Iowa State University Capstones, Theses and Graduate Theses and Dissertations Dissertations
2017 Pest management of soybean aphid and soybean cyst nematode: Host-plant resistance, entomopathogens, and seed-applied pesticides Eric Harold Clifton Iowa State University
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Pest management of soybean aphid and soybean cyst nematode: Host-plant resistance, entomopathogens, and seed-applied pesticides
by
Eric Harold Clifton
A dissertation submitted to the graduate faculty
in partial fulfillment of the requirements for the degree of
DOCTOR OF PHILOSOPHY
Major: Entomology
Program of Study Committee: Aaron J. Gassmann, Co-major Professor Erin W. Hodgson, Co-major Professor Gregory L. Tylka Brad S. Coates Joel R. Coats
The student author and the program of study committee are solely responsible for the content of this dissertation. The Graduate College will ensure this dissertation is globally accessible and will not permit alterations after a degree is conferred.
Iowa State University
Ames, Iowa
2017
Copyright © Eric Harold Clifton, 2017. All rights reserved. ii
TABLE OF CONTENTS
Page ACKNOWLEDGMENTS ...... iv
ABSTRACT ...... v
CHAPTER 1. GENERAL INTRODUCTION ...... 1 Dissertation Organization ...... 1 Soybean Aphid Ecology and Management ...... 1 Soybean Cyst Nematode Ecology and Management ...... 6 Interactions Between Soybean Aphid and Soybean Cyst Nematode...... 9 Entomopathogenic Fungi and Their Ecological Roles ...... 10 Induced Resistance by Microorganisms ...... 15 Focus of Dissertation ...... 20 References Cited ...... 21
CHAPTER 2. EFFECTS OF HOST-PLANT RESISTANCE AND SEED TREATMENTS ON SOYBEAN APHID (APHIS GLYCINES MATSUMURA), SOYBEAN CYST NEMATODE (HETERODERA GLYCINES ICHINOHE), AND SOYBEAN YIELD ...... 32 Abstract ...... 32 Introduction ...... 33 Materials and Methods ...... 37 Results ...... 44 Discussion ...... 47 Conclusions ...... 51 Acknowledgments...... 52 References Cited ...... 53 Tables ...... 59 Figures...... 65 Supplemental Tables ...... 70 Supplemental Figures ...... 77
iii
CHAPTER 3. EFFECTS OF ENDOPHYTIC ENTOMOPATHOGENIC FUNGI ON SOYBEAN APHID AND IDENTIFICATION OF METARHIZIUM ISOLATES FROM AGRICULTURAL FIELDS ...... 78 Abstract ...... 78 Introduction ...... 79 Materials and Methods ...... 82 Results ...... 90 Discussion ...... 91 Acknowledgments...... 97 References Cited ...... 98 Tables ...... 105 Figures...... 106 Supplemental Tables ...... 111 Supplemental Figures ...... 114
CHAPTER 4. GENERAL CONCLUSIONS ...... 115
APPENDIX A: EFFECTS OF HOST-PLANT RESISTANCE AND PESTICIDAL SEED TREATMENT ON SOYBEAN APHID AND SOYBEAN CYST NEMATODE ON SOYBEAN ...... 116 Preface ...... 116 Introduction ...... 117 Materials and Methods ...... 118 Results and Discussion ...... 121 References Cited ...... 125 Tables ...... 128 Figures...... 129
APPENDIX B: MISCELLANEOUS PROJECTS AND CO-AUTHORED PUBLICATIONS ...... 132 iv
ACKNOWLEDGMENTS
I would like to thank my Co-major Professors, Aaron Gassmann and Erin Hodgson, and my committee members, Gregory Tylka, Brad Coates and Joel Coats, for their guidance throughout the course of this research. I would also like to thank Stefan Jaronski for the additional training and encouragement on experiments.
This work was completed thanks to six or more years of guidance and help from fellow lab members, department faculty and staff, particularly Kelly Kyle, Mike Dunbar, Jennifer
Petzold-Maxwell, Melissa Rudeen, Nadya Keweshan, David Ingber, Mike McCarville, Adam
Varenhorst, Greg VanNostrand, Ram Shrestha, Sean Bradley, Benjamin Brenizer, Eric Yu,
Aubrey Paolino, Coy St. Clair, Kenneth Masloski, Christopher Marett, Gregory Gebhart, Mark
Mullaney, and David Soh. I also want to thank all of the undergraduate employees who helped me with the work, particularly Ethan Sprung and Sarah Rueger during the recent years.
To my parents, Jay and Nancy Clifton, for supporting my scientific curiosities. To my siblings, Brian and Jodie, and their partners, Anne and Jake, for their encouragement and the good times we have had together. I would also like to thank my nephew Luke for being a continual source of joy.
I also want to thank all of my friends in Ames and those back home for keeping in touch and for the memories. In no particular order, I would like to thank Edmund N., Lauren L., Ryan
T., Alex W., Holly G., Brian S., Scott P., Austin B., Eddie B., Cesar M., Mike C., Nathan R., Joe
G., Alex L., Jordan B., and everyone else that can’t fit on this page.
Last but not least, I want to thank Jillian and her parents, Steve and Nancy, for the love and support. Thank you for treating me as part of the family and for throwing the best pool parties in Carthage. v
ABSTRACT
Farmers can benefit from tools that are applied at planting and can still manage pest
populations throughout the growing season. To meet these demands, some of the tools include
host-plant resistance and seed-applied pesticides. However, prophylactic applications of
pesticides in a seed treatment may not always be needed to preserve yield. The complex biology
of induced plant defenses could also be exploited to make crops more tolerant to pest injury.
Here we focus on two major soybean pests: soybean aphid, Aphis glycines Matsumura, and
soybean cyst nematode, Heterodera glycines Ichinohe.
One field study was conducted in multiple years and locations with experimental plots to
compare the effects of host-plant resistant cultivars and pesticidal seed treatments on soybean
aphid, soybean cyst nematode, and soybean yield. Complementary studies were performed in
the greenhouse with the same treatments. Host-plant resistance effectively suppressed both soybean pests; however, pesticidal seed treatments were inconsistent.
A laboratory study was performed to explore inducible host plant defenses and its potential impacts on soybean aphid. Fungal entomopathogens were used as plant inoculum and molecular tools to identify isolates of these fungi that naturally occur in agricultural fields.
Fungal entomopathogens could establish as endophytes in soybean, but the fungus Metarhizium brunneum actually increased populations of soybean aphid on inoculated plants. All of the fungal isolates were Metarhizium robertsii, which confirms its prevalence throughout North
American soils. 1
CHAPTER 1. DISSERTATION ORGANIZATION AND LITERATURE
REVIEW
Dissertation organization
The research presented in this dissertation attempts to improve our understanding of the tools used for managing two major pests of soybean, soybean aphid and soybean cyst nematode, in addition to research on entomopathogenic fungi and their roles as endophytes. The dissertation is organized into four chapters. Chapter 1 is a general review of the relevant literature and outlines the research that will be presented in the later chapters. Chapter 2 will report the use of host-plant resistance and pesticidal seed treatments to manage soybean aphid and soybean cyst nematode and to preserve soybean yield. Chapter 3 is based on a laboratory experiment that used entomopathogenic fungi as soybean seed inoculum to determine their potential impacts on soybean aphids. This chapter also reports the identification of entomopathogenic fungi that were obtained from corn and soybean fields in the state of Iowa.
Chapter 4 contains a summary of the findings and the implications of the research on integrated pest management of soybean pests. Appendix A is a supplemental experiment and Appendix B is a summary of co-authored publications outside of my dissertation during my time at Iowa
State University.
Soybean aphid ecology and management
Soybean aphid, Aphis glycines Matsumura (Hemiptera: Aphididae), is an invasive pest of soybean in North America with prevalent populations throughout the Midwestern United States.
Soybean aphid was first observed in Wisconsin during the summer of 2000 and later observed in ten other northcentral states by September that year (Ragsdale et al. 2011). Soybean aphid 2 damages soybean by removing phloem sap from stems, leaves, and developing pods.
Populations of soybean aphid seem to increase more rapidly later in the growing season when plants have an increased concentration of nitrogen in the phloem (Myers et al. 2005). Even at low levels, soybean aphid can interfere with gas exchange and photosynthetic rates (Macedo et al. 2003). When infestations of the pest reach high levels, plants can become stunted, pods can be aborted (Lin et al. 1993), saprophytic molds can grow on honeydew excrement, and the resulting yield losses can be 40% or more (Ragsdale et al. 2007). The soybean aphid is also capable of transmitting multiple plant viruses to soybean and vectoring plant pathogens to other visited crops like snap bean and potato (Hill et al. 2001, Davis et al. 2005, Gildow et al. 2008).
Soybean aphid has a heteroecious holocyclic life cycle, with sexual reproduction on its primary host buckthorn (Rhamnus spp.) during the fall (Ragsdale et al. 2004). Some of the confirmed overwintering hosts for soybean aphid in North America include common buckthorn,
Rhamnus cathartica L., which is an invasive species of European origin, and the native alderleaf buckthorn, Rhamnus alnifolia L’Hér (Voegtlin et al. 2005). Some soybean aphids have been observed on glossy buckthorn, Frangula alnus P. Mill, in Minnesota (Gassmann et al. 2008).
After mating with males in the fall, female oviparae lay their overwintering eggs on buckthorn.
In the North Central states, deposited eggs have been observed on buckthorn in late October through mid-November (Ragsdale et al. 2004). In the spring, nymphs hatch and develop into wingless fundatrices. A few generations of wingless females remain on buckthorn until winged morphs develop and transition to secondary hosts in the summer, which is primarily cultivated soybean. Hatching has been observed in late March, and colonies of soybean aphid may stay on buckthorn until early May, depending on the region and temperatures. 3
Throughout the summer, winged and wingless morphs reproduce asexually for many
overlapping generations (Ragsdale et al. 2004). When soybean aphids first colonize a soybean
field, distribution is highly patchy. These small colonies are probably produced by winged
migrants that fed for a short time, deposited nymphs, and then moved in search of other host
plants in the field or those in other soybean fields. When soybean is in vegetative growth stages,
soybean aphid colonies typically aggregate near the growing points like young trifoliates,
petioles, and stems. As soybean plants mature and begin to set flowers, soybean aphid colonies tend to become more dispersed and they are typically observed on the undersides of mature
leaves, lower stems, lateral branches, and pods. As temperatures drop and photoperiod decreases
in the fall, winged females called gynoparae develop and then leave soybean in search of
buckthorn (Wu et al. 2004). After finding buckthorn and feeding, gynoparae produce nymphs
that will develop into oviparae. The oviparae will be the females that mate with the males on buckthorn.
After soybean aphid established in North America, field studies observed natural enemies
that attack and help to suppress the ability of aphid populations to grow. The natural enemies observed attacking soybean aphid include lady beetles, lacewings, bugs, and flies (Rutledge et al.
2004, Nielsen and Hajek 2005, Mignault et al. 2006). However, the success of these natural
enemies to stall soybean aphid population growth can depend heavily on the composition of the
surrounding landscape, which provides perennial habitats for predators (Gardiner et al. 2009) and
whether broad-spectrum insecticides are applied to a field (Ohnesorg et al. 2009). Aside from
these natural enemies that can attack soybean aphid in a given year, there are parasitoid wasps
being considered as candidates for importation biological control (Heimpel et al. 2004, Kaiser et
al. 2007). 4
Management of soybean aphid in the United States relies on foliar insecticides, with most
products containing organophosphates or pyrethroids. After its arrival to North America,
soybean aphid became one of the first insect pests in the North Central region to regularly impact soybean yields and motivated farmers to use insecticides more often in their fields (Ragsdale et al. 2004, Tilmon et al. 2011). Some soybean growers have even used prophylactic growth-stage- based applications of insecticides instead of scouting for soybean aphid populations, which is a discouraged practice as it could bring unnecessary harm to communities of beneficial insects that attack soybean aphids (Schmidt et al. 2007, Varenhorst and O’Neal 2012). Ragsdale et al.
(2007) recommends scouting soybean fields starting in July and then applying insecticides if populations reach the economic threshold of 250 aphids/plant. When using this recommendation, growers have a greater likelihood of profiting from an insecticide application.
The spraying of inconsequential soybean aphid populations would be a waste of insecticide
product, harm beneficial insects, and could accelerate insecticide resistance.
Soybean growers can also use seed-applied insecticides as a tool for managing soybean aphid. So far, seed treatments have had limited success in managing soybean aphid, mostly due to their limited residual activity and the timing of soybean aphid colonization. Neonicotinoids are one of the more prevalent groups of insecticides that are registered for seed treatments and for a variety of crops (Douglas and Tooker 2015). Some of the common active ingredients found
in these neonicotinoid seed treatments are clothianidin, imidacloprid, and thiamethoxam. Labels
for many neonicotinoid seed treatments on soybean are aimed at early-season pests like bean leaf
beetle, Cerotoma trifurcata (Coleoptera: Chrysomelidae) and wireworms (Melanotus spp.,
Agriotes spp., and Limonius spp.), which usually injure soybean prior to its reproductive stages
(Elbert et al. 2008, Johnson et al. 2009, Myers and Hill 2014). Some labels for neonicotinoid 5
seed treatments on soybean have been updated to include A. glycines as a target pest. However,
there is evidence that soybean aphid populations typically colonize soybean fields after the time
period when insecticidal seed treatments are most effective, which is typically ≤ 55 days after planting (McCarville and O’Neal 2013). McCornack and Ragsdale (2006) measured inconsistent
efficacy of these insecticidal treatments for suppression of soybean aphid populations in the
field. Insecticidal seed treatments may help to delay the establishment of soybean aphid
populations early in the growing season, but thus far there is little evidence to suggest that
neonicotinoid seed treatments can prevent economic damage to soybeans when populations of
the pest typically increase at a rapid pace in later months of the growing season, i.e. late July and
August (McCornack and Ragsdale 2006, Johnson et al. 2008). Furthermore, outbreaks of
soybean aphid populations are unpredictable in a given year and location, which makes it
difficult to predict the potential benefits of an insecticidal seed treatment on soybean (Johnson et
al. 2008, Gardiner et al. 2009). It appears that many soybean growers are using seed-applied
insecticides as an “insurance” approach, with more than 47% of producers using them without
targeting any pest of concern (Douglas and Tooker 2015, Hurley and Mitchell 2017).
Microbial pesticides, including entomopathogenic fungi, have shown potential for
managing aphids in cotton, potatoes, alfalfa, spinach, beans, small grains, and other crops
(Wilding and Perry 1980, Soper and Ward 1981, Feng et al. 1990, 1991, McLeod et al. 1998,
Steinkraus et al. 1997, Dara and Semtner 2001). However, few microbial pesticides have been
used for soybean aphid and succeeded. Entomopathogenic fungi, predominantly Pandora spp.,
were observed on mycosed cadavers of soybean aphids (Nielsen and Hajek 2005, Koch et al.
2010), but those particular fungi are difficult to culture. Furthermore, the unpredictable timing
of aphid infestations and the abiotic environmental factors, e.g. ultraviolet radiation and 6 humidity, could weaken attempts at aphid suppression. Soybean growers are unlikely to use microbial pesticides in this therapeutic manner when chemical-based insecticides are affordable and more dependable for aphid management.
Another tool for managing soybean aphid is the planting of soybean cultivars with host- plant resistance, which has little to no negative effects on the environment (Pedigo and Rice
2009). Soybean-aphid-resistant cultivars were released in 2010 for commercial growers. These first resistant cultivars used the gene Rag1, which stands for “resistance to Aphis glycines”, and uses antibiosis-based resistance to hinder the development of soybean aphids feeding on the plant. Other host-plant resistant cultivars have been developed that contain the genes Rag2,
Rag3, or Rag4, and there are now cultivars containing a pyramid of resistance genes such as
Rag1/Rag2 (Rouf Mian et al. 2008, Zhang et al. 2009). A pyramid of resistance genes provides greater protection from aphids, and this pyramid of traits can help to delay the development of aphid populations with virulent biotypes that could develop on plants with a single resistance gene (Gould 1998, Wiarda et al. 2012).
Soybean cyst nematode ecology and management
Soybean cyst nematode, Heterodera glycines Ichinohe (Tylenchida: Heteroderidae), is an invasive pest in North America and believed to originate from China, Korea, or Japan (Schmitt et al. 2004). It was first discovered in the United States in 1954 and is now distributed in all of the major soybean-producing states. In states like Illinois and Iowa, soybean cyst nematode has been confirmed in all counties. Among the major pests and pathogens that regularly affect soybean yield, soybean cyst nematode is still the most economically damaging (Wrather et al.
