and morphometric analyses is time consuming and requires considerable expertise (Stock 2015). However, molecular approaches have become standard alternatives or complements to morphological identification. In particular, DNA barcoding approaches can be used for species identification and to infer phylogenetic relationships of unknown taxa to known taxa using small genomic sequences from individual nematodes (Bhadury et al. 2006, Powers

2004). Two ribosomal regions are frequently used for EPNs studies: internal transcriber

126

128 spacer (ITS1, 5.8S and ITS2) and 28S rDNA (D2/D3 expansion region) (Stock et al. 2001,

Spiridonov et al. 2004, Stock 2015). Sequences from these markers can be extremely variable which can prevent them from being used as a universal nematode marker needed for DNA barcoding approaches (Prosser et al. 2013). The mitochondrial gene cytochrome oxidase subunit 1 (CO1) is considered to be a universal barcode for the animal kingdom as it translates into conserved proteins which provide the right specificity to differentiate species and strains (Hebert et al. 2003, Powers 2004, Prosser et al. 2013). Beyond providing biodiversity data, proper EPN identification can aid in biological control efforts. Identifying native persistent EPN strains that can be augmented in the laboratory and inoculated into agricultural fields can help reduce populations of soil pests (Shields et al. 1999; Shields 2015; Shields and Testa 2015)

In general, at the local scale, EPN species exhibit a patchy horizontal and vertical distribution in the soil (Stuart et al. 2006). Several factors influence EPN distribution and seasonality which include host availability, stress tolerance, soil conditions

(temperature, moisture, texture), agricultural management intensity, ecosystem type, and intra- and inter- specific resource allocation (Kung et al. 1991, Cabanillas and

Raulston 1994, Stuart and Gaugler 1994, Glazer 1996, Campbell et al. 1998, Efron et al.

2001, Grewal et al. 2002, Millar and Barbercheck 2002, Spiridonov et al. 2007, Campos-

Herrera et al. 2008, 2013, Salame and Glazer 2015, Stuart et al. 2015). Multiple EPN species can co-exist in the environment, but they must display different foraging behaviors, niche partitioning and dispersal strategies to avoid competition (Seesao et al.

2017).

126

129

Since EPN species can co-exist, studies have tested co-inoculation of two or more

EPN species for pest suppression with mixed success (Choo et al. 1996, Koppenhöfer et al. 2000, Neumann and Shields 2008, Shields 2015). Successful inoculations occur with the right combination of EPN species that can cover different soil profiles (Shields 2015).

Long term EPN persistence can also occur with the introduction of multiple EPN species in crop-rotation systems (Shields 2015, Shields and Testa 2015). Several Steinernema and Heterorhabditis species have been tested against the western corn rootworm

(WCR) (Diabrotica virgifera virgifera LeConte) in laboratory and field settings (Geisert et al. 2018, Wright et al. 1993, Journey and Ostlie 2000, Toepfer et al. 2005, 2008, Kurtz et al. 2009). However, Steinernema feltiae Filipjev, Steinernema carpocapsae Weiser and

Heterorhabditis bacteriophora Poinar are the species mostly studied under field conditions for WCR control. Field studies have indicated that EPN applications can be just as efficacious as insecticide treatments in killing WCR larvae and providing crop root protection (Wright et al. 1993, Jackson 1996, Toepfer et al. 2008, Pilz et al. 2009).

In an effort to understand how EPNs can be used for WCR management in the continuous corn system in Nebraska, we conducted a study with three objectives: 1) conduct a survey for native EPN that are adapted to irrigated cornfields in western

Nebraska; 2) determine efficacy, survival and seasonal distribution of inoculated EPNs in a cornfield artificially infested with WCR; and 3) determine the identity of EPNs from the previous objectives using a DNA barcoding approach.

126

130

Materials and Methods

Survey Project

Field sites. Five continuous cornfield sites were the same as described in Chapters 2 and

3. The details on field history, location, hybrids, transgenic traits, and insecticide and fungicide use are listed in Appendix Table 1.

Nematode detection from survey sites. Soil sampling and baiting assay procedures were the same as described in Chapter 3. In 2014, five Galleria mellonella (L.) larvae were placed into the baiting assays and in 2015, three G. mellonella larvae and three

Tenebrio molitor L. larvae were used. Dead larvae were placed onto white traps (White

1927) to allow for nematode isolation and detection of infective juveniles. White traps consist of cadavers being placed on a small petri-dish (60 x 15mm) lined with moist filter paper placed inside a larger (100 x 15 mm) “harvest” dish filled with water. Nematodes were then isolated in sterile water in 1.5 ml microcentrifuge tubes (Fisher Scientific™,

Lenexa, KS) and stored at 4 °C until DNA extraction.

Inoculation Project

Field Site. This field site was located at the University of Nebraska West Central Water

Resources Laboratory near Brule, NE (GPS coordinates at center of plots: N41.09.482’,

W102.01.452’). The site was first year corn, soybeans Glycine max (L.) Merr were planted in 2014. The corn hybrid DeKalbâ 52-61 VT Double Proâ that does not express rootworm-active traits (did express Lepidoptera-specific Cry1A.105 and Cry2Ab2 proteins) was planted at 32,000 seeds/acre and 76.2 cm row spacing on May.18.2015.

This was a no-tillage field with no at-plant insecticide applications. The field was

126

131 maintained with typical agronomic practices for irrigated cornfields in the region including fertilizer, herbicides and fungicides. Each plot measured 24.4 m (32 rows) x

24.4 m. There was a total of 36 plots arranged into a randomized complete block design of 6 blocks (Fig. 5.1). This site was planted to soybeans in 2016.

Western corn rootworm infestation and nematode inoculation. Western corn rootworm egg infestation occurred on June.18.2015 when corn plants were at the V1-

V2 growth stage (Ritchie et al. 1992). Eggs from French Agricultural Research, Inc.

(Lamberton, MN) were suspended in 0.15% agar solution and applied with a syringe at a depth of 10 cm in a single furrow adjacent to the plant along the planted row (Sutter and Branson 1980). The infestation zone within each plot consisted of 42 corn plants from the middle four corn rows (2.5 x 2.5 m) (Fig. 5.1). Each plant received 400 eggs; this infestation rate corresponded to the maximum infestation rate before density- dependent mortality often occurs (Hibbard et al. 2010).

EPN inoculation occurred in the evening of July.8.2015 when corn plants were at the V3-V5 growth stage and WCR larvae were in first and second instars. The three treatments evaluated were: 1) commercial EPN strains NemAttackä (Steinernema feltiae) and NemaSeekä (Heterorhabditis bacteriophora) (ARBICO Organics, Tucson, AZ);

2) persistent EPN strains from New York state of S. feltiae “NY 04” strain and H. bacteriophora “Oswego” strain, herein referred as persistent EPNs (Dr. Elson Shields,

Cornell University, Ithaca, NY); and 3) non-inoculated, control plots that received water only. Nematodes were applied at a rate of 2.5 x 109 IJ/ha total, representing 1.25 x 109

IJ/ha/species. Hence, each plot (12.5 m2) received approximately 3.12 x 106 IJs total

126

132

(1.56 x 106 IJs/species). Commercial EPN products were diluted in batches of four liters of non-chlorinated water and applied over the WCR infestation zone (42 plants) with a watering can. Persistent nematodes were applied via Galleria mellonella cadavers, with the calculation that each cadaver releases approximately 100,000 infective juveniles

(Shapiro-Ilan et al. 2003, Dolinski et al. 2007, 2015). Cadavers were buried at a 5 cm depth adjacent to corn roots every 122 cm, alternating the cadavers of each species to allow for an even distribution, for a total of 32 cadavers per plot (16 of each nematode species) or four cadavers/row followed by eight liters of water over the treatment area

(Fig 5.1).

Western corn rootworm population and damage assessments. Western corn rootworm abundance was monitored via single-plant beetle emergence cages (Pierce and Gray

2007). Each plot received 3 cages that were monitored weekly from August.5.2015 until

September.24.2015 for a total of eight collection dates. Larval feeding damage assessment was measured on August.12.2015 via the Iowa State Node Injury Scale

(Oleson et al. 2005) on five plants/plot. This scale ranges from a 0.00 - 3.00 rating; 0 = no feeding damage; 1 = one node, or the equivalent of an entire node of roots pruned by larval feeding to ≤ 3.8 cm from the stalk (Oleson et al. 2005); 2 = two nodes pruned; 3 = three or more complete nodes pruned.

Nematode detection and isolation from soil samples. Soil sampling for nematodes occurred at seven dates: 7 days pre-inoculation, 7 days post-inoculation (dpi), then at

14, 30, 60, 90 dpi and one-year post-inoculation. In an effort to prevent cross- contamination, controls were always sampled first, and different personnel groups

126

133 sampled the commercial and persistent treatments. Soil sampling was obtained from the top 15 cm of the corn root zone with a 2.2 cm x 83.8 cm soil probe (AMS Inc.,

American Falls, ID). Soil probes were rinsed with water and then sterilized with 70% ethanol between plots. In each block, ten soil samples were obtained per date per treatment. For each sample, the top 5 cm of soil (0-5 cm) was separated from the bottom 10 cm (5-15 cm) into two deli dishes (226.8 ml clear hinged deli container with high dome lid Genpak, Charlotte, NC). Soil cores were broken down with sterile forks prior to receiving G. mellonella. Three G. mellonella were placed in dishes with 0-5 cm soil samples and six larvae were placed in dishes containing 5-15 cm soil samples

(Shields et al. 1999). Deli dishes were kept in the dark and incubated at 22-23°C for 7 days. Larval cadavers were then placed onto white traps or dissected to confirm EPN infection. Isolated IJ from white traps were kept at 4 °C inside 1.5 ml microcentrifuge tubes filled with sterile water until DNA extraction.

Nematode Identification: DNA barcoding. Single nematodes in 1.5 ml microcentrifuge tubes were placed onto glass coverslips in 18 µl of sterile water. DNA extractions consisted of individual nematodes being macerated with a micropipette tip (Powers et al. 2014). Mashed nematodes in water were then stored at -20°C in 0.25 ml PCR reaction tubes until PCR was conducted. This study used a partial mitochondrial cytochrome oxidase subunit I (COI) gene primer set: COI-F1KF (29bp, 5’-

CCTACTATGATTGGTGGTTTTGGTAATTG-3’) and COI-R2KF (23bp, 5’-

GTAGCAGCAGTAAAATAAGCACG-3’) (Kanzaki and Futai 2002). Excluding the primers, amplification products yielded 658-bp for sequence analysis. PCR amplification reactions

126

134 consisted of 6.4 µl of ddH2O, 1.8 µl of each 20 µM primer, 15 µl of 2XJumpStart RED Taq

ReadyMix (Sigma-Aldrich, Inc. St. Louis, MO) and 5 µl DNA template from the macerated nematode, for a total reaction volume of 30 µl. PCR cycling protocol began with a hot- start and an initial denaturation of 5 minutes at 94°C followed by 45 cycles of 15 seconds at 94°C (denaturation), 15 seconds at 55°C (annealing), 60 seconds at 72°C

(extension) and a final extension step of 5 minutes at 72°C. All PCR was conducted in a thermal cycler (Techne Equipment, Staffordshire, UK). To confirm successful amplifications, 3 µl of PCR products were loaded into 1% agarose gels stained with ethidium bromide or GelRedä Nucleic Acid Gel Stain (Biotium, Hayward, CA, USA) in 1×

Tris-acetate-EDTA (TAE) buffer. Gels were placed into electrophoresis with 0.5X Tris-

Borate-EDTA (TBE) running buffer for 35 minutes at 155V. UV visualized gel images were digitally recorded. Positive PCR reactions were purified with a Gel/PCR DNA Fragment

Extraction Kit (IBI Scientific. Dubuque, IA) following the manufacturer’s guidelines. DNA templates were sequenced in both directions by the UCDNA Sequencing Facility at the

University of California – Davis. Sequences were edited and aligned on CodonCode

Aligner Version 4.2 (CodonCode Corp, Centerville, Massachusetts).

Reference nematode specimens. To provide standards for identification of the isolated nematodes, we obtained a set of nematode strains from Dr. David Shapiro-Ilan (USDA-

ARS-SE Fruit and Tree Nut Research Unit, Byron, GA). The isolates used were

Heterorhabditis bacteriophora (VS strain), H. bacteriophora (Oswego strain), H. bacteriophora (HB strain), H. megidis UK211, H. georgiana (Kesha strain), H. floridensis

(K22 strain), Steinernema carpocapsae (All strain), S. carpocapsae (Cxrd strain), S. feltiae

126

135

(SN strain) and S. rarum (17C&E). Live nematodes were received in 250 ml cell culture flasks with vent caps (Corning Incorporated, Corning, NY) in sterile water. These reference nematodes were subjected to the same procedures outlined in the

“Nematode identification: DNA barcoding” section.

Nematode Identification/phylogenetic analysis: Edited field-collected nematode COI sequences were compared to sequences in the National Center for Biotechnology

Information (NCBI) database via Basic Local Alignment Search Tool (BLAST) and sequences from USDA reference species. A Neighbor-joining phylogenetic analysis was also conducted with MEGA6: Molecular Evolutionary Genetics Analysis Version 6.0.6

(Tamura et al. 2013) using Kimura-2-Parameter model, 2,000 bootstrap replications, and treated with pairwise deletion gap treatment. Seventy nematode specimens were used in the construction of a neighbor-joining tree. Fifteen nematode specimens originated from the survey project, twenty from the inoculation project, twenty from USDA reference specimens (representing 2 nematodes for each of the 10 EPN strains) and fifteen nematode COI sequences from GenBank.

Data analyses. Survey Project: Soil samples were considered positive for nematode infection if at least one G. mellonella or T. molitor cadaver detected nematodes.

Detection frequency was expressed as proportional number of nematode-infected cadavers per soil sample for each date in all fields. Statistical analyses were performed using generalized linear mixed models with PROC GLIMMIX in SAS (SAS Institute Inc., version 9.4, Cary, NC). Two-way analysis of variance (ANOVA) was used to test for the effect of field site, date and their interaction on detection frequency for 2014 and 2015.

126

136

One-way ANOVA was conducted for 2015 data to determine if bait insect species (G. mellonella or T. molitor) had an impact on nematode detection. Prior to all analyses, all data was converted to beta-binomial distribution (Ferrari and Cribari-Neto 2004, Stroup

2015). Means, standard errors and confidence limits were converted back to the data proportional scale using the ILINK option in LSMEANS (Schabenberger 2005). Multiple comparisons were adjusted with Tukey’s honestly significant difference (HSD) test using the ADJUST option in LSMEANS.

Inoculation project: Nematode detection frequency was expressed as the proportion of infected cadavers at each depth per treatment per block. A three-way ANOVA under generalized linear mixed model was conducted using PROC GLIMMIX in SAS to evaluate the effect of treatment, soil sample depth, date and all interactions on nematode infection rate. Dates were treated as repeated measures with first order autoregressive covariance structure (AR-1). Root injury rates were converted to proportional data by dividing every rating by three (node injury scale 0-3, Oleson et al. 2005). Proportional root injury rates were analyzed in a one-way ANOVA of main effects of EPN treatment.

Nematode detection and root injury rates were fitted to a beta binomial distribution and proportional data were converted back to the data scale using the ILINK option in

LSMEANS. Emergence cage data were analyzed on PROC GLIMMIX using a one-way

ANOVA of main effects of EPN treatment. Emergence cage data was fitted to a negative binomial distribution (Tripathi 2006). Treatment effects and interactions at P< 0.05 were considered significant for both projects.