2010). 7
Heterodera glycines is a sexually dimorphic obligate endoparasite that infects soybean
roots (Niblack et al. 2006). Soybean is the predominant and preferred host plant for H. glycines,
but alternative host plants have been observed. Some annual weeds found in the Midwestern
United States, including henbit (Lamium amplexicaule), field pennycress (Thlaspi arvense) and purple deadnettle (Lamium purpureum), have supported H. glycines populations in a greenhouse environment (Venkatesh et al. 2000). When eggs hatch in the spring, the second-stage (J2) juveniles seek soybean roots. After reaching host plant roots, the nematode establishes a feeding site called a syncytium within or near the vascular tissue (Johnson et al. 1993). This feeding site will provide nutrients to the nematode for the duration of its life cycle. After three subsequent molts in the feeding site, adult males stop feeding and leave the soybean roots in search of females that remain adhered to the plant. The adult females are swollen and have a characteristic lemon-shaped appearance. After mating is completed, the females continue to grow and develop eggs. Females can produce 500 or more eggs under ideal conditions, with some eggs being pushed outside of her body in a gelatinous matrix. Depending on the region and temperatures throughout the growing season, H. glycines populations in a soybean can produce two to four generations during a single year (Schmitt et al. 2004). The eggs within the dead females (cysts) and the eggs dispersed outside of cysts in the soil can remain dormant for as many as 11 years
(Niblack et al. 2006). Because of their high fecundity and persistence in the environment, H. glycines populations are difficult to manage in fields that produce soybean, even after several years with a non-host crop like maize.
Management of H. glycines and other nematode pests of soybean has included chemical- based nematicides, with fumigants like ethylene bromide and 1,3-dichloropropene (1,3-D), and non-fumigants like aldicarb and ethoprop (Minton et al. 1980, Trevathan and Robbins 1995). 8
Aldicarb and 1,3-dichloroproene are seldom used today because of the risks of environmental
contamination and the current, stricter regulations and registration reviews (Thomas 1996, Davis
et al. 1997). Rather than treating large areas of farmland, newer nematicide products on the
market for H. glycines management utilize a seed coating – also referred to as a seed treatment.
Some of the new nematicidal seed treatments labeled for H. glycines contain a formulation of
pathogenic microorganisms and/or their toxic metabolites, including the bacterium Pasteuria
nishizawae (Pn1), the bacterium Bacillus firmus (I-1582), and there is even a fungicide
fluopyram that has nematicidal activity (Jones et al. 2017, Mourtzinis et al. 2017). Nematicidal
seed treatments like these are attractive to growers because of safer handling; the lower amount
of product needed per hectare and because they do not require the additional equipment and
labor that were previously used for applications of fumigants to the soil.
Management of H. glycines has long relied on resistant cultivars and rotation schemes
that include non-host crops. Soybean growers have access to hundreds of cultivars with host-
plant resistance for H. glycines and the three different source lines include PI 88788, PI 437654
and Peking. Despite these different sources of resistance, roughly 95% of the soybean cultivars that are resistant to H. glycines use the PI 88788 gene (Mitchum 2016, McCarville et al. 2017).
Rotating the sources of resistance in cultivars is encouraged to slow the buildup of H. glycines populations and simultaneously prevent the buildup of virulent biotypes (HG-types) (Niblack
2005, McCarville et al. 2017). Planting of non-host crops does not eliminate H. glycines populations from fields because the cysts and eggs can remain viable in soil for multiple years, but it does slow the buildup of populations compared to fields with two or more successive years
of soybean (Niblack et al. 2006).
9
Interactions between soybean aphid and soybean cyst nematode
Aphis glycines and H. glycines co-occur in a majority of the soybean-producing regions
of North America, and their life cycles on soybean plants overlap for most of the growing season. Heterodera glycines is the leading reducer of yield among soybean pathogens from year
to year (Wrather et al. 2010) and the interactions of H. glycines with aboveground herbivores
could worsen the reductions to yield. . McCarville et al. (2014) suggests that small infestations
of A. glycines could alter host-plant physiology in a way that improves the nutrient quality of soybean for H. glycines or suppresses plant defenses, and this interaction leads to greater numbers of H. glycines compared to plants that did not endure aboveground herbivory. Similar studies on these interactions have suggested that A. glycines infestations could increase or have no effect on H. glycines reproduction, but these studies differed greatly from one another in their methods (Heeren et al. 2011; Hong et al. 2010, 2011; McCarville et al. 2012).
A majority of the studies on plant-herbivore interactions have focused on aboveground defoliators (Johnson et al. 2006), but plants are often attacked at multiple sites, including soil- dwelling pests feeding on roots (Coleman et al. 2004). Despite the spatial separation between the roots in the soil and aboveground leaves, it is well-documented that roots can store some secondary plant compounds that can protect aboveground tissues from herbivores and transport these compounds to distal tissues when plant defenses are triggered (van der Putten et al. 2001).
Also, foliar attacks could trigger plants to reallocate nutrients to the roots and thereby benefit
root-feeding herbivores (Kaplan et al. 2008, 2009). Plant-pathogenic microbes (i.e., fungi and
viruses) also can play roles in the reallocation of plant nutrients and alter plant defenses (Rafiqi
et al. 2012). A later section in the literature review goes into more detail about inducible plant
defenses, with a particular focus on microorganisms. 10
Entomopathogenic fungi and their ecological roles
Entomopathogenic fungi (EPF), particularly those in the genera Metarhizium and
Beauveria, have been widely studied for their services of killing pest insects (Lord 2005). These
fungi can persist in the soil and phylloplanes in natural landscapes and those modified for
agricultural cropping systems (Leger 2008). The infection process begins when asexual conidia
come into contact with an insect cuticle. Numerous enzymes aid with penetration through the
cuticle, followed by hyphal bodies and blastospores forming in the body cavity, which sap nutrients and release toxins (Zimmermann 2007). Fresh conidia develop on external surfaces of the insect cadaver for dispersal in the environment (Hajek and St. Leger 1994). Under ideal conditions, EPF can cause epizootics that help to keep insect populations in check.
Identification of EPF species cannot always rely on the general morphology of conidia, blastospores, and other characters. Molecular tools including polymerase chain reactions (PCR) have revealed that some species of EPF have cryptic morphologies, and revisions to the taxonomy were made (Bischoff et al. 2006). Some PCR primers, including the internal transcribed spacer region (ITS), elongation factor 1-alpha (EF1-α), the largest (RPB1) and
second largest (RPB2) RNA polymerase II subunits, and Beta-tubulin (βt) have helped to
increase the resolution and confidence of the updated phylogeny (Enkerli and Widmer 2010).
For example, the genus Metarhizium now has a clade that serves as the core of the M. anisopliae
complex and includes unique species like M. brunneum, M. robertsii, and M. pingshaense which
used to be identified as the same species of M. anisopliae (Bischoff et al. 2009). This distinct
clade with the four species is often referred to as the Metarhizium PARB clade, where P = M.
pingshaense, A = M. anisopliae, R = M. robertsii, and B = M. brunneum. Similar phylogenetic 11
revisions were made to the fungus B. bassiana which now has polyphyletic clades (Rehner et al.
2005, 2011). Molecular studies also confirmed that EPF in the genus Cordyceps can produce
Beauveria anamorphs (Li et al. 2001).
Here we explore the genus Metarhizium because it naturally occurs in agricultural soils
around the world (Meyling and Eilenberg 2007, Schneider et al. 2012). Metarhizium isolates
collected from insects or soil in one sugarcane field revealed that a majority of the isolates were
identified as M. anisopliae or M. robertsii (Rezende et al. 2015). In order to accurately sample
the diversity of soilborne EPF in an agricultural system, one must consider the methods of
isolation that are used (i.e., baiting with insects or serial dilutions of soil samples) and the
potential bias that can arise from host specificity or selective media. One must also consider
how abiotic factors in the environment, the timing of sampling, and the presence of plants and
their root systems could bias the isolates collected (Hernández-Dominguez and Guzmán-Franco
2017). Wyrebek et al. (2011) observed that Metarhizium spp. may not be randomly distributed
in a soil environment but rather show rhizosphere specificity for certain plants, with M. robertsii
preferring grass roots and M. brunneum preferring the rhizosphere of shrubs and trees. Studies
on Metarhizium isolates that naturally occur in the North Central United States is lacking.
Clifton et al. (2015) collected Metarhizium isolates from corn and soybean fields with bait insects, and all of the isolates were identified as Metarhizium robertsii as part of the research in
this dissertation (see Chapter 3). Although Metarhizium spp. kill arthropods, one isolate of
Metarhizium was obtained from a H. glycines cyst, but there is no evidence that the fungus
infected the nematode, and it may have been trapped inside the cyst while the nematode
developed on the surface of the roots (Carris et al. 1989). 12
As far as we know, Metarhizium spp. have not been observed infecting and killing A.
glycines, but other EPF kill aphids, including A. glycines. Most of EPF observed to cause natural epizootics in A. glycines populations in the field includes Pandora spp., Zoophthora spp., and
Neozygites spp., with Pandora neoaphidis being the most prevalent (Nielsen and Hajek 2005).
These epizootics from fungal pathogens tend to occur at greater aphid density. Unfortunately, many of the EPF observed on A. glycines cadavers in the field are not easily cultured in the lab for subsequent inundative applications. In order for an epizootic of Pandora spp. to occur the environmental conditions need to be favorable. The Pandora propagules in the landscape typically become infective after a minimum period of 24 hours with >90% humidity (Brobyn et al. 1985).
Some EPF can be easily cultured and applied to cropping systems as a microbial insecticide, also called a mycoinsecticide. Many of the registered strains available to growers belong to the genera Metarhizium and Beauveria, e.g., Green Muscle®, BotaniGard®
Mycotrol® (Kassa et al. 2008). These mycoinsecticides are especially desirable to organic growers that have to maintain their certification and refrain from the use of synthetic pesticides.
Coffee producers in Hawaii can use B. bassiana to help suppress the coffee berry borer,
Hypothenemus hampei Ferrari (Coleoptera: Curculionidae) (Vega et al. 2009). The formulation of EPF products for biological control can vary. Some “technical powders” of pure conidia can be mixed with surfactants and then sprayed on crop foliage, similar to the mixing process used on chemical insecticides in the form of wettable powders. Metarhizium spp. can also be applied with substrates of barley flakes or as microsclerotial granules for the inoculation of potting media or soil (Jackson and Jaronski 2009). Other strains of EPF are not yet registered for sale but are screened in the laboratory or field as potential candidates for registration. Wraight et al. (2010) 13
showed that some unregistered isolates of B. bassiana killed lepidopteran pests of vegetable
crops more quickly than the widely used GHA strain. Rudeen et al. (2013) measured
significantly greater mortality of western corn rootworm with field-collected strains of
Metarhizium compared to the widely used F52 strain. Entomopathogenic fungi could also be
applied in the form of a seed treatment. Corn seeds coated with M. anisopliae conidia reduced
seedling injury by wireworms and increased yield (Kabaluk and Ericsson 2007).
Aside from killing insects, other ecological roles are being described for
entomopathogenic fungi. Vega et al. (2008) provide a review of the literature on the role of
entomopathogenic fungi as fungal endophytes, plant disease antagonists and plant growth
promoters. An endophyte can be defined as a microorganism that can dwell within plant tissues
without harm to the host plant for at least part of the microorganism’s life cycle (De Bary 1866,
Petrini 1991). Beauveria bassiana has been reported as an endophyte (either naturally or
artificially inoculated) in banana, cacao, cocklebur, coffee, cotton, date palm, ironwood,
jimsonweed, maize, Monterey pine, opium poppy, potatoes, white pine and more (Vega 2008,
Brownbridge et al. 2012). Metarhizium spp. have been reported as endophytes in cabbage,
haricot bean, oilseed rape, switchgrass, tomato and more (Hu and St. Leger 2002, Elena et al.
2011, Sasan and Bidochka 2012). Among these studies on Metarhizium and Beauveria
endophytes, some reported that the fungi benefited plant development. For example, M. brunneum increased root hair density and root length for switchgrass (Sasan and Bidochka
2012). Cotton plants inoculated with Beauveria bassiana exhibited greater biomass and more
rapid development of reproductive structures (Lopez and Sword 2015).
In some of the aforementioned studies on Beauveria and Metarhizium acting as endophytes, there were observations they protected host plants from disease. Haricot beans 14 exhibited healthier growth and lower disease indices from root rot, Fusarium solani f. sp. phaseoli, when plants were inoculated with M. robertsii (Sasan and Bidochka 2013). Grapevine leaves inoculated with B. bassiana showed significantly lower disease indices by downy mildew,
Plasmopara viticola (Berk. and Curt.) Berl. and de Toni., compared to the untreated controls
(Jaber 2015). Multiple studies have treated tomato seeds with B. bassiana conidia and improved the health of plants grown in soil containing Rhizoctonia pathogens (Seth 2001, Ownley et al.
2004). While some studies suggest that fungal endophytes improved overall plant vigor and may increase their tolerance to plant pathogens, some of these observations on disease antagonism by
EPF could be explained by enhanced plant defenses.
A review of B. bassiana endophytes and the different impacts on host plants and herbivores is provided by McKinnon et al. (2017). In that review, there was a general pattern where Beauveria bassiana impacted aphids negatively. Also, fava bean was the most widely- studied plant for B. bassiana endophytes in leguminous crops. There appears to be a lack of literature on EPF acting as endophytes in soybean and their potential impacts on herbivores. One study by Russo et al. (2015) inoculated a number of crops, including soybean, but had no insect bioassay. Khan et al. (2012) inoculated soybeans with M. anisopliae and measured greater plant growth under salt-stressed conditions and greater concentration of jasmonic acid (JA). The increased JA concentrations suggest that the plant’s enzymes were reprogrammed in a mutualistic relationship with the fungal endophyte, which suggests that plant defenses might be primed for future attacks by herbivores. However, the signaling pathways for host plant defenses, including JA, are complicated and vary greatly between different host plants. The following section will explore host plant defenses and how they could be manipulated by microorganisms. 15
Induced resistance by microorganisms
Plants are major contributors in food webs and have adapted constitutive defenses that protect them from abiotic stressors (i.e., desiccation and wind), and herbivores and pathogens
(Kaplan et al. 2008). Some adaptations for deterring herbivory and infection can be physical
(i.e., trichomes and waxes) or chemical (i.e., tannins and terpenoids). Plants can activate defense pathways after attacks to elevate overall plant defense, and this response is known as induced or systemic resistance (Sticher et al. 1997). Induced systemic resistance (ISR) is described by
Pieterse et al. (2014) as the induced state of resistance in plants prompted by chemical or biological inducers that primes the plant for future attacks by pathogens or herbivorous insects through an increased state of resistance. Systemic acquired resistance (SAR) is also used to describe similar phenomena, but the term is often used specifically for pathogens acting as the elicitor. The priming of plant defenses can go beyond localized infections and help to protect distal plant tissues (Spoel and Dong 2012). In general terms, the phenomenon of ISR is similar to immunization in mammals but through very different means.
In order for plant defenses to respond to these attacks by pathogens, plants have adapted to distinguish microbial compounds (i.e., fungal chitin) known as pathogen- or microbe- associated molecular patterns (PAMPs or MAMPs) (Zipfel 2009, Pieterse et al. 2014). Similar patterns arise when plants are damaged by enemy invasion or insect herbivory, known as damage-associated molecular patterns (DAMPs) (Boller 2009). In Arabidopsis and rice plants, pattern recognition receptors (PRRs) can recognize fungal pathogens when chitin comes into contact with extracellular spaces in the leaves (de Jonge et al. 2010). Thus, in order for successful infection and establishment to occur, pathogens have adapted mechanisms to bypass 16
PRRs and even suppress plant defenses. For example, the pathogen Cladosporium fulvum secretes an effector protein while invading a host plant that disrupts the defense mechanisms of plant chitinases (van Esse et al. 2007).
Underlying ISR and SAR events are signaling pathways mediated by levels of major plant hormones: the shikimate pathway using salicylic acid (SA), the octadecanoid pathway using JA, and the ethylene pathway. Some of the pioneering studies on induced resistance found that tobacco leaves treated with SA had decreased disease symptoms when challenged by tobacco mosaic virus (White 1979), and later in different systems confirmed the correlation between increased SA concentrations and SAR (Malamy et al. 1990, Métraux et al. 1990). In general, SA is associated with SAR whereas JA and ethylene are associated with ISR (van Loon et al. 2006). Jasmonic acid is found in many plants and can regulate the production of proteinase inhibitors that slow the development of pathogens or herbivores (Thaler 1999). Ethylene is a hormone that regulates plant ripening and senescence, but it also can stimulate or enhance defense responses, and its endogenous levels in the plant often correlates with levels of JA
(Thomma et al. 2001). Furthermore, the signaling of one defense pathway can lead to synergy or even antagonism with another defense pathway, but these outcomes are highly variable depending on the elicitors and host plant in question (Bostock 2005). Several reviews exploring induced resistance by microbes go into greater detail about the underlying mechanisms, modes of action and metabolites that make plant defenses so complex (Sticher et al. 1997, van Loon et al.
2006, Pieterse et al. 2014).
Beneficial microbes that promote plant development also are capable of triggering ISR.
Van Peer et al. (1991) revealed that a strain of plant-growth promoting rhizobacteria
(Pseudomonas fluorescens WCS417r) also could enhance plant defenses towards EPF like 17
Fusarium oxysporum. Similar studies with Pseudomonas spp. have found that the bacteria could
enhance cucumber defenses to pathogens in the genus Pythium (Zhou et al. 1994, Chen et al.
1999). Beneficial fungi, including EPF, may enhance plant defenses. Trichoderma spp. have been extensively studied for their ability to antagonize plant-pathogenic fungi and protect a
variety of crops (Harman et al. 2004), and strains have been formulated for mycopesticides
products (Shoresh et al. 2010). The beneficial fungi that colonize plant roots have evolved
mechanisms of immunosuppression that are similar to harmful pathogens that attempt to bypass
PAMPs and MAMPs so that they can form a symbiosis with the plant (Jacobs et al. 2011).
In addition to exploring how EPF can improve plant development, fungi could also trigger plant defenses towards pathogens and herbivores. Based on the “bodyguard” hypothesis
(Elliot et al. 2000), mutualism would occur when the plant readily recognizes a beneficial fungus and provides shelter in exchange for protection. More evidence is needed to support this hypothesis, but in recent years more studies on EPF have been published that consider tritrophic interactions and broaden their hypotheses beyond more traditional approaches that solely examined their potential as biological control agents (Cory and Ericsson 2010). Ownley et al.
(2010) provides an excellent review of endophytic EPF that can suppress plant pathogens. Sasan and Bidochka (2013) showed that endophytic Metarhizium robertsii could protect haricot beans
from root rot disease, but without additional evidence they hypothesized that the results arose
from water-soluble metabolites, competition of the fungi for nutrients and antibiosis (Qi et al.
2010). Other studies have shown that Beauveria spp. can metabolize volatiles in vitro, including ethanol and sesquiterpenes that could be toxic to other microorganisms (Crespo et al. 2008).
Literature that provides evidence of EPF triggering ISR or SAR is lacking. Some studies have shown that inoculation of cotton plants with B. bassiana could induce systemic resistance against 18 bacterial blight and lower disease ratings in the foliage, but more research is needed (Griffin
2007, Ownley et al. 2008).
Endophytic EPF may enhance plant defenses towards herbivores. When fungi have established endophytically in plants, some of the genes encoding jasmonic acid and ethylene pathways used in defense responses are induced (Vos et al. 2015). Beauveria bassiana is often used as a model EPF for studies on endophytism and the potential impacts on herbivores. Many of these studies with Beauveria used lepidopteran pests and found either negative or neutral impacts on pest fitness (McKinnon et al. 2017), but few studies provide evidence of induced plant defenses. When date palm was inoculated with B. bassiana, proteins related to plant defenses and stress response were induced (Gómez-Vidal et al. 2009). Other studies with endophytic B. bassiana are suggestive that plant defenses could be induced by the fungus.
Cotton aphids, Aphis gossypii, reared on cotton plants inoculated with B. bassiana were shown to have slower population growth compared to the aphids reared on plants in the untreated control
(Lopez et al. 2014).
As stated in the earlier section on EPF, most of the studies on fungal endophytes and/or inducible defenses that examined legume crops have focused on fava bean or common bean, and the relevant literature for soybean is lacking (Vallad and Goodman 2004, McKinnon et al. 2017).
One study by Dann et al. (1998) treated field plots of soybean with 2,6-dichloroisonicotinic acid
(INA), a chemical related to SA, resulting in reduced symptoms of white mold disease
(Sclerotinia sclerotiorum). Another study inoculated soybean with M. anisopliae and observed greater plant development under conditions of salt stress compared to the untreated control
(Khan et al. 2012). Russo et al. (2015) inoculated a number of crops with B. bassiana, including soybean, and measured a low amount of endophyte recovery from soybean leaves in younger 19
plants. Altogether, these studies on soybean have provided some indication that EPF could
establish as endophytes and promote plant development, but comprehensive studies that include pest populations and plant growth parameters are needed.
Microbes can have a positive impact on plant defenses, but sometimes the opposite occurs. Another term for this state of suppressed defenses is induced susceptibility. Beneficial mycorrhizal fungi and bacteria secrete effector proteins that, in essence, silence the plant’s
“alarm system” so that they can establish a symbiosis within the plant, but similar adaptations also are found in saprotrophic plant pathogens (Rafiqi et al. 2012). For example, the causal
agent of gray mold disease can release small RNAs to silence the defense pathways in
Arabidopsis and tomato (Weiberg et al. 2013). This process of hijacking plant defenses has also
been observed in insect herbivores, including aphids (Hillwig et al. 2016). Soybean aphid
feeding can alter fatty acid metabolism and thereby affect the precursors to jasmonate-dependent
defenses (Kanobe et al. 2015). Another study showed some evidence that A. glycines can
obviate plant defenses conferred by the Rag1 gene. When an inducer population of A. glycines is put on resistant soybean and isolated from the rest of the plant, the aphids that are later added to a different region of the plant reach significantly greater numbers compared to the aphids on the control plants lacking an inducer population (Varenhorst et al. 2015). One question is whether microbes could induce plant defenses in a way that affects A. glycines reproduction or prevents the hijacking of plant defenses. In order for ISR to benefit soybean and affect A glycines, plants could be inoculated at an early time point (i.e., time of planting or inoculation of seedlings), but the altered state of plant defenses would have to sustain itself before aphid infestations occur at a later time.
20
Focus of Dissertation
The research presented in this dissertation explores the management of soybean aphid
and soybean cyst nematode with the tools of host-plant resistance and seed-applied pesticides.
We determine whether one tool was sufficient to suppress either pest or if both tools used together would provide added suppression and greater preservation of yield. We also explore whether entomopathogenic fungi can establish endophytically in soybean when they are applied as a seed-soaking inoculum. Additionally, we determine how these fungi may affect soybean aphid populations on inoculated plants. Lastly, we identify cryptic species of Metarhizium that were collected from soils of agricultural fields in Iowa. 21
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32
CHAPTER 2. EFFECTS OF HOST-PLANT RESISTANCE AND SEED
TREATMENTS ON SOYBEAN APHID (APHIS GLYCINES
MATSUMURA), SOYBEAN CYST NEMATODE (HETERODERA
GLYCINES ICHINOHE), AND SOYBEAN YIELD
A paper to be submitted to Pest Management Science
Eric H. Clifton, Gregory L. Tylka, Aaron J. Gassmann, and Erin W. Hodgson
Abstract
BACKGROUND
Soybean cyst nematode, Heterodera glycines, and soybean aphid, Aphis glycines, are invasive, widespread and economically important pests of soybean, Glycines max, in North America.
Management of these pests relies primarily on use of pesticides and soybean germplasm with genetic resistance. A three-year field study and complementary greenhouse experiment were conducted to determine the effects of host-plant resistance and pesticidal seed treatments on pest populations and soybean yield.
RESULTS
Host-plant resistance significantly decreased the abundance of A. glycines and, in most study sites, suppressed H. glycines. Neonicotinoid seed treatment reduced A. glycines abundance on the aphid- and nematode-susceptible cultivar, but abamectin nematicide seed treatment had no 33 effect on H. glycines populations in the field or greenhouse. Seed treatments inconsistently improved soybean yield.
CONCLUSION
These results suggest that the seed treatments included in our experiments may suppress pests and preserve yields, but not consistently for all soybean cultivars or locations. Ultimately, host- plant resistance was more consistent for reducing pest numbers compared to the use of pesticidal seed treatments. The planting of host-plant resistant cultivars should be a main tool in the integrated pest management strategies aimed at both soybean pests.
Keywords: soybean aphid, soybean cyst nematode, yield, host-plant resistance, thiamethoxam, abamectin
1. Introduction
Soybean cyst nematode, Heterodera glycines Ichinohe (Tylenchida: Heteroderidae), and soybean aphid, Aphis glycines Matsumura (Hemiptera: Aphididae), are both invasive pests that can reduce soybean yield (Niblack et al. 2006, Tilmon et al. 2011). Heterodera glycines and A. glycines co-occur in many soybean-producing regions of the United States, where 34 million hectares of soybeans were grown in 2015 at a value of over $37 billion (Niblack et al. 2006,
Ragsdale et al. 2011, McCarville et al. 2012). Heterodera glycines has been present in North
America for more than 60 years and continues to be one of the leading suppressors of soybean yield among plant diseases and pests (Wrather et al. 2010). Aphis glycines has been present in 34
North America since 2000 and yield losses from this pest can be as high as 40% when
populations are not managed (Ragsdale et al. 2007).
One tool that can help manage crop pests is host-plant resistance, which for insects, functions through mechanisms of antibiosis, antixenosis, tolerance, or combinations of these mechanisms, and is conferred through genetically heritable traits (Dogimont et al. 2010). Since the discovery of H. glycines in North America in 1954, management of this pest has relied on soybean germplasm with genetic resistance (Winstead 1955, Niblack 2005). Some of the main sources of H. glycines-resistant germplasm include Peking, PI88788, and PI90763, with PI88788 found in the great majority of soybean cultivars that are currently labeled as resistant to H. glycines (Shannon et al. 2004, McCarville et al. 2017).
Management of H. glycines with host-plant resistance has been complicated in recent years due to the evolution of virulent biotypes to the most commonly used resistance gene,
PI88788 (McCarville et al. 2017). This trend likely resulted from farmers relying exclusively on
a single form of host-plant resistance. For growers in the North Central United States, >95% of the soybean cultivars with H. glycines resistance use resistance genes from the PI88788 source
(Mitchum 2016, McCarville et al. 2017). Populations of H. glycines virulent to PI88788,
according to the H. glycines HG type test (Niblack et al. 2002), have been observed in Iowa,
Illinois, Minnesota, Missouri and Ontario (Cary and Diers 2016). Rotation schemes that include
susceptible soybeans (with or without nematicides) and soybeans with different sources of
resistance can help to prevent the buildup of virulent biotypes that have increased reproduction
on resistant cultivars (Niblack 2005, McCarville et al. 2017). If the trend of increasing virulence
is not stalled, the yields of resistant cultivars could become no different than yields of susceptible
cultivars in future years. 35
Host-plant resistance also can be used to manage A. glycines (McCarville et al. 2013). To
date, four genes conferring resistance to A. glycines have been identified, specifically Rag1,
Rag2, Rag3, and Rag4 (Hill et al. 2006, Mian et al. 2008, Zhang et al. 2009). The first soybean cultivars with A. glycines resistance used a single resistance gene, Rag1, and were available in
2010 (Mardorf et al. 2010). Aphid-resistant cultivars are not completely devoid of aphid populations, but A. glycines populations feeding on those cultivars typically have overall lower reproduction (McCarville et al. 2014b). Although small populations of A. glycines may persist on resistant cultivars, one would expect the aphid-susceptible cultivars to have greater numbers of A. glycines in most cases.
Soybean producers have the option to manage A. glycines and H. glycines with insecticides and nematicides applied as seed treatments. Compared to conventional pesticides used on soil or plant foliage, seed treatments are desirable for their seemingly reduced risks to the environment (Monfort et al. 2006). The most widely used class of insecticides that are registered for seed treatments are the neonicotinoids, including clothianidin, imidacloprid, and thiamethoxam, and it is estimated that neonicotinoids make up one-third of the world’s insecticide market (Simon-Delso et al. 2014, Douglas and Tooker 2015). Many neonicotinoid seed treatments on soybean are aimed at early season pests like bean leaf beetle, Cerotoma trifurcata [Coleoptera: Chrysomelidae] and wireworms, Melanotus spp., Agriotes spp., and
Limonius spp. [Coleoptera: Elateridae], which usually injure soybean prior to the reproductive
stages of soybean (Elbert et al. 2008, Johnson et al. 2009, Myers and Hill 2014).
Some labels for neonicotinoid seed treatments on soybean include A. glycines as a target
pest. However, A. glycines populations typically colonize soybean fields after the time period
when insecticidal seed treatments are most effective, which is typically the first 55 days after 36 planting (McCarville and O’Neal 2013). Insecticidal seed treatments may help manage A. glycines populations early in the growing season, but there is little evidence thus far to suggest that neonicotinoid seed treatments can prevent economic damage to soybeans because aphid populations typically increase at a rapid pace in later months of the growing season, such as in late July and August (McCornack and Ragsdale 2006, Johnson et al. 2008). Furthermore, outbreaks of A. glycines populations are unpredictable in a given year and location, which makes it difficult to predict the potential benefits of using an insecticidal seed treatment on soybean, a decision that must be made prior to planting (Johnson et al. 2008, Gardiner et al. 2009).
Seed treatments have been shown to increase soybean stand density in fields containing multiple pests, including nematodes like H. glycines (Gaspar et al. 2014). Abamectin, a nematicide used in some seed treatment products, is not systemic but it moves along the channels of developing roots and interferes with the nervous system of nematodes. Many of the new nematicide seed treatments labeled for H. glycines management utilize pathogenic microorganisms and/or their toxic metabolites, including strains of Pasteuria nishizawae (Pn1) and the bacterium Bacillus firmus (I-1582 (Jones et al. 2017, Mourtzinis et al. 2017).
Integrated pest management (IPM) strategies that include growing soybean cultivars with host-plant resistance to manage both H. glycines belowground and A. glycines aboveground can protect crops with low environmental risk (Ragsdale et al. 2011, McCarville et al. 2014a). The co-occurrence of both H. glycines and A. glycines in the North Central region of the United
States is also of importance because there is evidence that A. glycines feeding may enhance the quality of soybean as a host plant for H. glycines (McCarville et al. 2014a). If feeding of both pests has additive and negative impacts on soybean yield, the planting of soybean cultivars with stacked resistance genes for both pests would be a good strategy for preserving yield. For 37
soybean cultivars that lack resistance genes, a pesticidal seed treatment may help to provide some pest suppression and consequently preserve yield, however, the question remains as to whether or not this gain in yield would be sufficient to compensate for the cost of the seed treatment. Under high numbers of insect and nematode pests, the addition of a pesticidal seed treatment to resistant cultivars may be profitable. However, in other situations pesticidal seed treatments may add unnecessary costs. Among soybean growers, 47-65% are using pesticidal seed treatments without considering the target pest for which the seed treatment is intended
(Douglas and Tooker 2015).
To address the effects of host-plant resistance and seed treatments on soybean pests, and to understand the value of these tactics for integrated pest management in soybean production, we conducted a field and laboratory study to address the following hypotheses:
1) Soybean cultivars with host-plant resistance will support lower populations of A. glycines
and H. glycines.
2) A pesticidal seed treatment containing an insecticide and nematicide will provide added
suppression of A. glycines and H. glycines.
3) Soybean cultivars with host-plant resistance will yield more than susceptible cultivars.
4) Soybean with pesticidal seed treatment will have increased stand density and greater yield.
2. Materials and Methods
2.1 Field plot experiments and pest sampling
Small-plot soybean field experiments were conducted for three years (2013, 2014, and
2015) at two locations each year for a total of six unique study sites. The Northeast study sites
were located at the Iowa State University’s Northeast Research and Demonstration Farm near 38
Nashua, Iowa. The Central and Northwest study sites were located in Ames and Newell, Iowa, respectively, and were on privately owned farms. The study sites within a location were changed
annually. The Northeast study site was planted on May 16, 2013; May 20, 2014; and May 12,
2015. The Central study site was planted on June 7, 2013, and May 23, 2015. The Northwest
study site was planted on May 17, 2014.
At each study site, plots were established in a randomized complete block design with
four blocks, 12 treatments, replicated eight times, for a total of 96 plots per study site. Plots
contained one of 12 treatments from a fully-crossed factorial design with four soybean cultivars
and three levels of seed treatment. Originally, we intended to include a foliar insecticide
application for A. glycines as an additional factor in the experiment; however, A. glycines
populations never reached the economic threshold of 250 aphids plant-1 before the R5
developmental stage in soybean. No insecticides were sprayed in field experiments. As a result,
the factor of an insecticide application was removed from the experimental design and each
block contained two replications per treatment.