126

137

Results

Survey Project. Nematodes were detected in 15.8% (54/342) of soil samples in 2014 with only 2.3% (8/342) coming from non-irrigated areas. In 2015, 13.2% (26/197) of soil samples were positive for nematodes. In 2014, 5.5% (91/1640) of larvae were infected and in 2015, 3.7% (44/1179) of larvae were infected. There was no significant interaction of field by date or main effects of proportional nematode detection per arena in 2014 or 2015 (Fig. 5.2, Table 5.1). No significant differences in proportional nematode detection between baiting insects (G. mellonella vs. T. molitor) were detected in 2015 (F1,48 = 2.01, Pr> F = 0.1624).

Inoculation Project. Across treatments, only 8% of all G. mellonella larvae (912/11340) detected nematodes; from those, 467/3779 (12.4%) came from the top 0- 5 cm soil samples and 445/7561 (5.9%) came from the bottom 5-15 cm soil samples. Nematode detection frequency in pre-inoculation sampling was 0.12% (2/1620). Overall, nematode detection frequency was numerically highest for the persistent treatment (10.2%,

386/3780), followed by the commercial (7.7%, 290/3780) and control treatments (6.2%,

236/3780). Three-way analysis of variance between treatment, soil sample depth and date revealed significant two-way interactions for treatment by date and depth by date were observed (Table 5.2). No other two-way or three-way interactions were significant, but a main effect of date was observed (Table 5.2). Multiple comparisons of interaction least square means between treatment and date, and depth by date are listed in tables

5.3 and 5.4, respectively. The treatment by date interaction was primarily driven by treatment differences at 90 dpi (Fig. 5.3). At this date, the persistent treatment was

126

138 significantly higher than the control. Mean differences across the collection period for the control treatment revealed that nematode detection at 7 and 14 dpi was significantly higher than the pre-inoculation sampling and at 90 dpi and 1 yr sampling dates (Fig 5.3). For the depth by date interaction, mean nematode detection frequency per plot was significantly higher in the 0-5 cm layer at 14 dpi than all others (Fig. 5.4).

Western corn rootworm emergence was detected from August.5.2015 until

September.24.2015. Mean cumulative WCR emergence per plant was not statistically different among treatments; 8.4 ± 1.15 for the control treatment, 6.2 ± 0.81 for the commercial treatment, and 9.7 ± 1.4 for persistent treatment (F2,105 = 2.72, Pr > F =

0.0706) (Fig. 5.6). Root injury ratings were also not significantly different between treatments (F2, 177 = 1.38, Pr> F = 0.2534), with mean values (on a 0.00 - 3.00 scale) of,

0.028 ± 0.004, 0.021 ± 0.003 and 0.025 ± 0.004 for the control, commercial and persistent treatments, respectively.

DNA Barcoding. A full description of all specimens added in the neighbor joining tree can be found in Table 5.5. The neighbor-joining analysis produced eight haplotype groups of EPNs, all strongly supported by bootstrap values of 100 (Figure 5.7). The EPN haplotype groups were structured within two clades, one representing Heterorhabditis species and the second comprised of Steinernema species. In the Heterorhabditis clade there were four haplotype groups. Haplotype group 1 consisted of three H. bacteriophora USDA references strains, the H. georgiana USDA reference strain, and a separate and distinct subgroup of specimens isolated from the survey fields. Haplotype group 2 in the Heterorhabditis clade consisted of two Genbank specimens identified as

126

139

H. bacteriophora, and three specimens from the inoculation plots that were identical to

EF043402.1 from GenBank. The third Heterorhabditis haplotype group (#3) consisted of

H. floridensis USDA reference strain and haplotype group 4 consisted of H. megidis

UK211 USDA reference strain and a GenBank sequence representing H. megidis.

Haplotype groups 5 through 8 belonged to the Steinernema clade. Haplotype group 5 was divided into two subgroups and consisted of four USDA reference strains and three GenBank sequences of S. carpocapsae. All but two of these strains (N6797 and N6798) represented different haplotypes. Haplotype group 6 consisted of S. rarum

USDA reference strain. Haplotype group 7 contained Steinernema sp. from the inoculation and survey projects and did not match any reference or GenBank sequences.

Haplotype group eight consisted of S. feltiae strains from the control plots in the inoculation project, USDA reference (SN strain) and GenBank sequences from S. feltiae.

A third clade was largely represented by specimens in the family Diplogasteridae isolated from the survey and inoculation projects. An additional four specimens had no close match in GenBank, although one specimen (N6691) was supported by a bootstrap value of 100 with a GenBank sequence from the genus Oscheius.

Discussion

The nematode detection frequency rates from insect cadavers: 5.5% (2014) and

3.7% (2015) and percentage of positive soil samples: 16% (2014) and 13.2% (2015) fit within the range reported by other studies in cultivated fields (Cabanillas and Raulston

1994, Liu and Berry 1995, Garcia del Pino and Palomo 1996, de Brida et al. 2017) and other ecosystems (Hara et al. 1991, Campbell et al. 1996, Glazer et al. 1996, Hazir et al.

126

140

2003, Campos-Herrera et al. 2007, 2008, 2013, Abd-Elbary et al. 2012). Proportional nematode detection rate from the survey field sites did not statistically vary through the season or in between fields in both years (Fig. 5.2, Table 5.1). The proportional nematode detection frequency varied considerably between arenas causing wide standard errors in the data (Fig. 5.2). High standard errors are most likely a reflection of the natural horizontal patchy distribution that nematodes have in the soil (Stuart and

Gaugler 1994, Stuart et al. 2015). In 2015, statistically similar numbers of insect cadavers with nematodes were detected in G. mellonella vs. T. molitor baiting species. While quantitatively, nematode detection for the two baiting species may be the same, qualitatively they may differ as host preferences can vary significantly between entomopathogenic nematode (EPN) species (Simões and Rosa 1996). The small number of samples identified from 2015 (n=10, Table 5.5) did not allow host-EPN species comparisons to be made. Therefore, further studies should investigate whether using multiple host species in baiting assays allow for the isolation of a greater diversity of nematodes.

Days post inoculation (dpi) nematode detection frequency in the inoculation project was comparable to other projects with similar EPN application rates (2.5 x 109

IJ/ha) and similar dpi (Shanks and Agudelo-Silva 1990, Klein and Georgis 1992, Shields et al. 1999, Wilson et al. 2003) but was higher than Wright et al. (1993) reported despite similar application rate. Wright et al. (1993) had fewer soil samples per plot and collected the top 10 cm of soil, while we collected the top 15 cm of soil. Hence, taking in account nematode patchy distribution (horizontal and vertical) in the soil, it is possible

126

141 that the differences in methodology accounted for the differences in results between studies.

In the inoculation project, it is unclear why significant differences of nematode detection rates occurred at 90 dpi (Tables 5.2-5.3, Fig. 5.3), but it is known that the persistent nematode strains utilized in the inoculation project, S. feltiae “NY 04” strain and H. bacteriophora “Oswego” strain, are able to survive and remain viable in the laboratory over >300 days without a host (Shields 2015). This long persistence has also been shown in the field, where S. feltiae was able to recycle and persist for 3 years in a variety of cropping systems (alfalfa, vineyards, cranberry and apple) and through 7-year alfalfa-corn rotations (Shields 2015, Shields and Testa 2015). In these multi-year alfafa- corn rotations, there was a large increase in Steinernema feltiae “NY04” recovered from soil samples in second-year corn (Shields 2015). The author inferred that this increase occurred because S. feltiae was responding to WCR invasion in non-rotated corn but did not test this hypothesis (Shields 2015). However, at the one-year date (July.2016), all treatments recovered comparable low frequencies of nematodes (<2%, Fig. 5.3), and were statistically comparable to pre-inoculation levels (Table 5.3).

The significant interaction between date and depth in the inoculation project, showed that, across treatments at 14 dpi, the mean frequency of nematodes detected were significantly higher for the top 0-5 cm of soil than all others (Fig. 5.4). Nematode vertical distribution in the soil is greatly affected by their foraging and dispersal behaviors (Ferguson et al. 1995, Neumann and Shields 2006, 2008). Both Steinernema and Heterorhabditis species are present throughout the soil strata (0 – 32.5 cm), but

126

142

Steinernema tend to be more dominant in the top layers (<10 cm) and Hetererorhabditis on the bottom (>10 cm) (Ferguson et al. 1995, Glazer et al. 1996, Millar and Barbercheck

2001, Neumann and Shields 2006, Salame and Glazer 2015). Current literature that has investigated nematode vertical distribution, has focused on species composition per layer instead of the overall EPN abundance per soil layer (Fig. 5.4). Ferguson et al. (1995) found that the overall percentage of nematode infections decreased as soil depth increased for all nematode isolates, which would support the data from 7-30 dpi in this present study. Moreover, the DNA barcoding approach revealed native Steinernema sp.

(haplotype group 7) in control, commercial and persistent plots and S. feltiae (haplotype group 8) in the control plots (Fig. 5.7, Table 5.5). Therefore, one can hypothesize that the superior detection of nematodes in the 0 - 5 cm layer (Fig 5.4) was facilitated by the naturally occurring Steinernema spp. in the experimental plots.

Pre-inoculation baiting assays from the inoculation project, revealed 0.12%

(n=1620) nematode detection frequency. A background population of native nematodes throughout the season was expected based on the survey project findings (Fig. 5.1).

However, non-inoculated control plots revealed a relatively high level of nematode presence when compared to the inoculated plots throughout the collection dates (Fig.

5.3). Mean nematode detection frequency in the control plots varied significantly between dates (Fig. 5.3). Soil samples from July.15.2015 (7 dpi) and July.22.2015 (14 dpi), recovered significantly more nematodes than July.1.2015 (Pre-inococulation),

October.12.2015 (90 dpi) and July.26.2016 (1 year) (Fig. 5.3). Western corn rootworm

(WCR) egg infestation occurred on June.18.2015; by mid-July, larvae were in the second

126

143 or third larval instars, and by late July WCR pupae were also present. The entomopathogenic nematode S. carpocapsae, has similar life cycle to S. feltiae, and it completes a full lifecycle in WCR second and third instars, and pupae within 120 – 144 hours (Jackson and Brooks 1995). Western corn rootworm biology, together with the presence of native Steinernema sp. and S. feltiae (haplotype groups 7 and 8, Fig 5.6) in the control plots suggest that the increase in nematode detection in July may be from native IJs emerging from WCR larval and pupal cadavers.

It is also important to consider the possibility of cross-contamination from the inoculated plots to the control plots. Entomopathogenic nematodes are capable of moving long distances through the movement of newly infected hosts or via farming equipment (Shields et al. 2009, Shields 2015). In our system, however, the potential for nematode movement was reduced, as western corn rootworm larvae have limited movement (up to 46 cm) (Hibbard et al. 2003), farming equipment did not travel through the plots after planting, and the center-pivot irrigation system ran parallel to the plots (Fig. 5.1). Procedures such as tool sterilization, sampling control plots before treated plots and having different personnel sampling controls were also taken to prevent cross-contamination. Evidence of native Steinernema spp. from the DNA barcoding analysis, and all the procedures taken against cross contamination, makes the possibility of cross-contamination unlikely.

Despite the potential for WCR control by EPN’s reported by others in the literature (Wright et al. 1993, Jackson 1996, Toepfer et al. 2008, Kurtz et al. 2009, Pilz et al. 2009, Hiltpold et al. 2012, Shields and Testa 2015, Geisert et al. 2018), the inoculation

126

144 of S. feltiae and H. bacteriophora in WCR-infested plots did not generate expected treatment effects (Fig. 5.5). Rootworm emergence was similar in all three treatments

(Fig. 5.5) and mean root injury ratings were lower than 0.1 in all treatments. Root injury rates under 0.25 are considered minor feeding and does not cause impactful yield loss

(Oleson et al. 2005). Low establishment of WCR infestation is the most likely explanation why minor impacts of WCR feeding were detected across plots. Another explanation for the absence of EPN treatment differences is the presence of endemic EPN populations in the plots (Figs. 5.3 and 5.6), which would’ve kept the WCR populations in the control plots lower than expected.

Through the use of the COI DNA barcoding approach we were able to identify native EPN in both the survey and inoculation projects. The phylogenetic tree showed a native H. bacteriophora population was present in fields A and D in 2014 and in field C in

2015 (haplotype group 1, Fig. 5.6) and to the best of our knowledge these fields have not received EPN applications in the past. Four out of the five H. bacteriophora from the survey are identical and form a subgroup within Haplotype group 1. This subgroup is most likely H. bacteriophora given that strains obtained from the USDA and identified as

H. bacteriophora (VS, Oswego, HB strains) and H. georgiana (Kesha strain) formed another sub-group. Heterorhabditis georgiana and H. bacteriophora vary slightly in morphology but are genetically identical and form a monophyletic group based on the internal transcriber spacer (ITS) gene and the LSU D2-D3 expansion region of 28S rDNA

(Nguyen et al. 2008). Haplotype group 2 contains two GenBank H. bacteriophora strains and three identical H. bacteriophora haplotypes from persistent EPN-treated plots and a

126

145 commercial EPN-treated plot. Having two haplotype groups to be considered H. bactoriophpora is problematic. The USDA references obtained support H. bacteriophora allocation in haplotype group 1 while the GenBank database support haplotype group 2 as H. bacteriophora. A question of validity of species nomenclature is then raised. Is H. georgiana truly a different species than H. bacteriophora? Moreover, what defines H. bacteriophora? Heterorhabditids are divided into 3 monophyletic groups:

“bacteriophora-group”, “indica-group”, and “megidis-group” (Andaló et al. 2006,

Nguyen et al. 2008). Haplotype groups 1 and 2 are both considered a part of the bacteriophora-group (Maneesakorn et al. 2011, Spiridonov and Subbotin 2016). A multi- gene approach together with morphological analyses may help solve species placement of the strains in this study (Andaló et al. 2006, Nguyen et al. 2008, Spiridonov and

Subbotin 2016).

The diversity of Heterorhabditis is highlighted by haplotype groups 3 and 4.

Haplotype group 3, a monospecies clade of H. floridensis is the only member of the indica-group in this phylogenetic tree (Andaló et al. 2006). Haplotype group 4 represents the megidis-group with identical matches of H. megidis from the GenBank database and the USDA reference H. megidis UK211 (Andaló et al. 2006, Nguyen et al. 2008).

The Steinernema group was represented in four haplotype groups. Haplotype group 5 consists of S. carpocapsae specimens from the USDA references and GenBank sequences. Contrary to H. bacteriophora classification, the S. carpocapsae classification was supported by both USDA references and Genbank sequences. Haplotype group 6, S. rarum is a monospecies clade, also consistent with previous reports (Nadler et al. 2006,

126

146

Spiridonov and Subbotin 2016). Haplotype group 7 remained under Steinernema sp. and was composed of specimens from the inoculation project, treated and control plots, and one specimen from the survey project (Fig. 5.6). Two identical haplotypes (N6741 and

N6762) originated from the persistent and commercial inoculated plots, respectively.

Those plots received S. feltiae inoculations, yet those strains do not match haplotype group 8, the S. feltiae group. Haplotype group 8 consisted of sequences from the control plots from the inoculation project, USDA reference and GenBank. Similar to S. carpocapsae, S. feltiae classification was supported by GenBank sequences and USDA reference strains.