We used four soybean varieties with relative maturities ranging from 2.1 to 2.5, as
indicated by the first two numbers in the following variety names: 1) the cultivar susceptible to
both pests was S24-K2, 2) the A. glycines-resistant (Rag1) cultivar was S25-F2, 3) the H.
glycines-resistant (PI88788) cultivar was S23-P8, and 4) the cultivar with both Rag1 and
PI88788 genes was S21-Q3 (Syngenta AG, Greensboro, North Carolina, U.S.A.). The seed
treatments used in this study were 1) ApronMaxx®, which contained the fungicides mefenoxam
(0.0113 mg AI seed-1) and fludioxonil (0.0038 mg AI seed-1), 2) Avicta Complete®, which
contained the same fungicides in ApronMaxx® at the same rates plus the nematicide abamectin
(0.15 mg AI seed-1) and the neonicotinoid insecticide thiamethoxam (0.0907 mg AI seed-1), and 39
3) seeds that were left untreated (Syngenta AG, Greensboro, North Carolina, U.S.A.). Plots were
planted at a rate of 346,000 seeds hectare-1 using standard farming practices. Plots were 6.10 m
long and four rows with 0.76 m between rows. Plots contained ca. 27 seeds m-1 and there was
0.91 m between plots.
Plots were harvested when at least 95 percent of soybean pods reached maturity. The
Northeast study sites were harvested on October 10, 2013; October 11, 2014; and October 5,
2015. The Central study sites were harvested on October 3, 2013, and October 5, 2015. The
Northwest study site was harvested on October 8, 2014. The center two rows of plots were
harvested using a plot combine. The values for moisture and total seed weight in each plot were
used to determine yield as kilograms hectare-1 at 13% moisture.
Heterodera glycines populations were quantified at both the time of planting and at the
time of harvest by collecting ten soil cores (19 mm in diameter, 15-20 cm in length) from both sides of the two center rows of each soybean plot. Heterodera glycines cysts and eggs were extracted from 100 cc subsamples of dried soil with a modified wet sieving and decanting technique (Gerdemann 1955, McCarville et al. 2012). Cysts are fully developed H. glycines females that contain eggs, and they can be found on roots or in the soil. The suspensions of H. glycines cysts were put on a 250-µm-pore sieve and crushed using water and a motorized rubber stopper to release the H. glycines eggs, which were collected on a 25-µm-pore sieve (Faghihi and
Ferris, 2000). Eggs were suspended in 100 mL water and stained with acid fuchsin before a 1 mL sample representative of the suspension was counted for eggs using a dissecting microscope.
Egg counts were performed twice for each sample and the average H. glycines density per plot was calculated. 40
Aphis glycines populations (both alate and apterous) per plant were counted once per
week from early June (i.e., June 7-11), to early September (i.e., September 6-10). Plants were
sampled at random within each plot. We started with 20 randomly selected plants. However, we
were forced to reduce the number of plants sampled per week later in the season because the
proportion of plants with aphids increased, the plants became larger, and the number of aphids
per plant increased; all of which increased the time required to sample a plot. The number of
plants sampled per plot (average ± standard deviation) were: 18.90 ± 2.14 in June, 11.67 ± 2.41
in July, 5.42 ± 2.30 in August, and 3.00 ± 0.00 in September. The average number of aphids per
plant counted in each plot on the day of data collection was summed over the growing season to
calculate cumulative aphid days (CAD), which serves as an estimate of the season-long aphid
abundance (Ruppel 1983).
The HG type of H. glycines populations in each study site were determined with the
Illinois SCN Type test described by Niblack et al. (2002) that uses a standardized set of soybean
cultivars with different genetic sources of H. glycines resistance. The protocol for HG type tests
was similar to the greenhouse experiment described below.
2.2 Greenhouse experiments on H. glycines populations
We performed 30-day greenhouse experiments modified from the Standard Cyst
Evalution-2008 (SCE-08) protocol to measure treatment effects on a single generation of H.
glycines reproduction for the H. glycines populations in our study sites (Niblack et al. 2009).
The experimental design followed McCarville et al. (2014a). Heterodera glycines typically
completes one generation every 25 to 32 days at 27 to 30 ºC, thus the 30-day experiment
estimates the amount of H. glycines reproduction in one generation (Alston and Shmitt 1988). 41
Because field conditions can greatly impact the reproduction of H. glycines, particularly soil
temperature and rainfall, we wanted to study the H. glycines populations and treatments within a
more controlled environment. The greenhouse experiments used the same 12 treatments of
soybean cultivar × seed treatment that were used in the field experiment. After completing H. glycines egg counts for the soil samples collected at the time of planting, all of the remaining soil from one study site was combined, mixed together and used for the greenhouse experiment. We performed five separate greenhouse experiments, with one experiment per study site. We could not conduct a greenhouse experiment for the Northeast 2015 study site because it had very low densities of H. glycines eggs. Depending on the H. glycines egg densities at the time of planting, the field soil from a study site was diluted with the appropriate amount of construction sand to adjust the H. glycines egg densities to ca. 10 eggs 1 cc soil-1. Single soybean plants were grown
in 125-mL cone-shaped containers (Stuewe & Sons, Tangent, OR), containing 100 mL field soil.
One container for each treatment was placed in a random arrangement inside a 7.5 L
sealed plastic bucket that was filled with construction sand to match the level of the soil inside
each container. In each experiment, six replications (i.e., buckets) were established, with each
bucket containing one container per treatment, for a total of 12 containers per bucket. Buckets
were kept in a temperature-controlled water bath (2.44 m × 1.22 m × 0.30 m; L × W × D) that
stabilizes the soil temperature between 27 and 30 ºC. The greenhouse used for these experiments contained three water baths. A water bath containing buckets and plants was housed on a greenhouse bench such that the tops of buckets were situated ca. 60 cm below 400 W high- pressure sodium growth lamps (16:8 (L:D)). Plants were watered as needed. At the end of the experiment, containers were removed from the buckets and roots were washed with a pressurized faucet sprayer to dislodge H. glycines females (cysts) on a 600-µm–pore sieve placed above a 42
250-µm-pore sieve. The cysts were then washed from the 250-µm-pore sieve into individual 100 mL beakers and later counted using direct microscope observation (Leica S6 E, Leica
Microsystems, Wetzlar, Germany).
2.3 Economic analysis
An economic analysis was conducted to compare the recent values of soybean and the cost of pesticidal seed treatments used in field plot experiments. The market value for soybeans in 2016 was obtained from the National Agricultural Statistics Service (2016) and converted to $
USD per kilogram. The cost of seed treatments for Avicta Complete® was estimated from Tylka
(2017) and converted to $ USD per hectare. A table was made that divides the cost of a seed treatment by the value of soybean (Table 6). The rows represent values for a seed treatment cost, ranging from $15 to $20 USD per acre depending on the product, and those values in the rows increase in increments of $1 USD. Those seed treatment costs were then converted to $ USD per hectare. The columns represent different market values for a bushel of soybeans, which has fluctuated between $6 USD and $14 USD in the last ten years, and those values in the columns increase in increments of $1 USD. Those values for a bushel of soybean were then converted to
$ USD per kg. The values within the table represent the minimum increase in yield for treated
soybeans that would compensate for the added cost of the seed treatment (i.e., the gain threshold
(Pedigo and Rice 2009). For each of the six study sites, we compared the average yield of each
soybean cultivar with the fungicide, insecticide, and nematicide (FIN) seed treatment to the
average yield of untreated soybeans for the same cultivar. These comparisons of yield were
calculated for each of the four cultivars at the six study sites, which will be referred to as site- cultivars (4 cultivars × 6 study sites = 24 site-cultivars total).
43
2.4 Data analysis
Unless otherwise stated, we analyzed data with a mixed-model analysis of variance
(ANOVA) using the PROC MIXED statement in SAS 9.4 (SAS 2017). Random effects were
tested using a log-likelihood ratio statistic (-2 RES log likelihood) based on a one-tailed χ2 test assuming one degree of freedom (Littell et al. 1996). Random factors were removed from the model to increase the statistical power when these factors were not significant at a level of α <
0.25 (Quinn and Keough 2002).
For the field study, we analyzed cumulative aphid days, H. glycines population densities at the time of harvest, soybean stand density, and soybean yield with an ANOVA that had the fixed factors of soybean cultivar, seed treatment, and the interaction of soybean cultivar × seed treatment. The analyses also included the random factors of study site, block (nested within study site), and all interactions of study site and block (nested within study site) with the fixed factors. When a significant interaction was present, pairwise comparisons were made using the
PDIFF statement in PROC MIXED. Pairwise comparisons were based on least-square means with a significance level of P < 0.05 after using the Bonferroni adjustment for multiple comparisons. For analyses of individual study sites (in the supplementary tables), we performed a mixed-model ANOVA with the same fixed factors. and the random factors of block and all interactions of block with the fixed factors. When a significant interaction was present, pairwise comparisons were made using Tukey’s Studentized Range (HSD) Test with a 95% confidence level.
HG types were determined following Niblack et al. (2002). If the average number of H. glycines females per host-plant resistant cultivar was more than 10% of the number of females 44
on the SCN-susceptible cultivar (Williams 82), the population was labeled with the appropriate
HG type (Tables S1 & S3).
For the greenhouse experiments, the number of H. glycines females (cysts) per soybean plant was analyzed with a mixed-model ANOVA that included the fixed factors of cultivar, seed treatment, and the interaction of cultivar × seed treatment. Because all of the study sites had H. glycines populations identified as HG type 2 (Table S1), study sites were combined for the overall analysis with that included the random factors of study site and all interactions of study site with the fixed factors in the model.
3. Results
3.1 Field plot experiments
For cumulative aphid days (CAD), there was a significant interaction between cultivar and seed treatment (Table 1, Fig. 1). For the soybean cultivars that was susceptible to A. glycines and H. glycines (NK S24-K2), plants grown from seeds treated with the insecticide thiamethoxam had significantly lower CAD than untreated soybeans (t138 = 4.71, P = 0.0004)
and those treated with fungicide (t138 = 5.26, P < 0.0001) (Fig. 1). However, for all other
soybean cultivars there were no significant differences among plants with or without seed
treatment (Fig. 1). The factor of cultivar also was significant, with Rag1 cultivars (NK S25-F2
and NK S21-Q3) having significantly lower CAD than the aphid-susceptible cultivars (NK S24-
K2 and NK S23-P8) (Table 1, Fig. 1). Between the two aphid-susceptible cultivars, we observed
that the cultivar with H. glycines resistance (NK S23-P8) had significantly lower CAD than the
cultivar without H. glycines resistance (NK 24-K2) (t69 = 2.95, P = 0.0262). 45
For data on density of H. glycines eggs in the soil, we found a significant effect of cultivar, but no significant effect of seed treatment or interaction between these factors (Table 2).
We also found that resistant cultivars (i.e., those with PI88788) had significantly fewer H. glycines at harvest compared to susceptible cultivars (Table 2, Fig. 2). However, analyses of H. glycines population densities separated by study site revealed that there was not a difference between resistant and susceptible cultivars in the Central 2015 location (Table S2).
We observed a significant effect of cultivar, seed treatment, and a significant interaction between cultivar and seed treatment for stand density of soybeans at the V1 stage (Table 3). The significant interaction arose because H. glycines-susceptible cultivars (S24-K2 and S25-F2) that received seed treatments displayed a greater increases in stand density, compared to the untreated control, than was observed for the H. glycines-resistant cultivars (S23-P8 and S21-Q3(Fig. 3).
However, for yield there was no significant effect of cultivar, seed treatment, or interaction between these factors (Table 4, Fig. 4).
All study sites had overall averages of >500 H. glycines eggs 100-1 cc soil before planting except for the Northeast study site in 2015, which averaged <150 eggs 100-1 cc soil (Table S2).
Because of the low number of H. glycines eggs in the Northeast study site in 2015, an HG type test could not be conducted. All study sites contained H. glycines populations with a biotype considered to be virulent on PI88788 (HG type 2), which is the source of resistance that was in our H. glycines-resistant cultivars (S23-P8 and S21-Q3; Tables S1 and S3). The female index values on PI88788 for each H. glycines population in a study site was calculated by dividing the number of females per plant on the PI88788 cultivar by the number of females per plant on 46
Williams 82, and then multiplying the value by 100. The female indices for these H. glycines
populations on the PI88788 soybean cultivar ranged from 12 to 43% (Table S1).
3.2 Greenhouse study with H. glycines
There was a significant effect of cultivar on the numbers of H. glycines females (cysts) per plant, but no significant effect of seed treatment or the interaction between cultivar and seed
treatment (Fig. 5, Table 5). Overall, there were approximately 20-30% fewer females on the H. glycines-resistant cultivars relative to the susceptible cultivars. Considering each study site and year separately, the numbers of H. glycines females per plant were significantly lower on resistant cultivars, except in the Northeast 2014 and Northwest 2014 study sites (Table S1).
3.3 Economic analysis
As of 2016, soybeans were valued at approximately $0.37 USD per kg (NASS 2016), and the cost of a fungicide, insecticide, and nematicide (FIN) seed treatment ranged from $37.07 to
$49.42 USD per ha. Using the lower estimated cost of $37.07 per ha for the seed treatment, the breakeven point (i.e. gain threshold) would be a yield increase of 110.18 kg per ha when a FIN seed treatment is used (Table 6). Compared to untreated seeds, soybean with FIN seed treatment exceeded the gain threshold in just 9 of the 24 combinations of site by cultivar (Fig. S1).
47
4. Discussion
In this study, we found that host-plant resistance suppressed both A. glycines and H. glycines populations (Figs. 1 & 2). We also found that the neonicotinoid thiamethoxam in the fungicide, insecticide, and nematicide (FIN) seed treatment reduced the cumulative aphid days
(CAD) on one of the aphid-susceptible cultivars (NK S24-K2; Fig. 1). The fungicide and FIN seed treatments improved stand densities for some cultivars, but there was no effect of either seed treatment on yield (Figs. 3 & 4). Additionally, the nematicide abamectin in the FIN seed treatment did not reduce population densities of H. glycines in the field or greenhouse (Figs. 2 &
5). They key result from these field and greenhouse experiments was that host-plant resistant cultivars consistently suppressed populations of both pests.
Our results with A. glycines populations on Rag1 cultivars support the results in previous studies that measured lower A. glycines populations on these resistant cultivars. In one field experiment, A. glycines populations on Rag1 cultivars peaked at a few hundred aphids per plant while susceptible cultivars had thousands of aphids per plant (O’Neal and Johnson 2010).
Another field experiment found that resistant cultivars yielded significantly higher than susceptible cultivars when field plots were exposed to natural populations of A. glycines
(Mardorf et al. 2010). In the same study, there were no significant differences in yield between resistant cultivars and susceptible cultivars in the absence of A. glycines populations (Mardorf et al. 2010). There are now soybean cultivars with the Rag1/Rag2 pyramid of resistance genes and they can suppress A. glycines populations more than single-gene cultivars of Rag1 or Rag2 alone
(McCarville et al. 2014b). 48
The results for H. glycines population densities in the field experiments demonstrate that
H. glycines-resistant soybeans can slow the buildup of H. glycines populations, even when those populations may have some virulence to a source of H. glycines resistance. Heterodera glycines injury does not always cause obvious symptoms to appear in the aboveground biomass, thus many growers would not even know that H. glycines populations are in their field without soil or root sampling (Wang et al. 2003). Heterodera glycines can cause up to a 30% reduction in yield and losses may increase in drier growing seasons (Niblack 2005). All of our study sites contained virulent H. glycines biotypes identified as HG type 2 (Table S3), and yet we measured lower H. glycines reproduction on resistant (PI88788) cultivars compared to the A. glycines- and
H. glycines-susceptible cultivar in the field and greenhouse experiments (Figs. 2 and 5). Virulent
H. glycines biotypes may be more widespread in North America than they were 10 years ago and this trend of increasing virulence could negatively impact soybean production, since host-plant resistant cultivars rely heavily on the PI88788 gene (Cary and Diers 2016, Mitchum 2016,
McCarville et al. 2017). In order to stall the buildup of H. glycines populations and simultaneously prevent the buildup of virulent biotypes, researchers have outlined rotation schemes for growers that should minimize yield losses by utilizing different sources of H. glycines resistance and by planting non-host crops such as corn, alfalfa, wheat, and cotton
(Niblack et al. 2006).