Steinernema phylogenetic classification is also heavily based on ITS and LSU D2-

D3 expansion region of 28S rDNA (Spiridonov and Subbotin 2016). Based on those gene sequences, species are currently classified under three monospecies clades and twelve multi-species clades, with ten of those clades forming three super-clades named:

Superclade 1:“Glaseri-Karii-Longicaudatum-Khoisanae”, Superclade 2: “Feltiae-Kushidai-

Monticolum” and Superclade 3: “ Carpocapsae-Bicornutum” (Spiridonov and Subbotin

2016). According to this classification, haplotype group 5 belong to Superclade 1 and haplotype group 8 belong to Superclade 2 (Nadler et al. 2006, Spiridonov and Subbotin

2016). The COI DNA barcoding approach herein highlights the importance of this work in contributing to understanding the phylogenetic diversity of EPNs. The COI phylogenetic tree was able to group Heterorhabdits and Steinernematids in similar patterns to ITS and LSU D2/D3 based phylogeny (Andaló et al. 2006, Nadler et al. 2006, Nguyen et al.

2008, Spiridonov and Subbotin 2016). This approach also enabled the identification of

126

147 nematodes from the control plots of the inoculation project and helped determine that resident EPN populations were present in treated and control plots, hence ruling out cross-contamination.

Several nematodes recovered from both projects were not entomopathogenic but were rather free-living nematodes belonging to the Diplogasteridae clade (Fig. 5.6).

Diplogasterids have diverse life-histories, they are primarily decomposers but can also be nematophagous or facultative parasites of insects and have also been shown to participate in insect phoresis (Poinar 1969, 1975, Colagiero et al. 2012). Biological control agents face a wide range of obstacles after application in the field, and one of them is their interaction with biotic factors including direct (natural enemies) and indirect antagonism (competition) predation and competition (Kaya and Koppenhöfer

1996). Hence, it can be speculated that it is possible that some of the S. feltiae and H. bacteriophora species applied in the inoculation project were eaten or displaced by the native diplogasterids in the plots, but we did not gather data to support this claim.

To our knowledge, this is the first description of naturally occurring EPNs in agroecosystems of the Midwest of the United States (Hominick 2002). Prior to human settlements, Nebraska was primarily composed of prairie, with mixed-prairie being predominant in Keith and Perkins Counties (Kaul and Rolfsmeier 1993, UNL

Conservation & Survey Division). The nematodes recovered from the cornfields in this study are most likely remnants of pre-agriculture ecosystems (Shields 2015). Nematodes have a wide range of survival mechanisms that allows them to persist in the environment (Glazer 1996). Adams (1998) found native S. feltiae and H. bacteriophora in

126

148 native and grazed prairie of Western Nebraska in Arthur Co. and Keith Co. Both EPN species were also found in this present study in cornfields in Keith and Perkins Counties.

Finding native EPN strains that can survive the intensive agricultural practices adopted in local fields may increase EPN efficacy against target pests (Glazer 1996, Shields 2015).

Local H. bacteriophora, Steinernema sp. and S. feltiae should be re-isolated from the field sites in this study and tested against WCR larvae and other soil pests. This work provides a foundation for future ecological and pest management studies with EPN in irrigated corn systems.

126

149

References

Abd-Elbary, N. A., M. S. Shamseldean, S. P. Stock, and N. M. Abu-Shady. 2012. Diversity of entomopathogenic nematode species (Heterorhabditidae and Steinernematidae) in Egypt. Egypt. J. Agronematology. 11: 333–353.

Adams, B. J. 1998. The ecology and evolution of Heterorhabditid nematodes and their symbiotic bacteria: phylogeny, species concepts, coevolution, and ecological observations. Ph.D. Dissertation. University of Nebraska- Lincoln.

Adams, B. J., A. Fodor, H. S. Koppenhöfer, E. Stackebrandt, S. Patricia Stock, and M. G. Klein. 2006. Biodiversity and systematics of nematode–bacterium entomopathogens. Biol. Control. 37: 32–49.

Bhadury, P., M. C. Austen, D. T. Bilton, P. J. D. Lambshead, A. D. Rogers, and G. R. Smerdon. 2006. Development and evaluation of a DNA-barcoding approach for the rapid identification of nematodes. Mar. Ecol. Prog. Ser. 320: 1–9.

Andaló, V., K. B. Nguyen, and A. Moino. 2006. Heterorhabditis amazonensis n. sp. (Rhabditida: Heterorhabditidae) from Amazonas, Brazil. Nematology. 8: 853– 867. de Brida, A. L., J. M. O. Rosa, C. M. G. de Oliveira, B. M. de C. e Castro, J. E. Serrão, J. C. Zanuncio, L. G. Leite, and S. R. S. Wilcken. 2017. Entomopathogenic nematodes in agricultural areas in Brazil. Sci. Rep. 7.

Cabanillas, H. E., and J. R. Raulston. 1994. Evaluation of the spatial pattern of Steinernema riobravis in Corn Plots. J. Nematol. 26: 25–31.

Campbell, J. F., E. Lewis, F. Yoder, and R. Gaugler. 1996. Entomopathogenic nematode (Heterorhabditidae and Steinernematidae) spatial distribution in turfgrass. Parasitology. 113: 473–482.

Campbell, J. F., G. Orza, F. Yoder, E. Lewis, and R. Gaugler. 1998. Spatial and temporal distribution of endemic and released entomopathogenic nematode populations in turfgrass. Entomol. Exp. Appl. 86: 1–11.

Campos-Herrera, R., M. Escuer, S. Labrador, L. Robertson, L. Barrios, and C. Gutiérrez. 2007. Distribution of the entomopathogenic nematodes from La Rioja (Northern Spain). J. Invertebr. Pathol. 95: 125–139.

Campos-Herrera, R., J. M. Gómez-Ros, M. Escuer, L. Cuadra, L. Barrios, and C. Gutiérrez. 2008. Diversity, occurrence, and life characteristics of natural entomopathogenic nematode populations from La Rioja (Northern Spain) under

126

150

different agricultural management and their relationships with soil factors. Soil Biol. Biochem. 40: 1474–1484.

Campos-Herrera, R., E. Pathak, F. E. El-Borai, R. J. Stuart, C. Gutiérrez, J. A. Rodríguez- Martín, J. H. Graham, and L. W. Duncan. 2013. Geospatial patterns of soil properties and the biological control potential of entomopathogenic nematodes in Florida citrus groves. Soil Biol. Biochem. 66: 163–174.

Choo, H. Y., A. M. Koppenhöfer, and H. K. Kaya. 1996. Combination of two entomopathogenic nematode species for suppression of an insect pest. J. Econ. Entomol. 89: 97–103.

Colagiero, M., L. C. Rosso, and S. N. Murga-Gutierrez. 2012. Observations on the biology of a predatory nematode belonging to Diplogasteridae. Redia. 94: 133– 135.

Dolinski, C., E. E. Del Valle, R. S. Burla, and I. R. Machado. 2007. Biological traits of two native Brazilian entomopathogenic nematodes (Rhabditida: Heterorhabditidae). Nematol. Bras. 31: 180–185.

Dolinski, C., D. Shapiro-Ilan, and E. E. Lewis. 2015. Insect cadaver applications: pros and cons, pp. 207–229. In Nematode Pathog. Insects pests, sustainability in plant and crop protection. Springer, Cham.

Efron, D., D. Nestel, and I. Glazer. 2001. Spatial analysis of entomopathogenic nematodes and insect hosts in a citrus grove in a semi-arid region in Israel. Environ. Entomol. 30: 254–261.

Ferguson, C. S., P. C. Schroeder, and E. J. Shields. 1995. Vertical distribution, persistence, and activity of entomopathogenic nematodes (Nematoda: Heterorhabditidae and Steinernematidae) in alfalfa snout beetle-(Coleoptera: Curculionidae) infested fields. Environ. Entomol. 24: 149–158.

Ferrari, S., and F. Cribari-Neto. 2004. Beta regression for modelling rates and proportions. J. Appl. Stat. 31: 799–815.

Garcia del Pino, F., and A. Palomo. 1996. Natural occurrence of entomopathogenic nematodes (Rhabditida: Steinernematidae and Heterorhabditidae) in Spanish Soils. J. Invertebr. Pathol. 68: 84–90.

Geisert, R. W., D. J. Cheruiyot, B. E. Hibbard, D. I. Shapiro-Ilan, K. S. Shelby, and T. A. Coudron. 2018. Comparative assessment of four Steinernematidae and three Heterorhabditidae species for infectivity of larval Diabrotica virgifera virgifera. J. Econ. Entomol.

126

151

Glazer, I. 1996. Survival mechanisms of entomopathogenic nematodes. Biocontrol Sci. Technol. 6: 373–378.

Glazer, I., E. Kozodoi, L. Salame, and D. Nestel. 1996. Spatial and temporal occurrence of natural populations of Heterorhabditis spp. (Nematoda: Rhabditida) in a Semiarid Region. Biol. Control. 6: 130–136.

Grewal, P. S., X. Wang, and R. A. J. Taylor. 2002. Dauer juvenile longevity and stress tolerance in natural populations of entomopathogenic nematodes: is there a relationship? Int. J. Parasitol. 32: 717–725.

Hara, A. H., R. Gaugler, H. K. Kaya, and L. M. Lebeck. 1991. Natural populations of entomopathogenic nematodes (Rhabditida: Heterorhabditidae, Steinernematidae) from the Hawaiian Islands. Environ. Entomol. 20: 211–216.

Hazir, S., N. Keskin, S. P. Stock, H. K. Kaya, and S. Özcan. 2003. Diversity and distribution of entomopathogenic nematodes (Rhabditida: Steinernematidae and Heterorhabditidae) in Turkey. Biodivers. Conserv. 12: 375–386.

Hebert, P. D., S. Ratnasingham, and J. R. de Waard. 2003. Barcoding animal life: cytochrome c oxidase subunit 1 divergences among closely related species. Proc. R. Soc. Lond. B Biol. Sci. 270: S96–S99.

Hibbard, B. E., D. P. Duran, M. R. Ellersieck, and M. M. Ellsbury. 2003. Post- establishment movement of western corn rootworm larvae (Coleoptera: Chrysomelidae) in central Missouri corn. J. Econ. Entomol. 96: 599–608.

Hibbard, B. E., L. N. Meihls, M. R. Ellersieck, and D. W. Onstad. 2010. Density- dependent and density-independent mortality of the western corn rootworm: impact on dose calculations of rootworm-resistant Bt corn. J. Econ. Entomol. 103: 77–84.

Hiltpold, I., B. E. Hibbard, B. W. French, and T. C. J. Turlings. 2012. Capsules containing entomopathogenic nematodes as a Trojan horse approach to control the western corn rootworm. Plant Soil. 358: 11–25.

Hominick, W. M. 2002. Biogeography. Entomopathog. Nematol. 1: 115–143.

Hominick, W. M., A. P. Reid, D. A. Bohan, and B. R. Briscoe. 1996. Entomopathogenic nematodes: biodiversity, geographical distribution and the convention on biological diversity. Biocontrol Sci. Technol. 6: 317–332.

Jackson, J. J. 1996. Field performance of entomopathogenic nematodes for suppression of western corn rootworm (Coleoptera: Chrysomelidae). J. Econ. Entomol. 89: 366–372.

126

152

Jackson, J. J., and M. A. Brooks. 1995. Parasitism of western corn rootworm larvae and pupae by Steinernema carpocapsae. J. Nematol. 27: 15–20.

Jackson, T. A., S. B. Alves, and R. M. Pereira. 2000. Success in biological control of soil- dwelling insects by pathogens and nematodes, pp. 271–296. In Gurr, G., Wratten, S. (eds.), Biol. Control Meas. Success. Springer Netherlands.

Journey, A. M., and K. R. Ostlie. 2000. Biological control of the western corn rootworm (Coleoptera: Chrysomelidae) using the entomopathogenic nematode, Steinernema carpocapsae. Environ. Entomol. 29: 822–831.

Kanzaki, N., and K. Futai. 2002. A PCR primer set for determination of phylogenetic relationships of Bursaphelenchus species within the xylophilus group. Nematology. 4: 35–41.

Kaul R. and S. Rolfsmeier. 1993. Natural vegetation of Nebraska. UNL Conservation & Survey Division.

Kaya, H. K., and A. M. Koppenhöfer. 1996. Effects of microbial and other antagonistic organism and competition on entomopathogenic nematodes. Biocontrol Sci. Technol. 6: 357–372.

Klein, M. G., and R. Georgis. 1992. Persistence of control of japanese beetle (Coleoptera: Scarabaeidae) larvae with Steinernematid and Heterorhabditid nematodes. J. Econ. Entomol. 85: 727–730.

Koppenhöfer, A. M. 2007. Nematodes, pp. 249–264. In Field Man. Tech. Invertebr. Pathol. Springer.

Koppenhöfer, A. M., M. Wilson, I. Brown, H. K. Kaya, and R. Gaugler. 2000. Biological control agents for white grubs (Coleoptera: Scarabaeidae) in anticipation of the establishment of the japanese beetle in california. J. Econ. Entomol. 93: 71–80.

Kung, S.-P., R. Gaugler, and H. K. Kaya. 1991. Effects of soil temperature, moisture, and relative humidity on entomopathogenic nematode persistence. J. Invertebr. Pathol. 57: 242–249.

Kurtz, B., I. Hiltpold, T. C. J. Turlings, U. Kuhlmann, and S. Toepfer. 2009. Comparative susceptibility of larval instars and pupae of the western corn rootworm to infection by three entomopathogenic nematodes. BioControl. 54: 255–262.

Liu, J., and R. E. Berry. 1995. Natural distribution of entomopathogenic nematodes (Rhabditida: Heterorhabditidae and Steinernematidae) in Oregon Soils. Environ. Entomol. 24: 159–163.

126

153

Maneesakorn, P., R. An, H. Daneshvar, K. Taylor, X. Bai, B. J. Adams, P. S. Grewal, and A. Chandrapatya. 2011. Phylogenetic and cophylogenetic relationships of entomopathogenic nematodes (Heterorhabditis: Rhabditida) and their symbiotic bacteria (Photorhabdus: Enterobacteriaceae). Mol. Phylogenet. Evol. 59: 271– 280.

Millar, L. C., and M. E. Barbercheck. 2001. Interaction between endemic and introduced entomopathogenic nematodes in conventional-till and no-till corn. Biol. Control. 22: 235–245.

Millar, L. C., and M. E. Barbercheck. 2002. Effects of tillage practices on entomopathogenic nematodes in a corn agroecosystem. Biol. Control. 25: 1–11.

Nadler, S. A., E. Bolotin, and S. P. Stock. 2006. Phylogenetic relationships of Steinernema Travassos, 1927 (Nematoda: Cephalobina: Steinernematidae) based on nuclear, mitochondrial and morphological data. Syst. Parasitol. 63: 159–179.

Neumann, G., and E. J. Shields. 2006. Interspecific Interactions among three entomopathogenic nematodes, Steinernema carpocapsae Weiser, S. feltiae Filipjev, and Heterorhabditis bacteriophora Poinar, with different foraging strategies for hosts in multipiece sand columns. Environ. Entomol. 35: 1578– 1583.

Neumann, G., and E. J. Shields. 2008. Multiple-species natural enemy approach for biological control of alfalfa snout beetle (Coleoptera: Curculionidae) using entomopathogenic nematodes. J. Econ. Entomol. 101: 1533–1539.

Nguyen, K. B., D. I. Shapiro-Ilan, and G. N. Mbata. 2008. Heterorhabditis georgiana n. sp. (Rhabditida: Heterorhabditidae) from Georgia, USA. Nematology. 10: 433– 448.

Oleson, J. D., Y.-L. Park, T. M. Nowatzki, and J. J. Tollefson. 2005. Node-injury scale to evaluate root injury by corn rootworms (Coleoptera: Chrysomelidae). J. Econ. Entomol. 98: 1–8.

Pierce, C. M. F., and M. E. Gray. 2007. Population dynamics of a western corn rootworm (Coleoptera: Chrysomelidae) variant in East Central Illinois commercial maize and soybean fields. J. Econ. Entomol. 100: 1104–1115.

Pilz, C., S. Keller, U. Kuhlmann, and S. Toepfer. 2009. Comparative efficacy assessment of fungi, nematodes and insecticides to control western corn rootworm larvae in maize. BioControl. 54: 671–684.

Poinar, G. O. 1969. Diplogasterid nematodes (diplogasteridae: Rhabditida) and their relationship to insect disease. J. Invertebr. Pathol. 13: 447–454.

126

154

Poinar, G. O. 1975. Entomogenous nematodes: a manual and host list of insect- nematode associations. Brill Archive.

Powers, T. 2004. Nematode molecular diagnostics: From bands to barcodes. Annu. Rev. Phytopathol. 42: 367–383.

Powers, T. O., E. C. Bernard, T. Harris, R. Higgins, M. Olson, M. Lodema, P. Mullin, L. Sutton, and K. S. Powers. 2014. COI haplotype groups in Mesocriconema (Nematoda: Criconematidae) and their morphospecies associations. Zootaxa. 3827: 101–146.

Prosser, S. W., M. G. Velarde-Aguilar, V. León-Règagnon, and P. D. Hebert. 2013. Advancing nematode barcoding: a primer cocktail for the cytochrome c oxidase subunit I gene from vertebrate parasitic nematodes. Mol. Ecol. Resour. 13: 1108–1115.

Ritchie, W. W., J. J. Hanway, and G. O. Benson. 1992. How a corn plant develops. Iowa State University of Science and Technology Cooperative Extension Service, Special Report 48. Ames, IA.

Salame, L., and I. Glazer. 2015. Stress avoidance: vertical movement of entomopathogenic nematodes in response to soil moisture gradient. Phytoparasitica. 43: 647–655.

Seesao, Y., M. Gay, S. Merlin, E. Viscogliosi, C. M. Aliouat-Denis, and C. Audebert. 2017. A review of methods for nematode identification. J. Microbiol. Methods, What’s next in microbiology methods? Emerging methods. 138: 37–49.

Schabenberger, O. 2005. Introducing the GLIMMIX procedure for generalized linear mixed models. SUGI 30 Proc. 196.

Shanks, C. H., and F. Agudelo-Silva. 1990. Field pathogenicity and persistence of Heterorhabditid and Steinernematid nematodes (Nematoda) infecting black vine weevil larvae (Coleoptera: Curculionidae) in cranberry bogs. J. Econ. Entomol. 83: 107–110.

Shapiro-Ilan, D. I., E. E. Lewis, Y. Son, and W. L. Tedders. 2003. Superior efficacy observed in entomopathogenic nematodes applied in infected-host cadavers compared with application in aqueous suspension. J. Invertebr. Pathol. 83: 270– 272.

Shields, E. J. 2015. Utilizing persistent entomopathogenic nematodes in a conservation or a more classical biological control approach, pp. 165–184. In Nematode pathog. insects pests, sustainability in plant and crop protection. Springer, Cham.

126

155

Shields, E. J., and A. M. Testa. 2015. New York Case Study: biological control of Otiorhynchus ligustici with Native persistent entomopathogenic nematodes using a more classical approach, pp. 285–307. In Nematode pathog. insects pests, sustainability in plant and crop protection. Springer, Cham.

Shields, E. J., A. Testa, J. M. Miller, and K. L. Flanders. 1999. Field efficacy and persistence of the entomopathogenic nematodes Heterorhabditis bacteriophora ‘Oswego’ and H. bacteriophora ‘NC’ on alfalfa snout beetle larvae (Coleoptera: Curculionidae). Environ. Entomol. 28: 128–136.

Shields, E. J., A. Testa, G. Neumann, K. L. Flanders, and P. C. Schroeder. 2009. Biological control of alfalfa snout beetle with a multi-species application of locally adapted persistent entomopathogenic nematodes: the first success. Am. Entomol. 55: 250–257.

Simões, N., and J. S. Rosa. 1996. Pathogenicity and host specificity of entomopathogenic nematodes. Biocontrol Sci. Technol. 6: 403–412.

Spiridonov, S. E., A. P. Reid, K. Podrucka, S. A. Subbotin, and M. Moens. 2004. Phylogenetic relationships within the genus Steinernema (Nematoda: Rhabditida) as inferred from analyses of sequences of the ITS1-5.8 S-ITS2 region of rDNA and morphological features. Nematology. 6: 547–566.

Spiridonov, S. E., M. Moens, and M. J. Wilson. 2007. Fine scale spatial distributions of two entomopathogenic nematodes in a grassland soil. Appl. Soil Ecol. 37: 192– 201.

Spiridonov, S. E., and S. A. Subbotin. 2016. Phylogeny and phylogeography of Heterorhabditis and Steinernema. Adv Entomopathog Nematode Taxon Phylogeny. 12: 413–427.

Stock, S.P., J. F. Campbell, and S. A. Nadler. 2001. Phylogeny of Steinernema Travassos, 1927 (Cephalobina: Steinernematidae) inferred from ribosomal DNA sequences and morphological characters. J. Parasitol. 87: 877–889.

Stock, S. P., and H. Goodrich-Blair. 2012. Nematode parasites, pathogens and associates of insects and invertebrates of economic importance. In Manual of techniques in invertebrate pathology. 375-425. Academic press, CA.

Stock, S. P. 2015. Diversity, biology and evolutionary relationships, pp. 3–27. In Nematode Pathog. Insects Pests. Springer.

Strong, D. R., A. V. Whipple, A. L. Child, and B. Dennis. 1999. Model selection for a subterranean trophic cascade: root-feeding caterpillars and entomopathogenic nematodes. Ecology. 80: 2750–2761.

126

156

Stroup, W. W. 2015. Rethinking the analysis of non-normal data in plant and soil science. Agron. J. 107: 811–827.

Stuart, R. J., M. E. Barbercheck, and P. S. Grewal. 2015. Entomopathogenic nematodes in the soil environment: distributions, interactions and the influence of biotic and abiotic factors, pp. 97–137. In Campos-Herrera, R. (ed.), Nematode pathog. insects pests, sustainability in plant and crop protection. Springer International Publishing.

Stuart, R. J., M. E. Barbercheck, P. S. Grewal, R. A. J. Taylor, and C. W. Hoy. 2006. Population biology of entomopathogenic nematodes: Concepts, issues, and models. Biol. Control, Third International Symposium on Entomopathogenic Nematodes and Symbiotic Bacteria. 38: 80–102.

Stuart, R. J., and R. Gaugler. 1994. Patchiness in populations of entomopathogenic nematodes. J. Invertebr. Pathol. 64: 39–45.

Sutter, G. R., and T. F. Branson. 1980. A procedure for artificially infesting field plots with corn rootworm eggs. J. Econ. Entomol. 73: 135–137.

Tamura, K., G. Stecher, D. Peterson, A. Filipski, and S. Kumar. 2013. MEGA6: Molecular Evolutionary Genetics Analysis Version 6.0. Mol. Biol. Evol. 30: 2725–2729.

Toepfer, S., C. Gueldenzoph, R.-U. Ehlers, and U. Kuhlmann. 2005. Screening of entomopathogenic nematodes for virulence against the invasive western corn rootworm, Diabrotica virgifera virgifera (Coleoptera: Chrysomelidae) in Europe. Bull. Entomol. Res. 95: 473–482.

Toepfer, S., A. Peters, R.-U. Ehlers, and U. Kuhlmann. 2008. Comparative assessment of the efficacy of entomopathogenic nematode species at reducing western corn rootworm larvae and root damage in maize. J. Appl. Entomol. 132: 337–348.

Tripathi,te R. C. 2006. Negative Binomial Distribution. Encycl. Stat. Sci., Major Reference Works.

White, G. F. 1927. A method for obtaining infective nematode larvae from cultures. Sci. Wash. 66.

Wilson, M. J., E. E. Lewis, F. Yoder, and R. Gaugler. 2003. Application pattern and persistence of the entomopathogenic nematode Heterorhabditis bacteriophora. Biol. Control. 26: 180–188.

Wright, R. J., J. F. Witkowski, G. Echtenkamp, and R. Georgis. 1993. Efficacy and persistence of Steinernema carpocapsae (Rhabditida: Steinemematidae) applied through a center-pivot irrigation system against larval com rootworms (Coleoptera: Chrysomelidae). J. Econ. Entomol. 86: 1348–1354.

126

Tables

Table 5.1. General linear mixed model analysis of nematode detection in the survey project.

Effect Df F P

2014 Field 4, 306 0.77 0.5439 Date 6, 306 0.79 0.5756 Field x date 23, 306 0.73 0.9905

2015 Field 4, 175 0.26 0.9015 Date 4, 175 0.14 0.9666 Field x date 16, 175 0.15 1.0000

Statistical analysis was conducted with PROC GLIMMIX in SAS using two-way generalized linear mixed model ANOVA. Nematode detection was expressed as the mean proportion of nematode-infected cadavers per arena for each sampling date in each field. Mean proportion was fitted to a beta-binomial distribution prior to analysis. Main effects and interactions at P < 0.05 were considered significant.

157

Table 5.2. General linear mixed model analysis of nematode detection in the inoculation project.

Effect Df F P

Treatment 2, 10 0.04 0.9619

Soil Depth 1, 195 0.63 0.4286

Date 6, 195 8.83 <.0001

Treatment x Soil Depth 2, 195 0.11 0.8981

Treatment x Date 12, 195 2.04 0.0229

Soil Depth x Date 6, 195 2.61 0.0185

Treatment x Soil Depth x Date 12, 195 0.30 0.9883

Statistical analysis was conducted with PROC GLIMMIX in SAS using a three-way generalized linear mixed model ANOVA. Nematode detection frequency was expressed as the mean proportion of infected baiting insects per sampling depth per replicate (30 larvae per replicate for 5 cm soil depth, and 60 larvae per replicate for 10 cm soil depths). Mean proportion was fitted to a beta- binomial distribution prior to analysis. Sampling dates were treated as repeated measures with first order autoregressive covariance structure (AR-1). Main effects and interactions at P < 0.05 were considered significant

158

Table 5.3. Differences of treatment by sampling date least square means with Tukey adjustment for multiple comparisons. All values were considered non-significant (NS) if adjusted P > 0.05.

159

Table 5.4. Differences of depth by sampling date least square means with Tukey adjustment for multiple comparisons. All values were considered non-significant (NS) if adjusted P > 0.05.

160

Table.5.5. Specimen origins for samples used in phylogenetic tree (Fig 5.6). Nematode Identification Number (NID), strains are in parenthesis. HG= Haplotype group. HG NID Year Species ID Locality Project Treatment Accession Number 7 6603 2014 Steinernema sp. Perkins Co., NE Survey Field B - 6605 2014 Diplogasteridae Perkins Co., NE Survey Field E 1 6607 2014 Heterorhabditis bacteriophora Keith Co., NE Survey Field A - 6609 2014 Koerneria sp. Perkins Co., NE Survey Field C 1 6616 2014 Heterorhabditis bacteriophora Perkins Co., NE Survey Field D - 6620 2015 Diplogasteridae Perkins Co., NE Survey Field C - 6629 2015 Diplogasteridae Perkins Co., NE Survey Field B - 6632 2015 Diplogasteridae Perkins Co., NE Survey Field B - 6635 2015 Diplogasteridae Perkins Co., NE Survey Field C 1 6642 2015 Heterorhabditis bacteriophora Perkins Co., NE Survey Field C 1 6649 2015 Heterorhabditis bacteriophora Perkins Co., NE Survey Field C - 6656 2015 Diplogasteridae Perkins Co., NE Survey Field B 1 6663 2015 Heterorhabditis bacteriophora Perkins Co., NE Survey Field C - 6667 2015 Diplogasteridae Perkins Co., NE Survey Field B - 6672 2015 Diplogasteridae Keith Co., NE Survey Field A - 6675 2015 Diplogasteridae Keith Co., NE Inoculation Control - 6678 2015 Diplogasteridae Keith Co., NE Inoculation Control 7 6682 2015 Steinernema sp. Keith Co., NE Inoculation Control - 6691 2015 Oscheius onirici Keith Co., NE Inoculation Control - 6700 2015 Diplogasteridae Keith Co., NE Inoculation Control - 6703 2015 Diplogasteridae Keith Co., NE Inoculation Control 8 6705 2015 Steinernema feltiae Keith Co., NE Inoculation Control - 6710 2015 Acrostichus sp. Keith Co., NE Inoculation Control - 6714 2015 Diplogasteridae Keith Co., NE Inoculation Control 8 6717 2015 Steinernema feltiae Keith Co., NE Inoculation Control 2 6723 2015 Heterorhabditis bacteriophora Keith Co., NE Inoculation Persistent

161

Table 5.5. (Continued) HG NID Year Species ID Locality Project Treatment Accession Number 2 6729 2015 Heterorhabditis bacteriophora Keith Co., NE Inoculation Commercial - 6734 2015 Koerneria sp. Keith Co., NE Inoculation Control 7 6738 2015 Steinernema sp. Keith Co., NE Inoculation Control 7 6741 2015 Steinernema sp. Keith Co., NE Inoculation Persistent 2 6752 2016 Heterorhabditis bacteriophora Keith Co., NE Inoculation Persistent 7 6762 2016 Steinernema sp. Keith Co., NE Inoculation Commercial - 6765 2016 Diplogasteridae Keith Co., NE Inoculation Control - 6769 2016 Diplogasteridae Keith Co., NE Inoculation Control 7 6770 2016 Steinernema sp. Keith Co., NE Inoculation Control 1 6755 2016 Heterorhabditis bacteriophora N/A Reference USDA (HB) 1 6757 2016 Heterorhabditis bacteriophora N/A Reference USDA (HB) 1 6778 2016 Heterorhabditis bacteriophora N/A Reference USDA (Oswego) 1 6779 2016 Heterorhabditis bacteriophora N/A Reference USDA (Oswego) 4 6782 2016 Heterorhabditis megidis (UK211) N/A Reference USDA 4 6783 2016 Heterorhabditis megidis (UK211) N/A Reference USDA 8 6791 2016 Steinernema feltiae (SN) N/A Reference USDA 8 6792 2016 Steinernema feltiae (SN) N/A Reference USDA 5 6797 2016 Steinernema carpocapsae (All) N/A Reference USDA 5 6798 2016 Steinernema carpocapsae (All) N/A Reference USDA 6 6801 2017 Steinernema rarum (17C&E) N/A Reference USDA 6 6802 2017 Steinernema rarum (17C&E) N/A Reference USDA 1 6811 2017 Heterorhabditis bacteriophora N/A Reference USDA (VS) 162