Although the thiamethoxam seed treatment reduced A. glycines populations on soybean cultivar that was susceptible to both A. glycines and H. glycines (Fig. 1), it is difficult to extrapolate from our results whether the same degree of pest suppression would occur A. glycines populations had reached overall greater numbers. In >95% of our field plots, the aphid populations did not reach the economic injury level of 674 aphids plant-1 (Ragsdale et al. 2007). 49
Compared to studies that observed A. glycines populations with >20,000 CAD on soybean
(McCornack and Ragsdale 2006, McCarville et al. 2014b), the A. glycines populations in our
study sites were relatively small and remained <5,000 CAD in most study sites (Fig 1 and Table
S2). Previous studies found that neonicotinoid seed treatments like thiamethoxam do not
consistently prevent A. glycines populations from reaching economically damaging levels
(Magalhaes et al. 2009, Seagraves and Lundgren 2012). This inconsistent reduction in A.
glycines populations with neonicotinoid seed treatments often is due to the timing of A. glycines
colonization, which typically occurs when the concentration of the neonicotinoid compounds in the soybean tissues has diminished greatly (Krupke et al. 2017). Furthermore, thiamethoxam did not provide additional suppression of A. glycines on Rag1 soybeans in our plots, which may have been due, in part, to the overall low A. glycines populations on those Rag1 cultivars. In these situations where host-plant resistance is already being used, an insecticide seed treatment may not be needed to suppress pests and preserve yield.
The abamectin seed treatment had no impact on H. glycines in either the field or greenhouse experiments (Figs. 2 & 5). Abamectin has been reported to suppress root-knot nematodes (Meloidogyne spp.) in cotton and vegetable crops, but otherwise there is little published evidence that it can suppress cyst-forming nematodes including H. glycines (Monfort et al. 2006, Frye et al. 2009, Vitti et al. 2014). In the course of completing this three-year field experiment, new seed treatment products have become available for management of H. glycines, including the bacterium Pasteuria nishizawae (Pn1), the bacterium Bacillus firmus (I-1582), and the fungicide fluopyram, all of which have nematicidal activity (Jones et al. 2017, Mourtzinis et al. 2017). Future studies should test how these nematicides affect H. glycines management and may fit with soybean IPM. 50
The lower populations of A. glycines on the H. glycines-resistant cultivar (NK S23-P8)
may represent an indirect interaction between A. glycines and H. glycines (Fig. 1). Constitutive
plant defenses in resistant cultivars that suppress H. glycines might confer similar effects on aboveground pests like A. glycines, as occurs in other systems (Bezemer and van Dam 2005).
However, it is currently unknown why this effect occurred. Hong et al. (2010) measured a greater intrinsic growth rate of A. glycines on soybeans that were attacked by H. glycines,
implying that feeding by H. glycines increases host-plant quality for A. glycines. Because H. glycines and A. glycines co-occur in many soybean-producing regions, there could be an added benefit to planting H. glycines-resistant soybeans if the belowground nematode pest could positively impact A. glycines populations and vice-versa (McCarville et al. 2014a).
For some soybean cultivars, we measured significantly greater stand density for soybeans with either fungicidal seed treatment or FIN seed treatment compared to untreated soybeans (Fig.
3, Tables 3 & S2). Previous field studies also have found that seed treatments with fungicides, insecticides, and/or nematicides increased the germination and establishment of soybeans in the field (Gaspar et al 2014). Despite our result of greater stand density, seed treatments did not significantly improve soybean yield (Fig. 4, Table 4). Soybean plots with lower stand density appeared to be able to compensate for yield, with greater canopy space per plant likely increasing the growth rate and size of individual plants. Additionally, we expected the cultivar with stacked resistance genes (both Rag1 and PI88788) to have the highest yield compared to the other treatments, however, yield did not differ among the cultivars. However, at higher pest densities host-plant resistance may have a positive effect on yield. 51
According to our economic analysis, the FIN seed treatment increased soybean yield beyond the breakeven point of 100.18 kg per ha for about one-third of the combinations of site by cultivars (Fig. S1, Table 6). Previous studies conducted in the Midwest also have found inconsistent benefits of these seed treatments on soybean for managing pest populations and increasing soybean yield (McCornack and Ragsdale 2006, Seagraves and Lundgren 2012,
Gaspar et al. 2014, Krupke et al. 2017). A review by North et al. (2016) reported how neonicotinoid seed treatments have helped reduce insect pest injury and increased soybean yield in the Mid-South growing region of the United States (e.g., Arkansas, Louisiana, Mississippi, and Tennessee). The southern region of the United States harbors a greater diversity of insect pests than the Midwest and overall has more pest activity early in the soybean-growing season than in the Midwest. In regions like the Mid-Southern United States, insecticidal seed treatments may be more essential tools for soybean IPM than they are in the North Central region.
5. Conclusions
Compared to pesticidal seed treatments, host-plant resistance is a more reliable tool for soybean IPM that can regularly suppress pest populations, especially when dealing with those that are difficult to monitor, such as nematodes in the soil, or those that can rapidly increase in population size, such as aphids (Ajayi-Oyetunde et al. 2016). Suppression of H. glycines with resistant cultivars may indirectly affect A. glycines reproduction through conferred host-plant defenses or alterations to host-plant quality, but further studies are needed on the interactions between these pests (Hong et al. 2010, McCarville et al. 2014a). Pesticidal seed treatments may help reduce A. glycines on susceptible cultivars, but resistant cultivars appear to not benefit from 52
a seed treatments. Furthermore, pesticidal seed treatments may not consistently preserve yield or
help reduce H. glycines populations (Figs. 3, 4, S1). In the North Central region, soybean
growers should consider using cultivars with host-plant resistance to manage A. glycines and H. glycines, but not use seed treatments with the nematicide abamectin or the insecticide thiamethoxam for these pests. If a grower does not have access to A. glycines-resistant cultivars, the most cost effective tool to preserve soybean yield would be scouting and application of foliar
insecticides if A. glycines populations reach the economic threshold (Myers et al. 2005, Hodgson
et al. 2012, Krupke et al. 2017). Soybean production in North America relies heavily on
cultivars with the PI88788 resistance gene. Thus, soybean producers should implement rotation
schemes with non-host crops and different sources of resistance to help delay virulence by H.
glycines to PI88788.
Acknowledgments
David Soh assisted with nematode experiments in the greenhouse and helped with data
collection. Present and past employees of Iowa State University, including Chris Marett, Greg
Gebhart, Stith Wiggs, Mark Mullaney, Greg VanNostrand, Michael McCarville, Adam
Varenhorst, Kiley Mackereth, Sarah Van Tol, Ethan Sprung, Anzelika Marie, Hailey Lambert,
Kate Gundry, and Sarah Rueger, assisted with experiments in the field and lab. This work was partially supported by the Iowa Soybean Association and the soybean checkoff.
53
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59
Tables
Table 1: Mixed model of analysis for cumulative aphid days in the field experiment.
Fixed effect d.f. F-value P-value
Cultivar 3, 69 19.36 <0.0001
Seed treatment 2, 46 3.54 0.0371
Cultivar × seed treatment 6, 138 4.68 0.0002
Random effect d.f. χ2 P-value
Study site 1 126.0 <0.0001
Study site (block) 1 0.4 0.2635
Study site (block) × cultivar 1 99.3 <0.0001
Study site (block) × seed treatment 1 2.3 0.0647
Study site (block) × cultivar × seed treatment 1 3.0 0.0416 60
Table 2: Mixed model of analysis for soybean cyst nematode (Heterodera glycines) population densities at the time of harvest in the field experiment.
Fixed effect d.f. F-value P-value
Cultivar 3, 69 11.96 <0.0001
Seed treatment 2, 46 1.24 0.2996
Cultivar × seed treatment 6, 426 0.72 0.6307
Random effect d.f. χ2 P-value
Study site 1 483.8 <0.0001
Study site (block) 1 240.3 <0.0001
Study site (block) × cultivar 1 3.9 0.0241
Study site (block) × seed treatment 1 4.9 0.0134 61
Table 3: Mixed model of analysis for soybean stand density in the field experiment.
Fixed effect d.f. F-value P-value
Cultivar 3, 69 8.78 <0.0001
Seed treatment 2, 46 14.86 <0.0001
Cultivar × seed treatment 6, 138 6.00 <0.0001
Random effect d.f. χ2 P-value
Study site 1 217.0 <0.0001
Study site (block) 1 8.4 0.0019
Study site (block) × cultivar 1 7.3 0.0034
Study site (block) × seed treatment 1 7.6 0.0029
Study site (block) × cultivar × seed treatment 1 0.1 0.3759 62
Table 4: Mixed model of analysis for soybean yield in the field experiment.
Fixed effect d.f. F-value P-value
Cultivar 3, 69 2.53 0.0646
Seed treatment 2, 46 1.38 0.2614
Cultivar × seed treatment 6, 138 0.75 0.6070
Random effect d.f. χ2 P-value
Study site 1 463.0 <0.0001
Study site (block) 1 443.3 <0.0001
Study site (block) × cultivar 1 32.7 <0.0001
Study site (block) × seed treatment 1 0.6 0.2193
Study site (block) × cultivar × seed treatment 1 3.9 0.0241 63
Table 5: Mixed model of analysis for soybean cyst nematode (Heterodera glycines) females per soybean plant in the greenhouse experiment.
Fixed effect d.f. F-value P-value
Cultivar 3, 12 5.54 0.0127
Seed treatment 2, 8 0.29 0.7555
Cultivar × seed treatment 6, 24 1.44 0.2392
Random effect d.f. χ2 P-value
Study site 1 44.4 <0.0001
Study site × cultivar 1 2.2 0.0690
Study site × seed treatment 1 4.2 0.0202
Table 6: Minimum yield increase, or the breakeven point, in kilograms per hectare that is needed to compensate for the cost of a seed treatment.
Soybean value ($/kg)a
ST ($/ha)b $0.24 $0.28 $0.31 $0.35 $0.37c $0.39 $0.43 $0.47 $0.51 $0.55 $37.07 154.44 132.38 119.56 55.80 100.18d 95.04 86.20 78.86 72.68 67.39 $39.54 164.73 141.20 127.54 69.74 106.85 101.37 91.94 84.12 77.52 71.88 $42.01 175.03 150.03 135.51 75.22 113.53 107.71 97.69 89.38 82.37 76.38 $44.48 185.33 158.85 143.48 97.63 120.21 114.05 103.44 94.63 87.21 80.87 $46.95 195.62 167.68 151.45 111.60 126.89 120.38 109.18 99.89 92.06 85.36 $49.42 205.92 176.50 159.42 125.54 133.57 126.72 114.93 105.15 96.90 89.85 64
a The column headings in bold represent the value of soybeans per kilogram. b The row headings in bold are the costs of a seed treatment (ST) per hectare based on the seeding rate of 346,000 seeds per hectare. c $0.37 USD kg-1 represents the recent market value of soybean. d 100.18 kg ha-1 would be the breakeven amount for the fungicide, insecticide, and nematicide (FIN) seed treatment used in the field experiment, which was estimated to cost at least $37.07 ha-1, when soybean was valued at $0.37 USD kg-1. 65
Figures
Figure 1: Average cumulative aphid days in the field experiment. Bars represent combinations of cultivar × seed treatment. Error bars represent standard error of the means. Bar groups are separated by soybean cultivar. SCN = soybean cyst nematode. SBA = soybean aphid. Shading or patterns in the bars represent seed treatment. F = fungicides; I = insecticide; N = nematicide. Capital letters above the bars represent significant means separation (P < 0.05) by soybean cultivar. Lowercase letters above the bars denotes a significant difference between means for seed treatments within a soybean cultivar. 66
Figure 2: Average number of H. glycines eggs per 100 cc soil at the time of harvest in the field experiment. Bars represent combinations of cultivar × seed treatment. Error bars represent standard error of the means. F = fungicides; I = insecticide; N = nematicide. Bar groups are separated by soybean cultivar. SCN = soybean cyst nematode. SBA = soybean aphid. Shading or patterns in the bars represent seed treatment. Letters above the bars represent significant means separation (P < 0.05) by soybean cultivar. 67
Figure 3: Average soybean stand density in the field experiment. Bars represent combinations of cultivar × seed treatment. Error bars represent standard error of the means. Bar groups are separated by soybean cultivar. SCN = soybean cyst nematode. SBA = soybean aphid. Shading or patterns in the bars represent seed treatment. F = fungicides; I = insecticide; N = nematicide. Capital letters above the bars represent significant means separation (P < 0.05) by soybean cultivar. Lowercase letters above the bars denotes a significant difference between means for seed treatments within a soybean cultivar. 68
Figure 4: Average soybean yield in the field experiment. Bars represent combinations of cultivar × seed treatment. Error bars represent standard error of the means. Bar groups are separated by soybean cultivar. SCN = soybean cyst nematode. SBA = soybean aphid. Shading or patterns in the bars represent seed treatment. F = fungicides; I = insecticide; N = nematicide. 69
Figure 5: Average number of H. glycines females per plant after 30 days in the greenhouse experiments. Bars represent combinations of cultivar × seed treatment. F = fungicides; I = insecticide; N = nematicide. Error bars represent standard error of the means. Bar groups are separated by soybean cultivar. SCN = soybean cyst nematode. SBA = soybean aphid. Shading or patterns in the bars represent seed treatment. Letters above the bars represent significant means separation (P < 0.05) by soybean cultivar.
Supplemental Tables
Supplementary Table 1: The number of soybean cyst nematode females (cysts) on single soybean plants grown in the greenhouse for 30 days. Uppercase letters following the value in a column represent separation of means for soybean cultivars for one study site and lowercase letters following the value in one column represent separation of means for the combinations of cultivar × seed treatment based on Tukey’s Studentized Range (HSD) Test using a 95% confidence level. Female index Year Study site HG-type on PI88788 Cultivara Seed treatmentb SCN females per plantc 2013 Northeast 2.5.7 12% S24-K2 Untreated 140.67 (18.28) Aa ApronMaxx 149.33 (11.78) Aa Avicta Complete 134.33 (13.40) Aa S23-P8 Untreated 68.50 (10.44) Bb ApronMaxx 62.00 (9.18) Bb Avicta Complete 67.00 (9.15) Bb S25-F2 Untreated 156.83 (11.28) Aa ApronMaxx 135.33 (27.03) Aa
Avicta Complete 143.40 (12.68) Aa 70 S21-Q3 Untreated 90.33 (24.41) Ba ApronMaxx 88.00 (7.43) Ba Avicta Complete 86.40 (11.99) Ba 2013 Central 2.5.7 43% S24-K2 Untreated 153.33 (46.53) Aba ApronMaxx 167.33 (24.57) ABa Avicta Complete 229.83 (39.59) ABa S23-P8 Untreated 121.40 (17.10) ABa ApronMaxx 168.33 (19.94) ABa Avicta Complete 167.17 (18.52) ABa S25-F2 Untreated 188.67 (48.86) Aa ApronMaxx 198.83 (26.10) Aa Avicta Complete 232.33 (36.86) Aa S21-Q3 Untreated 126.67 (12.15) Ba ApronMaxx 155.20 (27.18) Ba Avicta Complete 130.00 (26.44) Ba a Cultivar: S24-K2 is susceptible to both SBA and SCN; S23-P8 is resistant to SCN but susceptible to SBA; S25-F2 is resistant to SBA but susceptible to SCN; S21-Q3 is resistant to both SBA and SCN. b Seed treatment: untreated has no chemicals; ApronMaxx has the fungicides fludioxonil and mefenoxam; Avicta Complete has the same fungicides in ApronMaxx in addition to the insecticide thiamethoxam and nematicide abamectin. c SCN females per plant: average number of SCN cysts (females) washed from the roots of a single soybean plant with standard error in parentheses.
Supplementary Table 1 continued Female index Year Study site HG-type on PI88788 Cultivara Seed treatmentb SCN females per plantc 2014 Northeast 2.5.7 17% S24-K2 Untreated 190.83 (27.66) Aab ApronMaxx 214.67 (60.37) Aab Avicta Complete 343.00 (27.57) Aa S23-P8 Untreated 167.50 (43.78) Aab ApronMaxx 155.33 (30.18) Ab Avicta Complete 166.67 (43.73) Aab S25-F2 Untreated 202.80 (29.74) Aab ApronMaxx 207.83 (48.36) Aab Avicta Complete 198.33 (34.39) Aab S21-Q3 Untreated 164.67 (50.82) Aab ApronMaxx 169.00 (8.89) Aab
Avicta Complete 224.75 (16.30) Aab 71 2014 Northwest 2.5.7 19% S24-K2 Untreated 158.83 (21.68) Aa ApronMaxx 168.33 (18.42) Aa Avicta Complete 157.83 (34.83) Aa S23-P8 Untreated 195.50 (39.96) Aa ApronMaxx 186.60 (41.55) Aa Avicta Complete 85.80 (10.48) Aa S25-F2 Untreated 162.17 (48.92) Aa ApronMaxx 165.00 (25.85) Aa Avicta Complete 154.83 (20.52) Aa S21-Q3 Untreated 190.33 (27.55) Aa ApronMaxx 191.00 (22.87) Aa Avicta Complete 115.33 (26.03) Aa
a Cultivar: S24-K2 is susceptible to both SBA and SCN; S23-P8 is resistant to SCN but susceptible to SBA; S25-F2 is resistant to SBA but susceptible to SCN; S21-Q3 is resistant to both SBA and SCN. b Seed treatment: untreated has no chemicals; ApronMaxx has the fungicides fludioxonil and mefenoxam; Avicta Complete has the same fungicides in ApronMaxx in addition to the insecticide thiamethoxam and nematicide abamectin. c SCN females per plant: average number of SCN cysts (females) washed from the roots of a single soybean plant with standard error in parentheses.