Table 5.5. (Continued) HG NID Year Species ID Locality Project Treatment Accession Number 1 6813 2017 Heterorhabditis bacteriophora N/A Reference USDA (VS) 5 6816 2017 Steinernema carpocapsae (Cxrd) N/A Reference USDA 5 6817 2017 Steinernema carpocapsae (Cxrd) N/A Reference USDA 1 6822 2017 Heterorhabditis georgiana N/A Reference USDA (Kesha) 1 6823 2017 Heterorhabditis georgiana N/A Reference USDA (Kesha) 3 6826 2017 Heterorhabditis floridensis (K22) N/A Reference USDA 3 6827 2017 Heterorhabditis floridensis (K22) N/A Reference USDA 2 - Heterorhabditis bacteriophora Bari, Italy Reference GenBank LN912990 2 - Heterorhabditis bacteriophora Maynooth, Reference GenBank EF043402 (HP88) Ireland 4 - Heterorhabditis megidis Cardiff, Reference GenBank DQ285542 Wales - - Oscheius sp. (TGO) Bari, Italy Reference GenBank LN613269 - - Koerneria sp. Tuebingen, Reference GenBank JX163960 Germany - - Acrostichus sp. Tuebingen, Reference GenBank JX163961 Germany - - Parapristionchus giblindavisi Tuebingen, Reference GenBank JX163962 Germany - - Neodiplogaster sp. Tsukuba, Reference GenBank AB478642 Japan - Acrostichus sp. Tsukuba, Reference GenBank AB477082 Japan 5 - Steinernema carpocapsae Kiyotake, Reference GenBank AP017465 Japan

163

Table 5.5. (Continued) HG NID Year Species ID Locality Project Field/Plot Accession Number 5 - Steinernema carpocapsae Ponta Delgada, Reference GenBank AY591323 Portugal 5 - Steinernema carpocapsae Bari, Italy Reference GenBank LN912989 8 - Steinernema feltiae Bari, Italy Reference GenBank LM608088 8 - Steinernema feltiae Bari, Italy Reference GenBank LM608089 8 - Steinernema feltiae Bari, Italy Reference GenBank LM608090

164

165

Figures

Fig 5.1. Diagram of EPN inoculation project. Top: Plots were arranged into a randomized complete block design of 6 blocks, for a total of 6 replicates per treatment. Blocks are shown outlined in black. Each plot measured 24.4 m (32 rows) x 24.4 m. Bottom: Shown in shadow is the inoculation zone (2.5 x 2.5 meters) of the persistent treatment. Infected G. mellonella cadavers of S. feltiae (circle) and H. bacteriophora (triangle) were placed every 122 cm. Inoculation of commercial treatment consisted of equal concentration of each EPN species mixed together and applied over the inoculation zone with a watering can.

166 a)

b)

Figure. 5.2. Mean nematode detection per arena in survey sites in 2014 (a) and 2015 (b). Nematode detection was analyzed as proportion data and multiplied by 100 to obtain percentages. There were no significant interactions or main effects of field or date in both years (Table 5.1).

167

25% a ab a 20% a abc ab abc 15% abc abc

10% abc

5% bc c c Mean nematode detection per replicate detection Mean nematode abc abc

0% Pre-Inoc 7 dpi 14 dpi 30 dpi 60 dpi 90 dpi 1 year Sampling Period Control Commercial Persistent

Figure 5.3. Mean nematode detection per treatment over sampling dates. Treatment by time interaction was significant (F12, 195 = 2.04, Pr> F = 0.0229). Different letters represent mean detection differences at the P < 0.05 level.

168

35% a

30%

25%

20% b b b 15% b bc bcd 10% b b b 5% d Mean nematode detection per replicate detection Mean nematode abcd 0% Pre-Inoc 7 dpi 14 dpi 30 dpi 60 dpi 90 dpi 1 year Sampling Period 0-5 cm 5-15 cm

Figure 5.4 Mean nematode detection (averaged across treatments) by sampling depth over collection periods. Depth by time interaction significant (F6,195 = 2.61, Pr > F < 0.0185). Different letters represent mean detection differences at the P < 0.05 level. Overall interaction least square mean differences shown in table 5.4.

169

Figure 5.5. Cumulative WCR emergence distribution among treatments. Treatments received three cages per plot for a total of 36 cages per treatment. Black line on box- plot represent emergence median. Mean WCR emergence per cage was not significantly different among treatments (F2, 105 = 2.72, Pr > F = 0.0706).

170

N6811 Heterorhabditis bacteriophora VS strain N6813 Heterorhabditis bacteriophora VS strain

6 4 N6779 Heterorhabditis bacteriophora Oswego strain N6778 Heterorhabditis bacteriophora Oswego strain

9 9 N6757 Heterorhabditis bacteriophora HB strain N6755 Heterorhabditis bacteriophora HB strain 1 N6822 Heterorhabditis georgiana Kesha strain 100 6 7 N6823 Heterorhabditis georgiana Kesha strain N6616 Heterorhabditis bacteriophora N6607 Heterorhabditis bacteriophora 9 9 8 0 N6642 Heterorhabditis bacteriophora 6 9 N6649 Heterorhabditis bacteriophora N6663 Heterorhabditis bacteriophora 2 LN912990.1 Heterorhabditis bacteriophora COI GB 6 4 EF043402.1 Heterorhabditis bacteriophora COI GB 100 N6723 Heterorhabditis bacteriophora 9 0 N6729 Heterorhabditis bacteriophora N6752 Heterorhabditis bacteriophora 9 8 3 N6826 Heterorhabditis floridensis K22 strain 100 N6827 Heterorhabditis floridensis K22 strain 4 N6782 Heterorhabditis megidis UK211 strain

100 DQ285542.1 Heterorhabditis megidis COI GB N6783 Heterorhabditis megidis UK211 strain

100 N6609 Unknown N6710 Unknown 6 1 N6691 Oscheius sp. 100 LN613269.1 Oscheius sp. COI GB AB477082.1 Acrostichus sp. COI GB N6734 Unknown JX163960.1 Koerneria sp. COI GB JX163961.1 Acrostichus sp. COI GB AB478642.1 Neodiplogaster sp. COI GB JX163962.1 Parapristionchus giblindavisi COI GB N6672 Diplogasteridae

8 9 N6675 Diplogasteridae N6678 Diplogasteridae 9 9 9 9 N6629 Diplogasteridae N6667 Diplogasteridae

100 N6703 Diplogasteridae 100 N6769 Diplogasteridae

5 4 N6605 Diplogasteridae Diplogasteridae 7 0 9 9 N6765 Diplogasteridae N6635 Diplogasteridae

6 3 N6700 Diplogasteridae

7 3 N6620 Diplogasteridae N6714 Diplogasteridae 8 7 N6632 Diplogasteridae 9 2 N6656 Diplogasteridae

7 3 N6797 Steinernema carpocapsae All strain 6 5 N6798 Steinernema carpocapsae All strain 9 4 N6816 Steinernema carpocapsae Cxrd strain 5 9 4 AP017465.1 Steinernema carpocapsae COI GB AY591323.1 Steinernema carpocapsae COI GB 100 9 6 N6817 Steinernema carpocapsae Cxrd strain LN912989.1 Steinernema carpocapsae COI GB 6 N6801 Steinernema rarum 17C8E strain 100 N6802 Steinernema rarum 17C8E strain N6741 Steinernema sp. 6 2 N6762 Steinernema sp. 6 0 8 7 N6682 Steinernema sp. 6 3 7 N6603 Steinernema sp. 100 N6770 Steinernema sp. N6738 Steinernema sp.

7 6 LM608088.1 Steinernema feltiae COI GB

9 5 N6791 Steinernema feltiae SN strain 100 N6792 Steinernema feltiae SN strain

100 LM608089.1 Steinernema feltiae COI GB 8 LM608090.1 Steinernema feltiae COI GB N6705 Steinernema feltiae 5 4 N6717 Steinernema feltiae

0.02 Figure 5.6. Neighbor-joining tree of COI nucleotide sequence from 70 nematode specimens. Haplotype groups are enumerated in circles adjacent to terminal branch tips. NID (Nematode Identification number) sampling characteristics are provided in Table 5.5

171

CHAPTER 6: SUMMARY AND CONCLUSIONS

Management of western corn rootworm (WCR), Diabrotica virgifera virgifera

LeConte (Chrysomelidae: Coleoptera) is a complex, multivariate issue. There are many tactics used to control WCR populations to prevent or reduce yield losses in corn.

Currently used tactics to manage WCR are crop rotation, Bacillus thuringiensis (Bt) hybrids and chemical control. Crop rotation breaks the pest cycle by removing the host for larval feeding. However, there are many reasons why farmers keep their fields as continuous corn, including livestock needs, economics of corn production especially under irrigation, soil properties, and contractual obligations of rented land (Andow et al.

2017). All three controls may be effective but because they are widely used, WCR has adapted to one or more management practices in several areas of the Corn Belt (Meinke et al. 1998, Levine et al. 2002, Parimi et al. 2006, Gassmann et al. 2011, 2014, 2016,

Wangila et al. 2015, Zukoff et al. 2016, Ludwick et al. 2017). It is necessary to examine new tools for WCR control to provide new management practices as well as to help extend the lifetime of existing technologies.

The overall goal of this dissertation was to characterize the communities of natural enemies in commercial cornfields in the context of examining their potential as

WCR biological control agents (BCAs). Specifically, objectives were developed to look at arthropod predators (Chapter 2), entomopathogenic fungi (Chapters 3 and 4) and entomopathogenic nematodes (Chapter 5) in lab and field studies.

The survey for above ground arthropods (Chapter 2) revealed that commercial cornfields support an abundant and diverse community of predators. The predators

172 caught on the yellow-sticky cards are not known predators of the WCR (Kuhlmann and

Van der Burgt 1998, Toepfer et al. 2009). Also, none of the Carabid beetles from dry pitfalls tested positive for WCR DNA in molecular gut-content analyses. Western corn rootworm prey was available in all fields but especially abundant in fields A and C, even though these fields contained WCR- Bt traits. These results expand on findings that arthropod predators are likely having minimal impacts on WCR mortality in the fields

(Kirk 1982, Lundgren and Fergen 2014).

Looking at the below-ground microbial natural enemies, a diverse assemblage of entomogenous fungi were isolated from the soil of the same cornfields from Chapter 2 via Galleria mellonella (L.) (Lepidoptera: Pyralidae) and Tenebrio molitor L. (Coleoptera:

Tenebrionidae) baiting assays (Chapter 3). A total of 373 strains were isolated and 132 of those were selected for molecular identification. Entomogenous fungi were detected in every field site and the recovered strains had a variety of ecological roles such as phytopathogens, antagonists of phytopathogens, insect antagonists, and saprophytes.

Entomopathogenic fungi (EPF) made up the majority of fungi isolated with the most prevalent genus being Metarhizium, represented by M. robertsii J.F. Bisch., Rehner &

Humber and M. anisopliae (Metschn.). Other genera of potential and confirmed EPF isolates included Beauveria, Penicillium, Pseudogymnoascus, and Purpureocillium

(Paecilomyces). This study was similar to others that reported diverse fungal communities with multiple ecological functions from insect cadavers (Sun and Liu 2008,

Sun et al. 2008, Oliveira et al. 2011), but it was a unique study as it focused on commercial cornfields.

173

Forty-eight fungal strains from Chapter 3 and a commercial strain, Botanigard

22WPâ, Beauveria bassiana (Bals.-Criv.) Vuill. strain GHA, were tested against the WCR in the laboratory (Chapter 4). Those strains were selected to represent the diversity found in the soil and to determine their impact on mortality of WCR third-instar larvae.

In soil assays, fourteen strains caused mortality higher than the negative control: M. anisopliae (n=2), M. robertsii (n=11), Pseudogymnoascus sp. (n=1). Only one strain

(E1089, M. anisopliae) caused mortality significantly higher than BotaniGard, the commercial standard. In the immersion assay, eight strains caused mortality higher than the control: M. anisopliae (n=1), M. robertsii (n=5), Metarhizium sp. (n=1). BotaniGard

(B. bassiana). This study was novel as it tested EPF beyond B. bassiana and M. anisopliae and was able to determine that other pathogens can potentially be explored as BCAs of the WCR.

From the same soil samples of Chapter (3) we also examined the native entomopathogenic nematode (EPN) community (Chapter 5). EPN community showed no seasonal or field variance in the survey project. In 2015, there were no differences in the rate of nematodes or fungi recovered from G. mellonella or T. molitor indicating that both species are suitable baiting hosts for a general community of entomopathogens.

Through the use of a DNA barcoding approach it was determined that Heterorhabditis bacteriophora Poinar and Steinernema spp. are present in the commercial cornfields sampled. Both Heterorhabditis and Steinernema contain species that can infect the WCR

(Geisert et al. 2018, Wright et al. 1993, Journey and Ostlie 2000, Toepfer et al. 2005,

2008, Kurtz et al. 2009). Strains from the commercial fields have not yet been tested

174 against WCR larvae, but an inoculation project was set-up to determine the impacts of commercial and persistent H. bacteriophora and S. feltiae co-inoculation on WCR mortality in the field. Data from the inoculation project didn’t show any EPN treatment effects, potentially due to a high background of native nematodes in the control plots and relatively low infestation level of WCR. Some of the nematodes in the control plots were identified as resident strains of Steinernema feltiae and Steinernema spp. One hypothesis derived from these results is that the Steinernema spp. strains in the control plots were causing WCR mortality and therefore we didn’t see treatment effects.

Application of EPF and EPN individually, as species assemblages or in conjunction with Bt hybrids are able to reduce larval and adult WCR populations (Bruck and Lewis

2001, Journey and Ostlie 2000, Toepfer et al. 2005, 2008, Hoffmann et al. 2014, Kurtz et al. 2009, Meissle et al. 2009, Mulock and Chandler 2001, Petzold-Maxwell et al. 2012a,b,

Pilz et al. 2009, Rudeen et al. 2013, Geisert et al. 2018, Wright et al. 1993). Hence our isolates hold promising potential to be incorporated into inundative or conservation biological control programs in this system. Future studies are needed to determine if the strains isolated here have the requirements needed in a good BCA such as virulence, host specificity, compatibility with agrochemicals, and environmental persistence

(Glazer 1996, Kaya and Koppenhöfer 1996). If BCA(s) meet field innundation or conservation requirements, then biological control can become a reality within the WCR integrated pest management framework.

Cornfields sampled in this study are high-input systems intensely managed with

Bt hybrids, insecticides, nematicides, fungicides, herbicides, and fertilizers. Despite

175 these practices, natural enemies above ground (arthropods) and below-ground

(entomopathogens) are present in our system. Moreover, Cry3Bb1 WCR resistance has been documented in the counties where we conducted our studies (Wangila et al. 2015,

Wangila and Meinke 2016, Reinders 2017). In 2014, Field A (Keith Co.) and Field C

(Perkins Co.) had single trait Cry3Bb1 hybrids and high densities of WCR in those fields suggest some level of resistance may occur at those sites.

These findings beg the question: if all these natural enemies exist in cornfields, and a lot of them have been shown to kill WCR, then why is the WCR such a problem in

Nebraska?

The first contributing factor is the wide adoption of corn monocultures. Keith and Perkins Counties, Nebraska are high yielding counties in the state (USDA-NASS

2018). High densities of WCR are able to build up in these areas as continuous corn is grown on a regional level, not just in isolated fields (USDA-NASS, 2017). In addition,

WCR have evolved to suppress corn defenses and have high reproductive capabilities

(Robert et al. 2012). All these factors together, greatly favor large WCR populations in the area even though natural enemies are present in the same habitats. Hence, the sheer magnitude of WCR density enables the WCR to overcome both biotic and abiotic mortality factors and therefore maintain economically challenging densities over time in many continuous cornfields.

Western corn rootworms spend the majority of their life cycle in the soil and are most likely adapted to the overall soil environment, including environmental stressors and natural enemies. Natural enemies and hosts are constantly co-evolving in a way to

176 cause or evade mortality. For instance, although not present in North American corn hybrids, corn roots can emit the insect-induced volatile, (E)- b- caryophyllene, to recruit

EPNs to attack WCR larvae (Rasmann et al. 2005), but, WCR larvae sequester benzoxazinoids from plant roots and activate it upon EPN attack, killing the nematodes and its associated symbiotic bacteria required to kill the insect (Robert et al. 2017).

Moreover, physical protection from natural enemies can happen while WCR larvae feed within the roots (Strnad and Bergman 1987) or as they pupate in earthen cells (Chiang

1973). Some predators are also deterred by a sticky hemolymph defense that larvae possess when facing predation (Lundgren et al. 2010).

Western corn rootworm trophic interactions are not well understood, and so far, no keystone predator or pathogen has been identified. Nevertheless, studies that targeted WCR larvae with entomopathogens (single or multi-species compositions) together with Bt hybrids have shown the potential for corn plant protection (Meissle et al. 2009, Petzold-Maxwell et al. 2012a,b; Petzold–Maxwell et al. 2013, Rudeen and

Gassmann 2013, Hoffmann et al. 2014). In general, these papers found that entomopathogens can act in additive and complementary ways to Bt to increase WCR mortality and/or increase plant protection. Coupling entomopathogens with existing practices can lead to a more sustainable pest management program over time.

In summary, natural enemies are natural components of agroecosystems and a thorough understanding of factors that influence their success as BCAs is necessary.

Understanding the soil microbial community and arthropod predators can contribute to the exploration of sustainable agriculture practices. This dissertation generated

177 foundation work for biological control in irrigated corn in Nebraska. We determined that a wide range of natural enemies are present in fields including pathogens of the WCR such as M. anisopliae, B. bassiana, H. bacteriophora and Steinernema spp. We also determined that other pathogens that have not been reported before against the WCR,

M. robertsii, Pseudogymnoascus sp., Chaetomium sp., and P. lilacinum can cause WCR mortality. Future research should focus on understanding WCR-pathogen relationships and the suitability of biological control programs to increase the efficacy of native EPF and EPN strains. The results of this dissertation generated questions that should be explored in future studies:

1) Investigate pathogenicity and virulence of native H. bacteriophora and

Steinernema spp. against WCR larvae. This study would help us understand

whether the native EPN are capable of utilizing WCR as a host.

2) Seed treatments of endophytic EPF have reduced insect densities but also

promoted plant health (Sasan and Bidochka 2012, Lopez et al. 2014, Lopez

and Sword 2015, Bamisile et al. 2018). All the EPF from this study came from

the rhizosphere, hence, it would be interesting to characterize root-EPF

relationships to understand how the strains described here can help plant

protection beyond insect protection.

3) Determine modes of action of Pseudogymnoascus sp., Chaetomium sp., and

P. lilacinum against the WCR. Understanding pathogenic and virulence

factors of these strains, and others found in this work, can lead to a better

178

understanding of factors governing insect susceptibility to EPF, and provide

potential sources of insect-resistance genes (Lacey et al. 2015).

4) Study the impacts of standard agronomical practices (fertilizers, herbicides,

nematicides, insecticides, fungicides, tillage, and rotation) on the

entomopathogenic community found from this dissertation. Studies like this

are necessary to make biological control an existing management option for

WCR and other pests.

179

References:

Andow, D. A., R. J. Wright, E. W. Hodgson, T. E. Hunt, and K. R. Ostlie. 2017. Farmers perspectives on resistance in western corn rootworm to CRW-Bt corn in Midwest USA. J. Agric. Ext. Rural Dev. 9: 27–38.

Bamisile, B. S., C. K. Dash, K. S. Akutse, R. Keppanan, and L. Wang. 2018. Fungal endophytes: beyond herbivore management. Front. Microbiol. 9.

Bruck, D. J., and L. C. Lewis. 2001. Adult Diabrotica spp. (Coleoptera: Chrysomelidae) infection at emergence with indigenous Beauveria bassiana (Deuteromycotina: Hyphomycetes). J. Invertebr. Pathol. 77: 288–289.

Chiang, H. C. 1973. Bionomics of the northern and western corn rootworms. Annu. Rev. Entomol. 18: 47–72.

Gassmann, A. J., J. L. Petzold-Maxwell, E. H. Clifton, M. W. Dunbar, A. M. Hoffmann, D. A. Ingber, and R. S. Keweshan. 2014. Field-evolved resistance by western corn rootworm to multiple Bacillus thuringiensis toxins in transgenic maize. Proc. Natl. Acad. Sci. 111: 5141–5146.

Gassmann, A. J., J. L. Petzold-Maxwell, R. S. Keweshan, and M. W. Dunbar. 2011. Field- evolved resistance to Bt maize by western corn rootworm. PLoS ONE. 6: e22629.

Gassmann, A. J., R. B. Shrestha, S. R. K. Jakka, M. W. Dunbar, E. H. Clifton, A. R. Paolino, D. A. Ingber, B. W. French, K. E. Masloski, J. W. Dounda, and C. R. S. Clair. 2016. Evidence of resistance to Cry34/35Ab1 corn by western corn rootworm (Coleoptera: Chrysomelidae): Root injury in the field and larval survival in plant-based bioassays. J. Econ. Entomol. 109: 1872-1880.

Geisert, R. W., D. J. Cheruiyot, B. E. Hibbard, D. I. Shapiro-Ilan, K. S. Shelby, and T. A. Coudron. 2018. Comparative assessment of four steinernematidae and three heterorhabditidae species for infectivity of larval Diabrotica virgifera virgifera. J. Econ. Entomol.

Georgis, R., A. M. Koppenhöfer, L. A. Lacey, G. Bélair, L. W. Duncan, P. S. Grewal, M. Samish, L. Tan, P. Torr, and R. W. H. M. van Tol. 2006. Successes and failures in the use of parasitic nematodes for pest control. Biol. Control, Third International Symposium on Entomopathogenic Nematodes and Symbiotic Bacteria. 38: 103– 123.

Glazer, I. 1996. Survival mechanisms of entomopathogenic nematodes. Biocontrol Sci. Technol. 6: 373–378.

Hoffmann, A. M., B. W. French, S. T. Jaronski, and A. J. Gassmann. 2014. Effects of entomopathogens on mortality of western corn rootworm (Coleoptera:

180

Chrysomelidae) and fitness costs of resistance to Cry3Bb1 maize. J. Econ. Entomol. 107: 352–360.

Journey, A. M., and K. R. Ostlie. 2000. Biological control of the western corn rootworm (Coleoptera: Chrysomelidae) using the entomopathogenic nematode, Steinernema carpocapsae. Environ. Entomol. 29: 822–831.

Kaya, H. K., and A. M. Koppenhöfer. 1996. Effects of microbial and other antagonistic organism and competition on entomopathogenic nematodes. Biocontrol Sci. Technol. 6: 357–372.

Kirk, V. M. 1982. Carabids: minimal role in pest management of corn rootworms. Environ. Entomol. 11: 5–8.

Kuhlmann, U., and W. A. C. M. Van der Burgt. 1998. Possibilities for biological control of the western corn rootworm, Diabrotica virgifera virgifera LeConte, in Central Europe. Biocontrol News Inf. 19: 59N–68N.

Kurtz, B., I. Hiltpold, T. C. J. Turlings, U. Kuhlmann, and S. Toepfer. 2009. Comparative susceptibility of larval instars and pupae of the western corn rootworm to infection by three entomopathogenic nematodes. BioControl. 54: 255–262.

Lacey, L. A., D. Grzywacz, D. I. Shapiro-Ilan, R. Frutos, M. Brownbridge, and M. S. Goettel. 2015. Insect pathogens as biological control agents: Back to the future. J. Invertebr. Pathol. 132: 1–41.

Levine, E., J. L. Spencer, S. A. Isard, D. W. Onstad, and M. E. Gray. 2002. Adaptation of the western corn rootworm to crop rotation: evolution of a new strain in response to a management practice. Am. Entomol. 48: 94–107.

Lopez, D. C., and G. A. Sword. 2015. The endophytic fungal entomopathogens Beauveria bassiana and Purpureocillium lilacinum enhance the growth of cultivated cotton (Gossypium hirsutum) and negatively affect survival of the cotton bollworm (Helicoverpa zea). Biol. Control. 89: 53-60

Lopez, D. C., K. Zhu-Salzman, M. J. Ek-Ramos, and G. A. Sword. 2014. The entomopathogenic fungal endophytes Purpureocillium lilacinum (formerly Paecilomyces lilacinus) and Beauveria bassiana negatively affect cotton aphid reproduction under both greenhouse and field conditions. PloS One. 9: e103891.

Ludwick, D. C., L. N. Meihls, K. R. Ostlie, B. D. Potter, L. French, and B. E. Hibbard. 2017. Minnesota field population of western corn rootworm (Coleoptera: Chrysomelidae) shows incomplete resistance to Cry34Ab1/Cry35Ab1 and Cry3Bb1. J. Appl. Entomol. 141: 28–40.

181

Lundgren, J. G., and J. K. Fergen. 2014. Predator community structure and trophic linkage strength to a focal prey. Mol. Ecol. 23: 3790–3798.

Lundgren, J. G., S. Toepfer, T. Haye, and U. Kuhlmann. 2010. Haemolymph defence of an invasive herbivore: its breadth of effectiveness against predators. J. Appl. Entomol. 134: 439–448.

Meinke, L. J., B. D. Siegfried, R. J. Wright, and L. D. Chandler. 1998. Adult susceptibility of Nebraska western corn rootworm (Coleoptera: Chrysomelidae) populations to selected insecticides. J. Econ. Entomol. 91: 594–600.

Meissle, M., C. Pilz, and J. Romeis. 2009. Susceptibility of Diabrotica virgifera virgifera (Coleoptera: Chrysomelidae) to the entomopathogenic fungus Metarhizium anisopliae when feeding on Bacillus thuringiensis Cry3Bb1-expressing maize. Appl. Environ. Microbiol. 75: 3937–3943.

Mulock, B. S., and L. D. Chandler. 2001. Effect of Beauveria bassiana on the fecundity of western corn rootworm, Diabrotica virgifera virgifera (Coleoptera: Chrysomelidae). Biol. Control. 22: 16–21.

Oliveira, I., J. A. Pereira, T. Lino-Neto, A. Bento, and P. Baptista. 2011. Fungal diversity associated to the olive moth, Prays oleae Bernard: A survey for potential entomopathogenic fungi. Microb. Ecol. 63: 964–974.

Parimi, S., L. J. Meinke, B. Wade French, L. D. Chandler, and B. D. Siegfried. 2006. Stability and persistence of aldrin and methyl-parathion resistance in western corn rootworm populations (Coleoptera: Chrysomelidae). Crop Prot. 25: 269– 274.

Petzold-Maxwell, J. L., X. Cibils-Stewart, B. W. French, and A. J. Gassmann. 2012a. Adaptation by western corn rootworm (Coleoptera: Chrysomelidae) to Bt maize: inheritance, fitness costs, and feeding preference. J. Econ. Entomol. 105: 1407– 1418.

Petzold–Maxwell, J. L., S. T. Jaronski, E. H. Clifton, M. W. Dunbar, M. A. Jackson, and A. J. Gassmann. 2013. Interactions among Bt Maize, entomopathogens, and rootworm species (Coleoptera: Chrysomelidae) in the Field: effects on survival, yield, and root injury. J. Econ. Entomol. 106: 622–632.

Petzold-Maxwell, J. L., S. T. Jaronski, and A. J. Gassmann. 2012b. Tritrophic interactions among Bt maize, an insect pest and entomopathogens: effects on development and survival of western corn rootworm. Ann. Appl. Biol. 160: 43–55.

Pilz, C., S. Keller, U. Kuhlmann, and S. Toepfer. 2009. Comparative efficacy assessment of fungi, nematodes and insecticides to control western corn rootworm larvae in maize. BioControl. 54: 671–684.

182

Rasmann, S., T. G. Köllner, J. Degenhardt, I. Hiltpold, S. Toepfer, U. Kuhlmann, J. Gershenzon, and T. C. J. Turlings. 2005. Recruitment of entomopathogenic nematodes by insect-damaged maize roots. Nature. 434: 732–737.

Reinders, J.D. 2017. Spatial variation in western corn rootworm (Coleoptera: Chrysomelidae) susceptibility to Bacillus thuringiensis corn events in Nebraska. M.S. Thesis. University of Nebraska-Lincoln

Robert, C. A. M., M. Erb, B. E. Hibbard, F. B. Wade, C. Zwahlen, T. C. J. Turlings, and K. Thompson. 2012. A specialist root herbivore reduces plant resistance and uses an induced plant volatile to aggregate in a density-dependent manner. Funct. Ecol. 26: 1429–1440.

Robert, C. A., X. Zhang, R. A. Machado, S. Schirmer, M. Lori, P. Mateo, M. Erb, and J. Gershenzon. 2017. Sequestration and activation of plant toxins protect the western corn rootworm from enemies at multiple trophic levels. eLife. 6: e29307.

Rudeen, M. L., and A. J. Gassmann. 2013. Effects of Cry34/35Ab1 corn on the survival and development of western corn rootworm, Diabrotica virgifera virgifera. Pest Manag. Sci. 69: 709–716.

Rudeen, M. L., S. T. Jaronski, J. L. Petzold-Maxwell, and A. J. Gassmann. 2013. Entomopathogenic fungi in cornfields and their potential to manage larval western corn rootworm Diabrotica virgifera virgifera. J. Invertebr. Pathol. 114: 329–332.

Sasan, R. K., and M. J. Bidochka. 2012. The insect-pathogenic fungus Metarhizium robertsii (Clavicipitaceae) is also an endophyte that stimulates plant root development. Am. J. Bot. 99: 101–107.

Strnad, S. P., and M. K. Bergman. 1987. Distribution and orientation of western corn rootworm (Coleoptera: Chrysomelidae) larvae in corn roots. Environ. Entomol. 16: 1193–1198.

Sun, B.-D., and X.-Z. Liu. 2008. Occurrence and diversity of insect-associated fungi in natural soils in China. Appl. Soil Ecol. 39: 100–108.

Sun, B.-D., H. Yu, A. J. Chen, and X.-Z. Liu. 2008. Insect-associated fungi in soils of field crops and orchards. Crop Prot. 27: 1421–1426.

Toepfer, S., C. Gueldenzoph, R.-U. Ehlers, and U. Kuhlmann. 2005. Screening of entomopathogenic nematodes for virulence against the invasive western corn rootworm, Diabrotica virgifera virgifera (Coleoptera: Chrysomelidae) in Europe. Bull. Entomol. Res. 95: 473–482.

183

Toepfer, S., T. Haye, M. Erlandson, M. Goettel, J. G. Lundgren, R. G. Kleespies, D. C. Weber, G. C. Walsh, A. Peters, R.-U. Ehlers, H. Strasser, D. Moore, S. Keller, S. Vidal, and U. Kuhlmann. 2009. A review of the natural enemies of beetles in the subtribe Diabroticina (Coleoptera: Chrysomelidae): implications for sustainable pest management. Biocontrol Sci. Technol. 19: 1–65.