Supplementary Table 1 continued Female index Year Study site HG-type on PI88788 Cultivara Seed treatmentb SCN females per plantc 2015 Central 2.5.7 22% S24-K2 Untreated 156.33 (24.94) Aa ApronMaxx 154.50 (15.07) Aa Avicta Complete 181.00 (13.88) A S23-P8 Untreated 87.67 (9.70) Ba ApronMaxx 144.33 (30.64) Ba Avicta Complete 119.83 (17.96) Ba S25-F2 Untreated 129.17 (15.89) ABa ApronMaxx 142.00 (13.94) ABa Avicta Complete 121.00 (32.92) ABa S21-Q3 Untreated 126.67 (12.15) ABa ApronMaxx 155.20 (27.18) ABa
Avicta Complete 130.00 (26.44) ABa 72
a Cultivar: S24-K2 is susceptible to both SBA and SCN; S23-P8 is resistant to SCN but susceptible to SBA; S25-F2 is resistant to SBA but susceptible to SCN; S21-Q3 is resistant to both SBA and SCN. b Seed treatment: untreated has no chemicals; ApronMaxx has the fungicides fludioxonil and mefenoxam; Avicta Complete has the same fungicides in ApronMaxx in addition to the insecticide thiamethoxam and nematicide abamectin. c SCN females per plant: average number of SCN cysts (females) washed from the roots of a single soybean plant with standard error in parentheses.
Supplementary Table 2: The stand counts, spring soybean cyst nematode egg counts, cumulative aphid days, fall soybean cyst nematode egg counts, and soybean yield for all crosses of cultivar × seed treatment separated by study sites in each year. Lowercase letters following the value in one column represent separation of means for one study site in that year based on Tukey’s Studentized Range (HSD) Test using a 95% confidence level. Year Study site Cultivara Seed treatmentb Stand (SE)c Spring SCN (SE)d CAD (SE)e Fall SCN (SE)f Soybean yield (SE)g 2013 Northeast S24-K2 Untreated 74.69 (2.45) ab 1056.25 (295.87) a 3995.94 (777.66) ab 2756.25 (672.98) ab 4262.92 (104.13) abcd ApronMaxx 78.94 (1.99) ab 1068.75 (182.72) a 4963.09 (612.91) a 2506.25 (459.37) ab 4235.16 (78.22) abcd Avicta Complete 79.19 (1.83) ab 1143.75 (248.29) a 2436.46 (301.38) bc 2675.00 (427.20) ab 4355.02 (95.84) abcd S23-P8 Untreated 81.13 (1.96) ab 1275.00 (368.27) a 2079.02 (188.64) c 1206.25 (276.85) ab 4498.81 (40.74) ab ApronMaxx 79.00 (1.48) ab 762.50 (179.97) a 2520.32 (284.16) bc 1631.25 (403.66) ab 4448.05 (35.33) abc Avicta Complete 81.56 (1.43) ab 625.00 (224.20) a 1628.98 (316.68) cd 643.75 (223.29) b 4543.18 (49.95) a S25-F2 Untreated 83.88 (2.34) a 800.00 (182.98) a 389.55 (99.24) d 3331.25 (711.95) a 4078.34 (39.28) d ApronMaxx 74.75 (1.40) ab 950.00 (224.01) a 156.45 (30.59) d 2768.75 (1.60) ab 4078.56 (100.53) d Avicta Complete 84.38 (2.72) a 806.25 (162.96) a 201.08 (42.61) d 2937.50 (778.66) ab 4201.05 (59.65) bcd S21-Q3 Untreated 74.75 (2.90) ab 787.50 (128.43) a 454.42 (122.14) d 831.25 (276.77) ab 4071.03 (53.88) d ApronMaxx 82.50 (1.86) a 693.75 (123.72) a 206.15 (35.78) d 1468.75 (687.32) ab 4157.60 (49.06) cd 73 Avicta Complete 72.63 (1.56) b 1162.50 (367.88) a 233.58 (58.39) d 818.75 (188.49) ab 4154.74 (35.28) cd 2013 Central S24-K2 Untreated 76.13 (1.44) a 2981.25 (645.65) a 668.31 (56.87) a 9631.25 (1931.85) a 2385.15 (140.04) ab ApronMaxx 78.44 (2.02) a 1643.75 (192.59) a 517.37 (57.68) ab 12012.50 (2358.91) a 2428.96 (105.35) ab Avicta Complete 79.69 (1.23) a 2750.00 (893.38) a 417.62 (36.14) b 10731.25 (2115.97) a 2508.73 (125.69) ab S23-P8 Untreated 76.50 (2.30) a 2281.25 (437.32) a 498.84 (62.79) ab 4487.50 (637.92) a 2875.87 (115.34) a ApronMaxx 77.75 (2.18) a 2831.25 (844.68) a 506.30 (72.30) ab 5650.00 (1072.51) a 2894.59 (130.74) a Avicta Complete 72.38 (1.22) a 2156.25 (612.48) a 381.48 (53.53) b 4806.25 (1433.73) a 2866.06 (211.37) a S25-F2 Untreated 76.75 (2.02) a 2431.25 (747.19) a 41.51 (9.15) c 9668.75 (1455.74) a 2063.71 (214.11) b ApronMaxx 71.38 (1.97) a 1493.75 (285.43) a 58.78 (12.67) c 8737.50 (1464.57) a 2210.66 (216.70) ab Avicta Complete 79.18 (2.00) a 2293.75 (773.99) a 37.66 (11.11) c 10706.25 (2247.63) a 2304.14 (160.57) ab S21-Q3 Untreated 72.81 (1.63) a 1768.75 (341.99) a 63.24 (6.73) c 4968.75 (558.13) a 2555.50 (139.34) ab ApronMaxx 76.44 (1.55) a 2318.75 (688.55) a 44.35 (7.02) c 5056.25 (1078.63) a 2607.79 (108.1) ab Avicta Complete 73.88 (1.14) a 2931.25 (817.28) a 58.35 (15.90) c 6531.25 (1760.93) a 2398.53 (95.82) ab
a Cultivar: S24-K2 is susceptible to both soybean aphid (SBA) and soybean cyst nematode (SCN); S23-P8 is resistant to SCN but susceptible to SBA; S25-F2 is resistant to SBA but susceptible to SCN; S21-Q3 is resistant to both SBA and SCN. b Seed treatment: untreated has no chemicals; ApronMaxx has the fungicides fludioxonil and mefenoxam; Avicta Complete has the same fungicides in ApronMaxx in addition to the insecticide thiamethoxam and nematicide abamectin. c Stand: number of standing plants per 3.05 m when soybean plants were at the VC (unifoliate) stage and the standard error in parentheses. d Spring SCN: average number of SCN eggs per 100 cc soil at the time of planting and the standard error in parentheses. e CAD: cumulative aphid days, which is a measure of the seasonal abundance of soybean aphids that plants endured and the standard error in parentheses. f Fall SCN: average number of SCN eggs per 100 cc soil at the time of harvest and the standard error in parentheses. g Soybean yield: average yield in kilograms per hectare and the standard error in parentheses.
Supplementary Table 2 continued Year Study site Cultivara Seed treatmentb Stand (SE)c Spring SCN (SE)d CAD (SE)e Fall SCN (SE)f Soybean yield (SE)g 2014 Northeast S24-K2 Untreated 60.19 (3.05) a 5081.25 (1573.61) a 2088.54 (734.29) a 2856.25 (663.89) a 2803.36 (165.08) b ApronMaxx 66.00 (4.79) a 4737.50 (1711.22) a 1234.49 (295.18) ab 1700.00 (520.82) a 3106.97 (89.48) ab Avicta Complete 69.63 (3.49) a 6087.50 (1670.48) a 552.76 (206.07) ab 3006.25 (692.27) a 3523.49 (285.13) a S23-P8 Untreated 61.63 (2.65) a 4562.50 (1048.76) a 1264.19 (497.16) ab 1843.75 (189.79) a 2733.19 (144.02) b ApronMaxx 69.31 (2.35) a 3531.25 (683.61) a 781.24 (271.96) ab 2131.25 (424.89) a 3079.86 (172.28) ab Avicta Complete 59.81 (1.63) a 4575.00 (1076.91) a 1115.39 (605.45) ab 2062.50 (352.26) a 2990.38 (107.83) ab S25-F2 Untreated 63.56 (2.28) a 3456.25 (754.39) a 246.03 (104.37) b 2987.50 (755.74) a 3114.61 (141.44) ab ApronMaxx 64.75 (3.69) a 3731.25 (894.12) a 193.70 (85.55) b 2356.25 (204.07) a 2879.72 (74.82) ab Avicta Complete 70.81 (2.45) a 4575.00 (637.45) a 490.59 (187.94) ab 3137.50 (586.44) a 3257.53 (120.53) ab S21-Q3 Untreated 62.75 (4.68) a 3418.75 (1148.52) a 284.17 (117.78) b 1275.00 (362.16) a 3044.44 (144.41) ab ApronMaxx 71.13 (2.64) a 3875.00 (1335.87) a 191.30 (73.29) b 1606.25 (315.16) a 3127.10 (88.49) ab Avicta Complete 65.31 (3.57) a 3731.25 (746.72) a 786.67 (496.49) ab 1625.00 (409.59) a 3002.00 (110.07) ab 74 2014 Northwest S24-K2 Untreated 74.50 (3.08) a 3750.00 (568.92) a 188.41 (14.99) a 18262.50 (1991.58) a 3206.99 (428.11) a ApronMaxx 76.13 (2.43) a 6975.00 (1603.12) a 197.14 (19.57) a 14693.75 (3267.30) a 3239.38 (434.26) a Avicta Complete 76.75 (4.45) a 6300.00 (1051.57) a 206.43 (33.68) a 18281.25 (5024.57) a 3237.03 (439.91) a S23-P8 Untreated 69.63 (4.04) a 5168.75 (806.72) a 221.28 (37.20) a 14318.75 (2701.40) a 2760.19 (462.84) a ApronMaxx 73.88 (1.34) a 6275.00 (1534.83) a 155.78 (23.53) a 12518.75 (2577.82) a 3205.41 (531.02) a Avicta Complete 72.75 (2.00) a 3675.00 (847.74) a 157.95 (26.24) a 14000.00 (4595.14) a 2976.65 (513.11) a S25-F2 Untreated 70.63 (2.14) a 5256.25 (870.67) a 47.79 (5.56) b 15987.50 (1745.75) a 3316.31 (419.89) a ApronMaxx 76.75 (1.37) a 4768.75 (1206.45) a 38.24 (7.53) b 18437.50 (4489.59) a 3278.21 (461.33) a Avicta Complete 79.50 (3.00) a 4368.75 (833.56) a 37.33 (8.12) b 22450.00 (3948.25) a 3331.03 (397.27) a S21-Q3 Untreated 66.63 (2.46) a 5825.00 (641.91) a 27.67 (4.46) b 10918.75 (1801.24) a 3141.53 (392.16) a ApronMaxx 72.88 (2.58) a 4100.00 (998.39) a 37.63 (7.87) b 14437.50 (2415.27) a 3066.51 (391.41) a Avicta Complete 72.00 (2.01) a 4412.50 (544.92) a 51.43 (10.46) b 16806.25 (5215.71) a 3257.72 (320.09) a
a Cultivar: S24-K2 is susceptible to both soybean aphid (SBA) and soybean cyst nematode (SCN); S23-P8 is resistant to SCN but susceptible to SBA; S25-F2 is resistant to SBA but susceptible to SCN; S21-Q3 is resistant to both SBA and SCN. b Seed treatment: untreated has no chemicals; ApronMaxx has the fungicides fludioxonil and mefenoxam; Avicta Complete has the same fungicides in ApronMaxx in addition to the insecticide thiamethoxam and nematicide abamectin. c Stand: number of standing plants per 3.05 m when soybean plants were at the VC (unifoliate) stage and the standard error in parentheses. d Spring SCN: average number of SCN eggs per 100 cc soil at the time of planting and the standard error in parentheses. e CAD: cumulative aphid days, which is a measure of the seasonal abundance of soybean aphids that plants endured and the standard error in parentheses. f Fall SCN: average number of SCN eggs per 100 cc soil at the time of harvest and the standard error in parentheses. g Soybean yield: average yield in kilograms per hectare and the standard error in parentheses.
Supplementary Table 2 continued Year Study site Cultivara Seed treatmentb Stand (SE)c Spring SCN (SE)d CAD (SE)e Fall SCN (SE)f Soybean yield (SE)g 2015 Northeast S24-K2 Untreated 80.63 (3.49) bc 175.00 (59.01) a 1415.32 (380.72) ab 12.50 (12.50) a 4609.97 (132.91) a ApronMaxx 90.73 (4.42) ab 87.50 (22.66) a 2051.48 (1319.71) a 175.00 (94.02) a 4762.37 (153.63) a Avicta Complete 96.77 (3.66) a 112.50 (44.07) a 450.21 (69.94) ab 150.00 (50.00) a 4300.28 (195.22) a S23-P8 Untreated 77.19 (1.95) bc 100.00 (46.29) a 339.07 (33.37) ab 237.50 (155.77) a 4878.41 (204.60) a ApronMaxx 81.56 (2.32) bc 100.00 (42.26) a 625.09 (285.81) ab 50.00 (26.73) a 4686.43 (231.74) a Avicta Complete 90.94 (3.05) ab 112.50 (47.95) a 419.90 (54.95) ab 62.50 (26.31) a 4530.32 (179.77) a S25-F2 Untreated 76.46 (2.79) bc 75.00 (31.34) a 72.93 (33.41) b 287.50 (233.33) a 4373.37 (74.97) a ApronMaxx 78.33 (2.44) bc 87.50 (35.04) a 41.14 (10.32) b 75.00 (31.34) a 4324.78 (99.49) a Avicta Complete 87.19 (3.96) abc 112.50 (35.04) a 31.46 (3.90) b 150.00 (88.64) a 4372.70 (83.29) a S21-Q3 Untreated 73.65 (3.82) c 87.50 (29.50) a 410.75 (115.36) ab 50.00 (26.73) a 4611.75 (60.74) a ApronMaxx 79.17 (2.23) bc 62.50 (26.31) a 255.66 (51.35) ab 50.00 (26.73) a 4383.00 (109.45) a Avicta Complete 78.23 (2.16) bc 200.00 (46.29) a 383.38 (95.56) ab 25.00 (16.37) a 4627.38 (129.53) 75 2015 Central S24-K2 Untreated 78.44 (1.60) bc 2806.25 (678.72) a 926.27 (174.18) a 1837.50 (414.01) a 3764.96 (112.96) a ApronMaxx 89.79 (1.54) ab 5043.75 (436.86) a 852.30 (102.79) ab 2262.50 (414.44) a 3608.84 (156.32) a Avicta Complete 93.96 (2.53) a 3531.25 (664.60) a 637.86 (59.91) abcd 1325.00 (228.15) a 3680.30 (191.79) a S23-P8 Untreated 74.38 (2.82) c 4450.00 (743.60) a 495.88 (81.30) abcde 2037.50 (415.73) a 3751.46 (127.59) a ApronMaxx 80.00 (5.08) bc 2943.75 (327.11) a 559.09 (83.99) abcde 1087.50 (239.37) a 3886.58 (89.08) a Avicta Complete 85.83 (3.10) abc 3818.75 (1037.90) a 712.51 (179.04) abc 1512.50 (301.45) a 3687.05 (168.58) a S25-F2 Untreated 76.77 (2.15) c 5543.75 (1201.02) a 169.08 (75.64) de 1787.50 (448.98) a 3574.86 (109.97) a ApronMaxx 77.19 (2.38) c 2593.75 (887.28) a 133.37 (37.58) e 1275.00 (239.61) a 3663.08 (115.73) a Avicta Complete 84.06 (1.57) abc 5406.25 (1722.99) a 76.56 (17.94) e 1425.00 (191.56) a 3639.47 (142.52) a S21-Q3 Untreated 72.81 (1.63) c 3462.50 (963.33) a 179.39 (30.39) de 1412.50 (204.80) a 3811.17 (175.71) a ApronMaxx 76.44 (1.55) c 3881.25 (1126.43) a 360.43 (140.36) cde 1225.00 (220.19) a 3841.88 (180.93) a Avicta Complete 73.88 (1.14) c 5968.75 (1300.75) a 400.65 (96.67) bcde 1437.50 (207.83) a 4081.03 (61.45) a
a Cultivar: S24-K2 is susceptible to both soybean aphid (SBA) and soybean cyst nematode (SCN); S23-P8 is resistant to SCN but susceptible to SBA; S25-F2 is resistant to SBA but susceptible to SCN; S21-Q3 is resistant to both SBA and SCN. b Seed treatment: untreated has no chemicals; ApronMaxx has the fungicides fludioxonil and mefenoxam; Avicta Complete has the same fungicides in ApronMaxx in addition to the insecticide thiamethoxam and nematicide abamectin. c Stand: number of standing plants per 3.05 m when soybean plants were at the VC (unifoliate) stage and the standard error in parentheses. d Spring SCN: average number of SCN eggs per 100 cc soil at the time of planting and the standard error in parentheses. e CAD: cumulative aphid days, which is a measure of the seasonal abundance of soybean aphids that plants endured and the standard error in parentheses. f Fall SCN: average number of SCN eggs per 100 cc soil at the time of harvest and the standard error in parentheses. g Soybean yield: average yield in kilograms per hectare and the standard error in parentheses.