Toepfer, S., A. Peters, R.-U. Ehlers, and U. Kuhlmann. 2008. Comparative assessment of the efficacy of entomopathogenic nematode species at reducing western corn rootworm larvae and root damage in maize. J. Appl. Entomol. 132: 337–348.

(USDA-NASS) US National Agricultural Statistical Services. 2017. Corn for all purposes 2016. Planted acres by county for selected states. https://www.nass.usda.gov/Charts_and_Maps/graphics/CR-PL-RGBChor.pdf (Accessed 20. April. 2018).

(USDA-NASS) US National Agricultural Statistical Services. 2018. Quick Stats: Corn crop in 2017. https://quickstats.nass.usda.gov/results/5C430FA2-1E94-3C98-A0CB- 5AF511557E51#96E20545-047C-304A-8B63-39CE3B0C0944 (Accessed 20. April.2010)

Wangila, D. S., A. J. Gassmann, J. L. Petzold-Maxwell, B. W. French, and L. J. Meinke. 2015. Susceptibility of Nebraska Western Corn Rootworm (Coleoptera: Chrysomelidae) Populations to Bt Corn Events. J. Econ. Entomol. 108: 742–751.

Wangila, D. S., and L. J. Meinke. 2016. Effects of adult emergence timing on susceptibility and fitness of Cry3Bb1-resistant western corn rootworms. J. Appl. Entomol. 141: 372-383.

Wright, R. J., J. F. Witkowski, G. Echtenkamp, and R. Georgis. 1993. Efficacy and persistence of Steinernema carpocapsae (Rhabditida: Steinemematidae) applied through a center-pivot irrigation system against larval com rootworms (Coleoptera: Chrysomelidae). J. Econ. Entomol. 86: 1348–1354.

Zukoff, S. N., K. R. Ostlie, B. Potter, L. N. Meihls, A. L. Zukoff, L. French, M. R. Ellersieck, B. W. French, and B. E. Hibbard. 2016. Multiple assays indicate varying levels of cross resistance in Cry3Bb1-Selected field populations of the western corn rootworm to mCry3A, eCry3.1Ab, and Cry34/35Ab1. J. Econ. Entomol. 109: 1387- 1398.

APPENDIX 1. FIELD SITES LOCATION AND AGRONOMIC CHARACTERISTICS. Years Bt Proteins Mean GPS in Foliar Soil Expressed WCR Planting Seed Field Coordinates at Conti- Year Hybridb Insecticides Typea for emergence/ Date Treatmentd Center of Field nuous & Fungicides Rootworm plantc Corn DeKalb 2014 Cry3Bb1 48.9 07 May 1 none 41°06'57.64"N Lex DKC52-04 A 5 101°38'55.35"W loam DeKalb Cry3Bb1 + 2015 47.5 03 May 1 none DKC52-84 Cry34/35Ab1 Woodly DeKalb 2014 Cry3Bb1 15.4 28 April 2 none 40°45'12.04"N fine DKC51-19 B >30 101°47'09.25"W sandy Pioneer 2015 Cry34/35Ab1 6.5 30 April 3 none loam 35F50AM1 Chemigation: Bifenthrin (6.4 fl oz/ac) Haxtun DeKalb 2014 Cry3Bb1 33.2 29 April 2 and 40°47'46.03"N fine DKC51-19 C >30 dimethoate 101°45'27.63"W sandy (1 pt/ac) loam on 22 July Pioneer 2015 Cry34/35Ab1 72.7 01 May 3 none P0419AMX DeKalb Cry3Bb1 + 2014 4.2 26 April 1 none 40°46'55.53"N Altvan DKC54-38 Cry34/35Ab1 D 13 101°48'57.68"W loam DeKalb Cry3Bb1 + 2015 4.2 30 April 1 none DKC54-38 Cry34/35Ab1 Pioneer 1.4 2014 none 03 May 3 none 1151AM Haxtun Aerial 40°51'12.52"N fine E 0 application: 101°42'4.78"W sandy Pioneer 2015 Cry34/35Ab1 5.6 17 May 3 Bifenthrin loam 1151AMX (6.4 fl oz/ac) on 26 July 184

aAs determined by the United States Department of Agriculture-Natural Resources Conservation Service Web Soil Survey. bDKC=DeKalb; P=Pioneer cMean WCR/plant: Eight plants/field were monitored with single-plant emergence cages throughout beetle emergence period (July-September). Mean emergence is the cumulative emergence per cage for 11 dates each year. dSeed treatments of Insecticides and Fungicides: 1: Clothianidin (0.50 mg/seed); metalaxyl, prothioconazole, and fluoxastrobin. 2: Clothianidin (0.25 mg/seed); metalaxyl, prothioconazole, and fluoxastrobin. 3: Thiamethoxam (0.25 mg/seed); thiabendazole, fludioxonil, mefenoxam, azoxystrobin, and ethaboxam

185

186

APPENDIX 2: PRELIMINARY TEST OF ENTOMOPATHOGENIC FUNGI AGAINST THE WESTERN BEAN CUTWORM, STRIACOSTA ALBICOSTA SMITH.

Introduction

The western bean cutworm (WBC), Striacosta albicosta Smith, is a significant pest of corn, Zea mays L., and dry beans, Phaseolus spp., in the United States

(Blickenstaff and Jolley 1982, Seymour et al. 2004, Michel et al. 2010). In corn, larval feeding on ears reduces yield by direct consumption and by providing entry points for quality-reducing fungal infections (Seymour et al. 2004, Michel et al. 2010). Sixth-instar larvae drop to the soil and construct earthen chambers in which they overwinter as prepupae in a quiescent state (Michel et al. 2010). Pupation occurs in late May and adult emergence starts in the beginning of July (Seymour et al. 2004). Any application for larval control of the western corn rootworm (WCR), Diabrotica virgifera virgifera

LeConte, in the soil can potentially impact secondary targets that spend part of their lifecycle within the soil matrix, such as S. albicosta. Thus, the objective of this experiment was to screen selected fungal strains that were previously tested against the

WCR (see Chapter 4) to determine their pathogenicity to WBC prepupae in soil cup

assays. 185

Materials and Methods

Western bean cutworm source. Western bean cutworm prepupae were obtained from the laboratory colony of the Agroecosystems Entomology Laboratory (University of

Nebraska-Lincoln, North Platte, NE). Colony was initiated in 2017 from field-collected egg masses from the Nebraska cities: Benkelman, Grant, Brule, North Platte, Kearney,

187

Grand Island, O’Neill, and Scottsbluff. Prepupae (the quiescent stage of the 6th larval instar) utilized in the experiment had been in the prepupal stage for 10-15 days (50-55 days since egg hatch). Individual prepupae were placed into moist play sand (ca. 2 ml/cup) (QUIKRETE® Premium Play Sand®, # 1113) in 59 ml plastic soufflé cups (Solo Cup

Company, Highland Park, IL) with small holes on the lids to allow for ventilation.

Fungal sources and inoculum preparation. The 10 native fungal strains used in this experiment were isolated via Galleria mellonella F. and Tenebrio molitor L. baiting assays of Nebraska soil samples from cornfields (see Chapter 3). Eight Metarhizium robertsii J.F. Bisch. strains, one Metarhizium anisopliae (Metschn.) Sorokin strain, and one Pseudogymnoascus sp. strain were selected from strains previously tested against the WCR in soil cup assays (see Chapter 4). Nine strains were selected because they caused mortality higher than the control for WCR and one strain (E211) was a poor performer for WCR but was selected to check for WBC-specific mortality. Fungal culture and spore suspensions were prepared as described in Chapter 5. In addition to the native EPF strains, Botanigardâ 22WP, Beauveria bassiana (Bals. -Criv.) Vuill. strain GHA

(Arbico Organics, Oro Valley, AZ) was included as a commercial comparison product. All

strains, including Botanigard, had inoculum concentrations of 1x107 viable spores ml-1, 185 with the exception of strains E1000 and E1034, which showed poor germination or sporulation. For those strains, the maximum obtained concentration of viable spores was used (4.2 x 106 spores ml-1 for E1000 and 2.7 x 106 spores ml-1 for E1034).

Pathogenicity screening. Bioassays were conducted in 59 ml cups containing approximately 37 g of moist sand (ca. 2ml /cup) and one prepupa per cup. Each cup

188 received a total of 3 ml of spore suspension topically applied evenly across the cup.

Bioassay cups were sandwiched between café-trays lined with moist paper towels

(100% RH) and kept in an incubator set at 26.3 ± 0.5°C for 9 days. There was only one replication per strain and each strain was tested against 16 insects (15 insects for strains

E1026, E211, and E376 due to low availability of insects). Control insects received 3 ml of 0.1% Tween-80.

Data analysis. Western bean cutworm mortality was analyzed using the FREQ procedure in SAS (SAS Institute Inc., version 9.4, Cary, NC) (O’Rourke and Hatcher 2013). A contingency table between number of dead insects and treatment was created with the

TABLE function. Differences between treatment mortality frequencies were analyzed via two-tailed Fisher’s exact test (McDonald 2014). If one or more cadavers showed fungal growth consistent with gross morphology of genera tested, then fungal growth was considered positive.

Results

Nine strains of Metarhizium spp., Pseudogymnoascus sp. and Botanigard 22 WP were tested against WBC prepupae. Negative control mortality was 6.25%, Botanigard

mortality was 31.3%, Metarhizium spp. strains mortality ranged from 18.8 to 60 %, and 185

Pseudogymnoascus sp. had 33.3% mortality (Fig. A1). Out of all the strains tested

(including Botanigard), six showed external sporulation: E1000, E1022, E1030, E161,

E328, and E380. Treatments, including the control, were not significantly different based on Fisher’s exact test (Pr < P = 0.22).

189

Discussion

Biological control is an understudied area of WBC management, but studies have explored the efficacy of predators and parasitoids of WBC egg masses (Archibald 2017,

Ostdiek 2012). To our knowledge this is the first study to have tested non-microsporidia entomopathogenic fungi against the WBC (Dorhout 2007, Helms and Wedberg 1976).

The relationship between Nosema and WBC is still unclear; however, it is known that

Nosema spp. are frequently found in WBC moths and its infection has been implicated to be one of the causes of cyclical fluctuations of WBC populations (Dorhout 2007;

Hutchison et al. 2011).

The data herein are preliminary, and strain derived mortality were not statistically different from the control. Additional studies with a larger quantity of test insects are needed to determine the impact of those strains on WBC mortality.

However, fungal growth on cadavers confirmed that the WBC is a host for six EPF strains

(E1000, E1022, E1030, E161, E328, and E380) and five of those strains inflicted significant mortality to WCR larvae (Chapter 4). The WBC prepupal larval stage overwinters in the soil and pupates in May, with adult emergence beginning in July

(Seymour et al. 2004). Therefore, there is an overlap in which prepupae and pupae of 185 the WBC and larvae of the WCR are present in the soil at the same time. A strain that can be used for both species simultaneously may benefit fields in which both pests are a problem. Based on the WCR soil assays (see Chapter 4) and the WBC preliminary assays, it would be worth exploring the feasibility of M. robertsii strains E380 and E1022 for the control of both pests in future studies.

190

References:

Archibald, W. R. 2017. Conservation biological control of western bean cutworm: Molecular gut content analysis of arthropod predators, feeding trials for key predators and agricultural surveys for integrated pest management. M.S. Thesis. University of Nebraska – Lincoln. https://digitalcommons.unl.edu/entomologydiss/49

Blickenstaff, C. C., and P. M. Jolley. 1982. Host Plants of Western Bean Cutworm. Environ. Entomol. 11: 421–425.

Dorhout, D. L. 2007. Ecological and behavioral studies of the western bean cutworm (Lepidoptera: Noctuidae) in corn. M.S thesis. Iowa State University, Ames. https://lib.dr.iastate.edu/rtd/14793/

Helms, T. J., and J. L. Wedberg. 1976. Effects of Bacillus thuringiensis on Nosema- infected midgut epithelium of Loxagrotis albicosta (Lepidoptera: Noctuidae). J. Invertebr. Pathol. 28: 383–384.

Hutchison, W. D., T. E. Hunt, G. L. Hein, K. L. Steffey, C. D. Pilcher, and M. E. Rice. 2011. Genetically engineered Bt corn and range expansion of the western bean cutworm (Lepidoptera: Noctuidae) in the United States: A response to greenpeace Germany. J. Integr. Pest Manag. 2: B1–B8.

McDonald, J.H. 2014. Handbook of Biological Statistics (3rd ed.). Sparky House Publishing, Baltimore, Maryland. pp: 77-85

Michel, A. P., C. H. Krupke, T. S. Baute, and C. D. Difonzo. 2010. Ecology and management of the western bean cutworm (Lepidoptera: Noctuidae) in corn and dry beans. J. Integr. Pest Manag. 1: 1–10.

O’Rourke, N., and L. Hatcher. 2013. A step-by-step approach to using SAS for factor

analysis and structural equation modeling, Second Edition. SAS Institute. pp: 356- 185

358.

Ostdiek, D. 2012. Biological control of western bean cutworm promising; further research needed - March 1, 2012. Univ. Nebraska-Lincoln CropWatch. (https://cropwatch.unl.edu/biological-control-western-bean-cutworm- promising-further-research-needed-march-1-2012).

Seymour, R. C., G. L. Hein, R. J. Wright, and J. B. Campbell. 2004. Western bean cutworm in corn and dry beans. University of Nebraska-Lincoln Extension. Lincoln, NE. G2013.

Figure A1. Western bean cutworm mortality from entomopathogenic fungi treatments.

Means not significantly different based on two-tailed Fisher’s exact test (Pr < P = 0.22). 1 91

192

APPENDIX 3: CURRICULUM VITAE CAMILA OLIVEIRA HOFMAN

Camila Oliveira Hofman, Ph.D. Entomology Department, University of Nebraska-Lincoln 103 Entomology Hall, Lincoln, NE, USA 68583-0816 E-mail: [email protected]

Education

Ph.D. Entomology, University of Nebraska-Lincoln. 2014-2018 M.S. Entomology, University of Nebraska-Lincoln. 2011-2013 B.S. Biology, University of Missouri-Columbia. 2006-2010 B.A. Anthropology, University of Missouri-Columbia. 2006-2010

Academic Research Experience

Graduate Research Assistant, Agroecosystems Entomology Lab, North Platte, NE Jan.2014-Present University of Nebraska-Lincoln, Dept. of Entomology Mentors: Drs. Julie A. Peterson and Lance J. Meinke Research goal: Characterization of the natural enemy community, with emphasis on entomopathogens, in pest management of western corn rootworm (Diabrotica virgifera virgifera) in West Central Nebraska. Skills: Experimental design (field and lab); entomopathogen (fungi and nematodes) isolation, identification and inoculation; insect pathology bioassays; mycology laboratory techniques; western corn rootworm rearing and infestation techniques; root feeding damage rating; DNA extraction, polymerase chain reaction (PCR), gel electrophoresis, DNA editing, BLAST and phylogenetic tree analysis.

Graduate Research Assistant, Vector Ecology Lab, Lincoln, NE Aug.2011-Dec.2013 University of Nebraska-Lincoln, Dept. of Entomology Mentor: Dr.Gary L. Hein Research goal: Determine the impact of Wheat streak mosaic virus and Triticum mosaic virus on transmission by Aceria tosichella and virus epidemiology. Skills: Wheat curl mite colony maintenance; virus inoculations; enzyme-linked immunosorbent assay (ELISA); data collection and analysis of laboratory, greenhouse and field experiments.