Supplementary Table 3: The numbers of soybean cyst nematode females (cysts) per plant (± SEM) from HG-type tests that characterized populations in our study sites
Study site and year
Cultivar Indicator # CENa 2013 NEb 2013 NWc 2014 NEb 2014 CENa 2015
PI 548402 1 40.0 ± 6.31 3.3 ± 0.42 7.3 ± 1.51 9.0 ± 1.15 10 ± 2.00
PI 88788 2 84.0 ± 2.20 25.9 ± 2.82 35.3 ± 8.50 59.3 ± 16.80 33 ± 4.70
PI 90763 3 2.5 ± 1.3 1.6 ± 0.57 2.4 ± 0.37 2.6 ± 0.78 6.86 ± 4.97
PI 437654 4 0 ± 0 2.1 ± 0.59 0 ± 0 0 ± 0 1.29 ± 0.42
PI 209322 5 68.0 ± 5.34 43.6 ± 9.05 80.7 ± 16.02 68.3 ± 18.16 42.43 ± 5.79 76
PI 89772 6 7.3 ± 1.65 4.5 ± 1.15 1.3 ± 0.52 2.7 ± 0.52 3.33 ± 2.0
PI 548316 7 98.3 ± 8.53 84.9 ± 9.91 100.2 ± 30.35 115.3 ± 23.42 45.43 ± 6.97
Williams 82 or Lee 74d Control 195.25 ± 11.51 224.1 ± 35.77 182.0 ± 36.02 351.5 ± 89.08 152 ± 18.56
HG-type 1.2.5.7 2.5.7 2.5.7 2.5.7 2.5.7 aCEN: Central location bNE: Northeast location cNW: Northwest location dWilliams 82 used as the control for all study sites, except Central 2013 which used Lee 74.
Supplemental Figures 77
Supplementary Figure 1: The differences in average yield for a soybean cultivar with the fungicide, insecticide, and nematicide (FIN) seed treatment minus the average yield for the same cultivar that was left untreated. Each bar represents the mean value for one cultivar and bars are arranged by different study sites. Error bars represent standard error of the means. The red line is the breakeven point of 110 kg ha-1, or the minimum increase in yield, that compensates for the cost of the FIN seed treatment based on the recent value of soybean. SCN = soybean cyst nematode; SBA = soybean aphid; Res = resistant.
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CHAPTER 3. EFFECTS OF ENDOPHYTIC ENTOMOPATHOGENIC
FUNGI ON SOYBEAN APHID AND IDENTIFICATION OF
METARHIZIUM ISOLATES FROM AGRICULTURAL FIELDS
A paper to be submitted to PLoS ONE
Eric H. Clifton, Stefan T. Jaronski, Brad S. Coates, Erin W. Hodgson, and Aaron J. Gassmann
Abstract
Terrestrial plants can harbor endophytic fungi that may induce changes in plants that in turn
affect interactions with herbivorous insects. We evaluated whether the entomopathogenic fungi
Beauveria bassiana and Metarhizium brunneum applied to soybean seeds could establish as endophytes in soybean and affect interactions with soybean aphid (Aphis glycines Matsumura).
We also used DNA sequence data to identify species of Metarhizium obtained from agricultural
fields in Iowa. We found M. brunneum strain F52 increased populations of A. glycines but B.
bassiana strain GHA had no effect. We were able to recover both fungi as endophytes in
soybean, but B. bassiana was more prevalent. Phylogenetic analyses, based on DNA sequence data, for Metarhizium isolates from agricultural fields indicated that all isolates were
Metarhizium robertsii, which is consistent with past studies indicating a cosmopolitan
distribution and wide host range for this species. Our work confirms that some entomopathogenic fungi can be endophytic in soybean, however, these fungi may have a negative effect on the plants by increasing susceptibility of soybean to A. glycines.
Keywords: endophyte, entomopathogenic fungi, induced resistance, soybean aphid, phylogeny
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1. Introduction
An endophyte is a microorganism that can dwell within plant tissues for at least part of
the microorganism’s life cycle (De Bary 1866, Petrini 1991). Some fungal endophytes can
promote plant development and/or suppress plant diseases and herbivorous pests in exchange for
nutrition, refuge and transmission of spores (Saikkonnen et al. 2004). Previous research on
plant-herbivore-endophyte interactions has focused on fungis that can produce toxic alkaloids in
grasses, and subsequently can harm grazing cattle (Cheeke 1995). However these fungal-derived
alkaloids also can deter phloem-feeding insects such as the bird cherry-oat aphid,
Rhopalosiphum padi Linnaeus, and greenbug, Schizaphis graminum Rondani (Hemiptera:
Aphididae) (Siegel et al. 1990, Wilkinson et al. 2000).
Following direct contact with potential insect hosts, entomopathogenic fungi (EPF) infect
by penetrating the cuticular exoskeleton (Zimmermann 2007). Due to the ability to suppress pest
populations and persist in the environment, EPF can serve as an alternative to chemical
insecticides in certain cropping systems (Chandler 2016). Some EPF strains in the genera
Beauveria (Hypocreales: Cordyciptaceae) and Metarhizium (Hypocreales: Clavicipitaceae) are
already registered and marketed as biopesticides (Lacey 2016). Some of the EPF strains used as
biopesticides are known to naturally occur in agricultural fields, including B. bassiana and M.
robertsii, and may sometimes cause epizootics in pest populations (Klingen et al. 2002, Meyling
and Eilenberg 2007). Research also has shown that EPF can be endophytes in a diversity of
crops, including banana, corn, cotton, fava bean, poppy, tobacco, and wheat (Vega et al. 2009).
Entomopathogenic fungi are typically applied to crops using foliar sprays or soil
inoculum, but endophytism can be exploited by treating the seeds or inoculating seedlings via
root dips, leaf sprays, and injection of stems (Quesada-Moraga et al. 2006, Akello et al. 2007,
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Russo et al. 2015). Coating of corn seeds with M. anisopliae sensu lato (s.l.) conidia reduced
seedling injury by wireworms and increased yield (Kabaluk and Ericsson 2007). Endophytic
EPF are hypothesized to establish a zone of protection around the plant roots, increase the rate of
plant development and nutrient uptake, and even induce systemic plant defenses (Sasan and
Bidochka 2012, Barelli et al. 2016). In general terms, induced systemic resistance is defined as the induced state of resistance in plants, prompted by chemical or biological inducers, that primes the plant to defend against future attack by pathogens or herbivorous arthropods (Pieterse et al. 2014). The signaling molecules salicylic acid (SA), jasmonic acid (JA), and ethylene could mediate defense pathways that respond to attacks and prime the plant for future attacks.
Recent studies have shown that endophytic EPF may induce systemic resistance that reduces injury from aphids, spider mites, and other pests (McKinnon et al. 2017). The specific
mechanisms underlying these systemic responses in the plant are unclear, but was suggested that
fungal metabolites could be excreted and transported through plant vasculature, either directly
affecting herbivores or mediating indirect effects by upregulation plant defenses (Jaber and
Ownley 2017). Gomez-Vidal et al. (2009) reported that B. bassiana endophytes could induce
proteins used in plant defense and stress responses for date palm. However, the alteration of one
defense pathway to combat a pest can have an antagonistic effect on another defense pathway, a
phenomenon described as cross-talk (Spoel et al. 2003), and subsequently, increase the
susceptibility of a plant to future attacks by herbivores or pathogens.
There are a few reports of endophytic EPF affecting interactions with aphids (Hemiptera:
Aphididae). Inoculation of cotton, Gossypium hirsutum L. (Malvales: Malvaceae), with various
fungi, including B. bassiana and Lecanicillium lecanii, significantly slowed reproduction of
cotton aphid, Aphis gossypii Glover, in greenhouse and field settings (Gurulingappa et al. 2010,
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Lopez et al. 2014). An analogous study found that B. bassiana reduced populations of bean
aphid, Aphis fabae Scopoli (Hemiptera: Aphididae) and pea aphid, Acyrthosiphon pisum Harris
(Hemiptera: Aphididae) on fava bean, Vicia faba L. (Fabales: Fabaceae), but Metarhizium
anisopliae s.l. had no effect on these aphids (Akello and Sikora 2012).
To date, few studies have considered the effect of endophytic EPF on soybean, Glycine
max L. Merr (Fabales: Fabaceae). A recent study inoculated crop plants, including soybean, with
B. bassiana and recovered the fungus from leaves of young plants, but other tissues were not
examined nor were insects included in the experiments (Russo et al. 2015). Another study that
inoculated soybean with M. anisopliae s.l. found that this fungus increased the rate of plant
development under salt stress conditions compared to controls, but aspects of endophyte
establishment or effects on herbivorous insects were not investigated (Khan et al. 2012).
Soybean growers in the North Central region did not regularly treat their fields with
insecticides until the arrival of the invasive soybean aphid, Aphis glycines Matsumura
(Hemiptera: Aphididae), in 2000 (Ragsdale et al. 2011). After the establishment of A. glycines,
the pest has quickly spread across most of the North Central region, and as a result the amount of
foliar insecticides used in soybean production has increased (Tilmon et al. 2011). Alternative
technologies, like resistant germplasm, are being considered for management of A. glycines in
order to reduce use of conventional insecticides (Ragsdale et al. 2011). Evidence suggests that
soybean defenses may be suppressed by A. glycines shortly after feeding begins (Studham and
MacIntosh 2013, Varenhorst et al. 2015), and thus, methods to enhance plant defenses may be of
value in integrated pest management (IPM) for A. glycines. In the present study, we addressed
the following questions: (1) Does seed treatment with either Beauveria bassiana or Metarhizium
brunneum establish as an endophytes in soybean? (2) Does EPF seed treatment, with or without
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endophytic establishment, affect the populations of A. glycines on soybean? (3) Which species of
Metarhizium occur in agricultural fields in Iowa?
2. Materials and Methods
2.1 Aphid experiment
2.1.1 Preparation of conidial suspensions
Conidia were produced for two strains of EPF, M. brunneum F52 and B. bassiana GHA
by solid substrate fermentation as described by Jaronski and Jackson (2012), and viability was
determined 24 h prior to seed inoculations via germination on Sabouraud dextrose agar after 18 h
of incubation at 27°C following Goettel and Inglis (1997). Conidia were suspended in
autoclaved 0.10% sorbitan mono-oleate surfactant (Tween 80) and titers were determined by a
hemocytometer using diluted conidial suspensions. Soybean seeds were inoculated with
suspensions of 1.0 × 108 conidia mL-1 of M. brunneum, B. bassiana, or a 1:1 blend of M.
brunneum and B. bassiana. Inoculations for the control contained the same autoclaved surfactant
but no fungal conidia.
2.1.2 Inoculation of soybean seeds
Seeds of the aphid-susceptible soybean cultivar (IA3027) were obtained from the Iowa
State University soybean breeding program. Seeds were briefly surface sterilized under a sterile
hood in the following order: 30 s in 0.01% autoclaved Tween 80 surfactant, 60 s in 2% sodium
hypochlorite with 5.5 pH, 30 s in 70% ethanol, and two consecutive rinses with sterile distilled water for 30 s each. After surface sterilization, seeds were air dried in the sterile hood on filter paper for 5 min. Approximately 16 seeds were designated for each treatment and placed in a
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sterile 50 mL conical tube (Cat. no. 14-959-49A, Fisher Scientific, Waltham, MA) containing 20
mL of a conidial suspension. Tubes containing the seeds and conidial suspensions were placed
in a dark incubator at 27ºC for 24 h. Tubes were laid sideways in plastic boxes in the incubator so that all seeds could be evenly distributed and soaked inside the conical tubes.
2.1.3 Planting and transfers
Inoculated seeds were individually planted in 266 mL plastic cups (Solo, Lake Forest, IL) with three small holes on the bottom for drainage. Cups were filled with unsterilized Metro mix
900 potting medium (Sun Gro Horticulture, Agawam, MA). After planting, cups with seeds received 20 mL of distilled deionized water at room temperature, and this was repeated 4 d and 7 d after planting. Eleven days after planting, seedlings were transferred to 8 cm diameter pots containing the same SB 900 potting medium, and plants were watered as needed for the duration of the experiments. From planting through the conclusion of the experiment, plants were grown in a biological incubator (27ºC, 60% RH, 14:10 L:D), with illumination provided by fluorescent lights (F25T8/TL841/ALTO, Philips, Amsterdam Netherlands) that produced 650 µmoles m-2 s-1.
2.1.4 Inoculation with A. glycines and data collection
The A. glycines used in this study were from a biotype-1 strain initiated from individuals collected from an Iowa State University research and demonstration farm in Boone County, Iowa during July 2015. The biotype identity was confirmed using detached leaf assays (Michel et al.
2010). Aphis glycines populations identified as biotype-1 are considered non-virulent and reproduce poorly on soybean cultivars with Rag genes (Rag is an abbreviation for resistance to
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A. glycines) compared to aphid-susceptible cultivars (Michel et al. 2011). However, the soybean variety used in this study did not contain any Rag genes.
Five A. glycines nymphs were placed on each plant 14 d after seeds were planted.
Nymphs were placed on the underside of the middle leaf in the first soybean trifoliate using a fine-hair paintbrush. Individual potted plants then were covered with a mesh net (Item #6LGK4,
Grainger, Lake Forest, IL) that was secured to the pots with rubber bands to prevent aphids from escaping. Mesh nets were carefully removed to count aphids on individual plants 1, 4, 7, 11 and
14 d after initial placement of aphids on plants (15, 19, 21, 25 and 28 d after planting). The experiment was replicated six times between December 2015 and October 2016, with eight plants per treatment during each replication, for a total of 192 plants for the entire experiment.
However only five plants were used for the untreated control in one replication due to accidental damage to 11 d-old plants during pot transfers, and this subsequently reduced the total sample size to 189 plants.
After counting A. glycines at the last time point (day 28), the soybean biomass above the base of the stem was removed and the potting media gently shaken off the roots. Roots were then gently rinsed under a faucet. The washed roots were placed on aluminum foil trays and placed in a drying oven for 48 h at 65ºC before recording the dry root mass.
2.2 Determination of endophytism
Inoculation of IA3027 soybean seeds, planting, and incubation were performed as previously described except that plants were not infested with aphids and were grown without the mesh coverings. Plants were arranged in a randomized complete block design with one plant per treatment randomly arranged on each of the six trays (blocks). We measured endophyte
85 establishment at three time points for the four treatments of B. bassiana, M. brunneum, the 1:1 blend of both fungi and the untreated control. At each time point (14, 21 and 28 d after planting), two trays (blocks) were randomly selected and removed from the chamber to determine endophytism. The experiment was repeated four times between May 2016 and
December 2016 for a total of 96 soybean plants (2 plants per treatment × 4 treatments × 3 time points × 4 replicates).
To determine endophytism, the stem and one unifoliate leaf from each plant was assessed for colonization by B. bassiana or M. brunneum. Specifically, one of the unifoliate leaves from inoculated plants and the uninoculated control plants was excised and surface sterilized with sterile 0.10% Tween 80 for 30 sec, 2% sodium hypochlorite with 5.5 pH for 2 min, 70% ethanol for 30 sec, and then rinsed with sterile distilled water two times. Surface-sterilized leaves were first pressed onto potato dextrose agar (PDA) in 10 cm diameter petri dish plates to determine whether any conidia were present on the surface and had the potential to germinate, which could give a false positive result. If subsequent growth of the fungus was observed on the PDA agar then any outgrowth from the corresponding plant tissues would be suspect (McKinnon et al.
2017). Leaves were then aseptically cut into ten 1 × 1 cm segments around the central vein of the leaf and placed onto selective media plates (McKinnon et al. 2017). Concurrently with cutting leaves, approximately 7 to 8 cm of the soybean stem, starting from the soil surface, was cut and surface sterilized, for each plant, in the same manner as the leaves. The surface- sterilized stems were rolled on PDA plates to determine whether any conidia were present.