Undergraduate Research Technician May.2009–July.2011

193

University of Missouri- Columbia, Dept. of Plant Science Mentor: Dr. Deborah L. Finke Research goal: Investigate interactions among the plant pathogen Cereal Yellow Dwarf virus, its aphid vector, Rhopalosiphum padi, and the aphid’s parasitoid Aphidius colemani. Skills: Greenhouse and insect colony maintenance; virus inoculations; collection, sorting and counting of insects; insect behavior experiments, data collection and analysis.

Plant Phylogenetics Lab Jan.2008-May.2009 University of Missouri- Columbia, Biological Sciences Dept. Mentor: Dr. Maria Alejandra Jaramillo Responsibilities: DNA extraction of plants in the genus Piper; PCR and gel electrophoresis; data analysis, input and organization; greenhouse maintenance.

Department of BioChemistry, Proteomics Lab Dec.2006-May.2007 University of Missouri-Columbia, Biochemistry Dept. Mentor: Dr. Jay J. Thelen Responsibilities: Cleaning duties; Arabidopsis seedling counting; data input in Excel; greenhouse maintenance.

Publications

Peer-reviewed

Oliveira-Hofman, C., Victor, V.S., Meinke, L.J., and Peterson, J.A. Diverse community of ground beetles in a corn agroecosystem do not predate key pest. In preparation. To be submitted to Food Webs.

Oliveira-Hofman, C., Adesemoye, A.O., Mollet, K., Wei, H.H., Meinke, L.J., and Peterson, J.A. In review. Characterization of potential fungal entomopathogens in irrigated cornfields of Nebraska. Submitted to Applied Soil Ecology.

Camargo, C., McCullough, C., Oliveira-Hofman, C. and Ribeiro, M. 2017. 2015 Student Debates, Molecular Biology and Entomology: Partnering for Solutions. Detate topic: What is the single best tool for managing pesticide resistance? Pesticide rotation. Invited summary. American Entomologist .63(2): 120.

194

Peterson, J. A., Ode, P. J, Oliveira-Hofman, C. and Harwood, J. D. 2016. Integration of plant defense traits with biological control of arthropod pests: challenges and opportunities. Frontiers in Plant Science. 7: 1794. doi: 10.3389/fpls.2016.01794

Wosula, E., McMechan, A., Oliveira-Hofman, C., Wegulo, S. N., and Hein, G. 2015. Differential transmission of two isolates of Wheat streak mosaic virus by five wheat curl mite populations. Plant Disease. 100(1): 154-158

Oliveira-Hofman, C., Wegulo, S.N., Tatineni, T., and Hein, G.L. 2015. Impact of Wheat streak mosaic virus and Triticum mosaic virus co-infection of wheat on transmission rates by wheat curl mites. Plant Disease. 99(8): 1170-1174. de Oliveira, C.F., E.Y. Long and D.L. Finke. 2014. Negative effect of a pathogen on its vector? A plant pathogen increases the vulnerability of its vector to attack by natural enemies. Oecologia. 174(4): 1169-1177.

Editor-Reviewed

Mollet, K.A., Hirzel, G.E. Oliveira-Hofman, C and JA Peterson. In Review. Performance of seed treatments and in-furrow at-plant insecticides for protection against Cry3Bb1- resistant western corn rootworm, 2016. Submitted to Arthropod Management Tests.

Mollet, K.A., Macedo, J.V., Oliveira-Hofman, C, Hirzel, G.E. and JA Peterson. In Review. Evaluation of seed treatments and at-plant soil insecticides for the control of wireworms in field corn, 2015. Submitted to Arthropod Management Tests.

Thesis de Oliveira, C.F. 2013. Impact of Wheat streak mosaic virus and Triticum mosaic virus on transmission by Aceria tosichella Keifer (Eriophyidae) and virus epidemiology in wheat. M.S. Thesis. University of Nebraska- Lincoln

Published abstracts

Suárez Victor V, C Oliveira-Hofman, KA Mollet & JA Peterson. 2016. Identification of potential predators of the western corn rootworm in maize fields of West Central

195

Nebraska. Proceedings of the 25th International Congress of Entomology. doi: 10.1603/ICE.2016.114783

Oliveira-Hofman C, AO Adesemoye, LJ Meinke & JA Peterson. 2016. Identification of entomopathogenic fungi from West Central Nebraska and pathogenicity against the western corn rootworm. Proceedings of the 25th International Congress of Entomology. doi: 10.1603/ICE.2016.112656

Oral Presentations

Invited

Oliveira-Hofman, C., Adesemoye, T.O., Meinke, L. J. and Peterson, J.A. 2017. Assessment of soil entomopathogens from maize fields of western Nebraska, USA: Can they be useful for western corn rootworm control? 26th International Working Group on Ostrinia and other maize pests (IWGO) Conference. Beijing, China.

Oliveira-Hofman, C., Peterson, J.A., Meinke, L. J. , and Adesemoye, T. 2016. Evaluation of entomopathogenic fungi on mortality of the western corn rootworm and other corn insects. Nebraska Corn Board Grant selection meeting, Lincoln-NE

Hein, G.L., McMechan, A.J., Wosula, E., Oliveira-Hofman, C. 2016. Complexity of virus transmission to wheat by the wheat curl mite, Aceria tosichella. North Central Branch (NCB)-Entomological Society of America (ESA). Cleveland, OH.

Oliveira-Hofman, C., Mollet, K.A., Wei, H-H., Adesemoye, A., Meinke, L.J. and Peterson, J.A. 2015. Evaluation of native entomopathogens from west central Nebraska for use in integrated western corn rootworm management programs. PIE-Section Symposium: Synergy in Agricultural Pest Control: Use of Interdisciplinary Approaches to feed a growing population. Annual meeting ESA. Minneapolis, MN.

Camargo, C., McCullough, C., Oliveira-Hofman, C. and Ribeiro, M. 2015. Detate topic: What is the single best tool for managing pesticide resistance? Pesticide Rotations. Student Debate competition. Annual Meeting ESA. Minneapolis, MN

Extension

196

Peterson, J. A. , Oliveira-Hofman, C. and Goulart Montezano, D. 2016. From beneficials to pests: understanding insects in agroecosystems for conservation and management. Nebraska Extension Women in Agriculture Conference. Grant,NE.

Peterson J.A., Archibald, W.R., Goulart Montezano, D., Oliveira-Hofman, C., Steward, R., Suárez Victor, V., and Mollet, K.A. 2016. Agroecosystems Entomology: Conserving beneficial insects and managing resistant pests. Nebraska Extension Eureka! Conference. Lincoln, NE.

Peterson, J.A., Oliveira-Hofman, C., Adesemoye, A.O. 2015. Understanding Soil Biodiversity for Managing Insect Pests and Plant Pathogens. 11th Annual West Central Water & Crops Field Day. Grant, NE. Student Competitions

Oliveira-Hofman, C., Powers, T.O., Meinke, L.J. and Peterson, J.A. 2017. DNA barcoding of nematodes recovered from soil baiting in Nebraska corn fields. Annual meeting ESA. Denver, CO.

Oliveira-Hofman, C., Adesemoye, T.A. Meinke, L.J, and Peterson, J.A. 2016. Identification of entomopathogenic fungi from West Central Nebraska and their pathogenicity against the western corn rootworm. International Congress of Entomology. Orlando, FL.

Oliveira-Hofman, C., Meinke, L.J., and Peterson, J.A. 2014. Potential biological control agents of the Western corn rootworm (Diabrotica virgifera virgifera) in continuous corn of West central Nebraska. Annual Meeting ESA. Portland, OR. de Oliveira, Camila F., S. N. Wegulo and G.L. Hein. 2014. Impact of co-infection of wheat streak mosaic virus and Triticum mosaic virus on virus transmission rates and wheat curl mite reproduction in the field. NCB-ESA. Des Moines, IA. de Oliveira, Camila F. and G.L. Hein. 2013. Impact of co-infection of Wheat streak mosaic and Triticum mosaic viruses on transmission rates by the wheat curl mite. Annual Meeting ESA. Austin, TX. de Oliveira, Camila F., E.Y. Long and D.L. Finke. 2010. The influence of a plant virus on the interaction between an aphid vector and its parasitoid. Annual Meeting ESA. San Diego, CA.

197

Poster Presentations

Oliveira-Hofman, C., Meinke, L.J., Adesemoye, A.O. and J.A. Peterson. 2018. Screen of entomopathogenic fungi from west central Nebraska against key pests of corn. 9th International IPM symposium. Baltimore, MD.

Daniel, S.R., Oliveira-Hofman, C., Owen, G.M., and Peterson, J.A. 2017. Spider diversity and abundance in Nebraska agroecosystems and the implication for biological control. Kansas Entomological Society. Lincoln NE.

Suárez Victor, V., Oliveira-Hofman, C., Mollet, K.A., and Peterson, J.A. 2016. Identification of potential predators of the western corn rootworm in maize fields of west central Nebraska. International Congress of Entomology. Orlando, FL.

Oliveira-Hofman, C., Daniel, S.R., and Peterson, J.A. 2017. Western corn rootworm integrated pest management. Nebraska Crop Management Conference. Kearney,NE.

Oliveira-Hofman, C., Suárez Victor, V., Mollet, K.A., Adesemoye, A.O., Meinke, L.J., and Peterson, J.A. 2016. Potential predators and entomopathogens of western corn rootworm in continuous cornfields in Nebraska. NCB-ESA. Cleveland,OH.

Oliveira-Hofman, C., Mollet, K. A.,Wei, H-H., Peterson, J. A, and Adesemoye, A. O. 2015. Multi-marker characterization of fungi and bacteria for biological control of western corn rootworm, Diabrotica virgifera virgifera. 10th Plant Growth-Promoting Rhizobacteria (PGPR) Workshop. Liège, Belgium.

Wosula, E.N., McMechan, A.J., de Oliveira, C., and Hein, G.L. 2014. Differential transmission of two strains of Wheat streak mosaic virus by five mite populations. American Phytopathological Society Meeting. Minneapolis, MN. de Oliveira, Camila F. and G.L. Hein. 2013. Impact of wheat streak mosaic and Triticum mosaic viruses on transmission by Aceria tosichella and virus epidemiology. NCB-ESA. Rapid City, SD.

Long, E.Y., C. F. de Oliveira and D.L. Finke. 2012. The cost of being a vector: plant pathogen-mediated interactions between an aphid and its natural enemy. Life Sciences Week. University of Missouri-Columbia.

198

Awards

3rd Place, Graduate Student Poster Competition. 2017. Kansas Ent. Soc.

1st Place, Ph.D. Student Poster Competition. 2016. NCB-ESA Cleveland, OH ($300)

Topic winner and 3rd Place general competition. 2015. Debate topic: What is the single best tool for managing pesticide resistance? Student debate competition. Annual Meeting ESA- Minneapolis, MN. Team members: Camargo, C., McCullough, C., Oliveira- Hofman, C. and Ribeiro, M. Team coach: Dr. Joe Louis.

3rd Place, M.S. Student 10-minute Presentations Competition. 2014. NCB-ESA Des Moines, IA ($100)

3rd Place, M.S. Student Poster Competition. 2013. NCB-ESA Rapid City, SD ($100)

Travel Grants ($1766 total)

Myron H. Swenk Memorial Fund. Dept. of Entomology - UNL. • September.2016 ($185) • October.2015 ($145) • October.2014 ($96) • October.2013 ($164) • June.2013 ($226) NCB Student Travel Scholarship. North Central Branch-ESA. • June.2016 ($250) • March.2014 ($500) Dept. of Entomology - UNL Funds to attend Insect Path. Course. June.2015. ($200)

Research Grants ($18,169 total)

Oliveira-Hofman, C., Peterson, J.A., Meinke, L. J. , and Adesemoye, T. 2016. Evaluation of entomopathogenic fungi on mortality of the western corn rootworm and other corn insects. Nebraska Corn Board. $18,169. Awarded.

Oliveira-Hofman, C. and Peterson,J.A. 2015. Evaluation of native entomopathogens from West Central Nebraska for use in integrated western corn rootworm management programs. North Central Region Sustainable Agriculture Research and Education (NCR- SARE) Graduate Student Grant. $9,999. Declined.

199

Scholarships and Fellowships ($1000 total)

Milton E. Mohr Graduate Fellowship. 2016. ($1000)

Professional Development Workshops

2017 Write Winning Grant Proposals Workshop. University of Nebraska-Lincoln. Lincoln, Nebraska.

Soft Skills workshop: Skills you need to succeed. Offered by the Graduate Student Assembly. University of Nebraska-Lincoln. Lincoln, Nebraska.

2016 Campuswide Workshops for Graduate Teaching Assistants. Sponsored by the Office of Graduate Studies. University of Nebraska-Lincoln. Lincoln, Nebraska.

2015 Insect Pathology Course. Offered by Cornell University and International Organisation for Biological Control. Ithaca,NY.

Teaching Experience

Teaching Assistant, Entomology 806: Insect Ecology. Online Course. Fall 2017 Duties: Content development for homeworks and assignments.

Teaching Assistant, Entomology 116: Insect Identification Fall 2016 Duties: Deliver lectures, engage students in in-class exercises and demonstrations. Answer students’ questions; help with insect collection, specimen identification and preparation techniques.

Teaching Assistant’s Assistant, Entomology 116: Insect Identification Spring 2014 Duties: Assist teaching assistant with class exercises and demonstrations. Answer students’ questions; help with insect collection, specimen identification and preparation techniques.

200

Outreach

2017 • Dept. of Entomology-UNL. Annual BugFest. 2016 • Dept. of Entomology-UNL. Annual BugFest. • Women in Agriculture Conference. West Central Research Station, UNL. Grant, NE • Women in Science UNL Workshops for High School Students. Insects: The good and the bad workshop.

2015 • Women in Science UNL Workshops for High School Students. Insect Pollinators Workshop. • Lawrence Bruner Club Educational Entomology Booth at Lincoln’s Farmer’s Market.

2014 • Dept. of Entomology-UNL. Annual BugFest. • College of Agricultural Sciences and Natural Resources. Weatherfest. Educational Entomology Booth. • Lawrence Bruner Club Educational Entomology Booth at Lincoln’s Farmer’s Market. • AppleJack Festival. Educational Entomology Booth. Nebraska City.

2013 • Dept. of Entomology-UNL. Annual BugFest. • AppleJack Festival. Educational Entomology Booth. Nebraska City.

Professional Memberships

Society for Invertebrate Pathology (July.2015-Present)

International Organization for Biological Control (Nearctic Regional section) IOBC-NRS (Sept.2014-Present)

Lawrence Bruner Club. University of Nebraska-Lincoln (Aug.2011-Present) • Secretary (May.2015- May.2016) • Library Committee Member (Aug.2012- Present) • Education Committee Member (Aug.2012- Present) o Chair 2017-2018 • President (Aug.2012- May.2013) • Student Representative to Faculty (Aug.2013- May.2014) • Fundraising Committee Member (Aug.2012- May.2015)

201

• Outreach Committee Member (Aug.2011-Aug.2012)

Entomological Society of America (ESA) Member (Nov.2010- Present) • North Central Branch Student Affairs Committee (NCB-SAC) (Oct.2015- June.2017)

C.V. Riley Entomology Club. University of Missouri-Columbia (May.2009- May.2011)

Service

Symposium moderator at Urban Pest Management Conference. Feb.2014. Lincoln, NE. Symposium moderator at Urban Pest Management Conference. Feb.2013. Lincoln, NE.

Additional Information

Portuguese: Native language | English: Fluent | Spanish: reading knowledge