Segments of 1 cm length were aseptically excised from both ends of the sterilized stems before aseptically cutting stems into five 1-cm pieces. Each 1-cm stem piece was aseptically cut in a
86 longitudinal direction and each half of a 1-cm stem piece was subsequently placed onto selective media so that the inner stem surface (pith) was facing down.
Selective media plates were sealed with parafilm and then placed in a biological incubator (27 °C, 0:24 L:D). The medium, selective for B. bassiana (GHA), was 2.0% oatmeal agar with 0.62 g L-1 dodine (Syllit 65W, Platte Chemical Inc., Greenville, MS), 0.25 g L-1 chloramphenicol (C0378, Sigma, Saint Louis, MO), and 10 mg L-1 crystal violet (C6158, Sigma,
Saint Louis, MO) and is based on that of Chase et al. (1986). To isolate M. brunneum (F52) the dodine concentration was decreased to 0.39 g L-1 and crystal violet was excluded. The growth of endophytic fungi from stems or leaves was determined after 11 d of incubation, and expressed as the proportion of plants with endophyte establishment, where proportion = number of plants with endophytes/total number of plants.
2.3 Metarhizium isolate phylogenetics
The 17 Metarhizium isolates used in this study were obtained from previous experiments that were described in Rudeen et al. (2013) and Clifton et al. (2015). Metarhizium isolates came from mycosed cadavers of greater wax moth larvae, Galleria mellonella (Lepidoptera:
Pyralidae), mealworm, Tenebrio molitor (Coleoptera: Tenebrionidae), and western corn rootworm Diabortica virgifera virgifera (Coleoptera: Chrysomelidae) that were exposed to soil collected from corn and soybean fields and their respective margins (Table S1). The data on mealworm cadavers was not previously published in the study by Clifton et al. (2015) because mealworms were only used in one of the two years of the study. Metarhizium isolates from the mycosed cadavers were recultured on oatmeal dodine agar and genomic DNA subsequently extracted.
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DNA extractions were performed with the established cetyltrimethyl ammonium bromide
(CTAB) protocol with slight modifications (Zhang et al. 2010). Before DNA extractions were
performed, 7 d old cultures of each Metarhizium isolate were prepared so that a dense mat of
mycelium had developed on the surface of a plate containing oatmeal dodine agar. Mycelia were
scraped from the surface of plates, flash frozen with liquid nitrogen, and ground in CTAB
extraction buffer. The solution was then transferred to sterile 1.5 mL microfuge tubes and
incubated overnight on a heating rack set at 55°C with 50 ng ul-1 proteinase K. Extracts were
further purified by the addition of 200 µL 1:1 phenol:chloroform, vortexed vigorously,
centrifuged at 14,000 rpm for 5 min, and then the top aqueous phase was carefully transferred to
a new 1.5 mL tube. DNA was then precipitated following addition of an equal volume of
isopropanol and a sodium chloride solution at a final concentration of 0.2 M, after which it was vortexed, and then centrifuged at 14,000 rpm for 20 min in a 4 °C refrigerated centrifuge (model
5417R, Eppendorf, Hamburg, Germany). The DNA pellets were washed with 70% ethanol and then mixed with 150 µL distilled deionized water for storage at -20°C prior to polymerase chain reactions (PCR).
Polymerase chain reactions were carried out using primers for elongation factor 1-alpha
and β–tubulin. To identify species of Metarhizium, we compared our isolates to a subset of
Metarhizium isolates described in Bischoff et al. (2009) that have sequence data for the same
genes on GenBank (Table S2). The primers used to amplify these genes were the same as those
used in Bischoff et al. (2009). Amplification of EF-1α was performed in 40 µL reaction volumes
consisting of 1X GoTaq® Flexi Buffer (Promega, Madison, WI), 1 µM of each primer, 3 mM
MgCl2, 200 µM dNTP, 2U Taq DNA polymerase, and 20 ng genomic DNA. Thermocycler
conditions consisted of a touch-down protocol with 30 s initial denaturation at 94°C followed by
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10 cycles of 10 s at 94°C, 30 s at 65–55°C (reducing annealing temperature by 1°C per cycle),
and 30 s at 72°C. Subsequently, 35 cycles were performed with the same conditions, however,
with a fixed annealing temperature of 55°C, a final extension of 10 min at 72°C, followed by a
4°C hold until PCR products were used in gel electrophoresis or stored at -20°C. Amplification
of β-tubulin was performed with the same reaction volumes and reagents, but thermocycler
conditions consisted of 40 cycles of 35 s at 94°C, 55 s at 52°C, and 2 min at 72°C, followed by a
4°C hold. Electrophoresis of 5 µL for each PCR product was performed on 1.5% agarose gel containing 0.70 mg ethidium bromide L-1, and products were visualized with a UV
transilluminator (model M-20E, UVP, Upland, CA). The remaining 35 µL of successful PCR
products were purified using the QIAquick PCR purification kit (Cat. No. 28104, Qiagen,
Germany), and submitted to the DNA Facility at Iowa State University for Sanger DNA
sequencing with an Applied Biosystems 3730 DNA analyzer (Life Technologies Corporation,
Carlsbad, CA). Sequence data were individually analyzed in SeqTrace 0.9.0 software (Stucky
2012) to confirm sequence quality and to trim ambiguous ends that lacked consensus.
In total, we analyzed elongation factor 1-alphaand β-tubulin sequence data from 45
Metarhizium isolates. Seventeen of these isolates were from samples collected in Iowa (Tables
S1 & S2). The remaining 28 isolates are described in Bischoff et al. (2009) and were downloaded from GenBank (National Center for Biotechnology Information 2017). The isolates from Bischoff et al. (2009) consisted of DNA sequence for eight species of Metarhizium: M. acridum, M. anisopliae, M. brunneum, M. guizhouense, M. lepidiotae, M. majus, M. pingshaense, and M. robertsii. Individual genes were concatenated by isolate, and sequence alignment generated with MEGA 6.0 (Tamura et al. 2013) using the Clustal W algorithm with the default settings. The best model tool in MEGA 6.0 was used to select the Jones-Taylor-
89
Thornton (JTT) model for sequence evolution, and the subsequent phylogeny was inferred using
the maximum likelihood method with 1000 bootstrap replicates (Saitou and Nei 1987) with all
aligned positions containing gaps and missing data eliminated.
2.4 Data analysis
Data on the number of aphids per plant for each data collection period were converted to
cumulative aphid days (CAD) per plant following Ruppel (1983) and Hodgson et al. (2012),
which provides a single value for the total aphid abundance on a plant over the duration of the
experiment. Data on CAD and dry root mass were analyzed with SAS Enterprise Guide 6.1
software (SAS 2017). Cumulative aphid days were transformed with the log10 function to
normalize residuals. Data were analyzed with a mixed-model of analysis of variance (ANOVA)
(PROC MIXED). The model used the fixed effect of treatment (control, B. bassiana, M.
brunneum, or B. bassiana: M. brunneum blend). The random effects were the experimental run
and the interaction of run with the fixed effect of treatment. When the effect of treatment was
significant, pairwise comparisons were made using the PDIFF statement in PROC MIXED.
Pairwise comparisons were based on least-square means with an experimentwise error rate of P
< 0.05 after using the Bonferroni adjustment for six comparisons.
For the endophyte experiment, the frequency of plants with successful recovery of endophytic EPF was analyzed with a G-test of independence using the PROC FREQ statement in
SAS 9.4 (Sokal and Rohlf 1995). The numbers of plants with or without endophytes were combined for all time points and were analyzed separately by soybean tissue (stem vs. leaf). We compared the frequency of B. bassiana endophytes to M. brunneum endophytes in the single treatments. We also compared the frequency of B. bassiana endophytes in the single treatment
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to the frequency of B. bassiana endophytes in the 1:1 blend treatment to determine if a
combination of inoculum affected endophyte recovery. The same comparison between the single
treatment and 1:1 blend treatment was performed for M. brunneum endophytes.
For the Metarhizium isolates collected from Iowa farms we identified a total of 4
haplotypes based on DNA sequence data, and a haplotype number of 1 to 4 was assigned to each
isolate. To test whether the frequency of Metarhizium haplotypes were randomly distributed
among different isolates and their location of origin, we performed a log-linear response function
(maximum likelihood analysis of variance) using the PROC CATMOD statement in SAS 9.4, with the counts of different haplotypes (1 through 4) tested with the main effects of practice
(organic vs. conventional), host (Lepidoptera vs. Coleoptera), crop (corn vs. soybean), location
(field vs. margin), and all interactions of the main effects (Ries and Smith 1963) (Table S1).
Because there were limited numbers of isolates and haplotypes distributed among the response
variables, the model was reduced until it could successfully produce an ANOVA for haplotype
frequencies. The reduced model contained the main effects of practice, host, and the interaction
of these main effects.
3. Results
For CAD there was a significant effect of treatment on A. glycines populations (Table 1,
Fig. 1). The treatment with B. bassiana GHA alone did not differ from the untreated control (df
= 15; t = 0.48; P = 1.00). However, both the treatment with M. brunneum F52 alone and the 1:1 blend had significantly greater CAD than the untreated control and the treatment with B. bassiana GHA alone (Fig. 1). However, we did not measure a significant effect of treatment on
dry root mass (Table 1, Fig. 2).
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Overall, the occurrence of B. bassiana as an endophyte was significantly greater than M. brunneum (df =1, G = 4.57, P = 0.0325). In total, with all time points combined, B. bassiana was recovered from the stems in 10 out of 48 inoculated plants, while M. brunneum was only recovered from the stems in 3 out of 48 inoculated plants. For the leaves, we recovered B. bassiana in 6 out of 48 inoculated plants, while M. brunneum was recovered from zero out of 48 inoculated plants. The frequency of B. bassiana recovery as a single inoculum did not differ significantly from its frequency as an endophyte in the 1:1 blend treatment (df = 1, G = 0.51, P =
0.48), and the same pattern was observed for M. brunneum (df = 1, G = 0.36, P = 0.55).
Our phylogenetic analysis separated Metarhizium species with strong bootstrap support
(Fig. 5). All of the Metarhizium field isolates from this study clustered within the same clade as the species Metarhizium robertsii (Fig. 5). Based on DNA sequence data, four unique haplotype sequences were observed among our 17 isolates (Fig. 5). Three transition mutations were located in the beta-tubulin gene and one transition mutation was observed in the elongation factor 1-alpha gene (Fig. S1). However, the four sites of mutation that produced these haplotypes were all synonymous substitutions that did not change the final amino acid sequences. Based on the isolate source, the frequency of different haplotypes varied by insect host and location (Table 2). Haplotypes 1 and 2 were more prevalent among Lepidoptera, whereas haplotypes 3 and 4 were more prevalent among Coleoptera (Table 2, Fig. S1).
Haplotypes 3 and 4 were more prevalent in fields, and haplotype 2 was more prevalent in margins outside of cropping fields.
4. Discussion
One goal of this study was to determine the effects of EPF on A. glycines when used as a seed inoculum and to determine whether EPF can establish endophytically in soybean. We
92
found that plants inoculated with M. brunneum alone or in the blend treatment (1:1 mixture of M.
brunneum and B. bassiana) had overall greater abundance of A. glycines compared to the plants
inoculated with B. bassiana or the untreated controls (Table 1, Fig. 1). We also found a
significantly lower prevalence of plants with endophytic M. brunneum compared to B. bassiana
(Figs. 3 and 4). Metarhizium brunneum was only isolated from stems whereas B. bassiana was isolated from stems and leaves, and we were able to recover both fungi from the stems of individual plants treated with the 1:1 blend (Fig. 3). Another goal of this study was to identify the species of Metarhizium found in cropping systems. We found that all of our Metarhizium isolates were identified as M. robertsii (Fig. 5) and that there was an association of different haplotypes with the insect host (Table 3).
We hypothesize two potential explanations for the increased aphid abundance on plants inoculated with M. brunneum: 1) the M. brunneum fungus reduced plant defenses, or 2) M. brunneum improved host plant quality. Soybean plants may have responded to M. brunneum as if it were a pathogen with defensive pathways mediated by salicylic acid (SA), and the subsequent cross-talk between the SA pathway and thejasmonic acid (JA) pathway could have made plants more vulnerable to aphids (Kunkel and Brooks 2002, Spoel et al. 2003). The observed lower levels of endophytism by M. brunneum compared to B. bassiana suggests that soybean plants did not support the presence of Metarhizium endophytes as well as Beauveria,
and that plants may have removed the fungus with a mechanism that is also employed to remove
saprophytic pathogens, many of which are regulated by the SA pathway (Conrath 2006, Pieterse et al. 2014). Regarding host plant quality, Metarhizium spp. can increase root hair development and translocate nitrogen from insect cadavers in the soil to nearby plants (Behie. et al 2012,
Sasan and Bidochka 2012), and perhaps M. brunneum did the same for soybean plants in this
93
experiment, although we did not observe any differences in root biomass among treatments (Fig.
2).
Plants are not passive bystanders to attack and can recognize microbial compounds (i.e.,
fungal chitin), and thus trigger defense pathways when challenged by a pathogen (Zipfel 2009,
Newman et al. 2013). Beneficial fungi and saprophytic pathogens have adapted mechanisms
(i.e., the secretion of effector proteins) to bypass plant defenses and colonize extracellular spaces
of plant tissues (van Esse et al. 2007). In this study, it appears that M. brunneum may elicit a
plant response that facilitates a greater abundance of A. glycines. Past research has suggested a trend that EPF, particularly B. bassiana, may induce plant defenses that would in turn suppress
insects (Lopez et al. 2014, McKinnon et al. 2017), but we did not observe this result for A.
glycines. Akello and Sikora (2012) used a similar method to ours of soaking fava bean seeds in
conidial suspensions and found that M. anisopliae s.l. had no effect on A. fabae and A. pisum
aphids, whereas B. bassiana significantly reduced aphid fecundity. The high concentration of M.
brunneum conidia in the seed-soak inoculum may have upregulated SA-mediated defenses that
are often triggered in response to saprophytic pathogens, e.g. Fusarium spp. or Pythium spp.
(Shoresh et al. 2010). The ensuing cross-talk between the SA and JA pathways may have come
at a cost that made plants more susceptible to insect herbivory. However, we did not measure
levels of endogenous SA or JA in soybean tissues that might explain the hypothesized alterations to plant defenses (Durrant and Dong 2004).
We did not observe any difference in root biomass among treatments (Fig. 2) suggesting that effects on nutrient acquisition were not present. Previous studies have observed that B.
bassiana and M. anisopliae s.l. can colonize leguminous plant roots and boost plant development
(Akello and Sikora 2012, Behie et al. 2015, Parsa et al. 2016). In a study of EPF inoculations on
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soybean, Khan et al. (2012) showed that M. anisopliae s.l. improved soybean shoot length and
shoot dry weight when plants were grown under salt-stressed conditions. Endophytic EPF may
increase plant growth in a way that allows plants tolerate more insect herbivory and compensate
for lost biomass (Waller et al. 2005, McKinnon et al. 2017). In the case of phloem-feeders like
aphids, M. brunneum may have increased levels of acquirable nutrients in the plant. Behie et al.
(2012) showed that M. robertsii could translocate nitrogen from soilborne insect cadavers to the
leaves of nearby haricot bean seedlings. Aphids have been observed to be more fecund and increase their populations more rapidly on plants supplemented with nitrogen fertilizer (Aqueel and Leather 2011). However, it is unknown to what extent M. brunneum may have altered host-
plant quality in this experiment.
Based on our results with endophyte establishment in the 1:1 blend of M. brunneum and
B. bassiana, there does not seem to be a negative effect of combining inoculum on either strain
(Figs. 2 & 3). In general, B. bassiana seems capable of establishing as an endophyte
aboveground in the phylloplanes of many plants, whereas Metarhizium spp. tend to establish as
endophytes in roots (Hu and St. Leger 2002, Meyling and Eilenberg 2007, Wyrebek and
Bidochka 2013). Past research has shown that Beauveria spp. and Metarhizium spp. are capable
of establishing as endophytes in some of the same crops, including tomato and haricot bean
(Powell et al. 2009, Elena et al. 2011, Behie et al. 2015). A few studies with insect assays have
found that combinations of Beauveria spp. and Metarhizium spp. can increase pest mortality in
an additive or synergistic manner (Inglis et al. 1997, Geetha et al. 2012, Kryukov et al. 2015),
but the relevant literature is lacking on whether combinations of EPF used as crop inoculum can
improve plant development or host-plant resistance relative to single-strain inocula.
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For Metarhizium isolates collected from different insects and fields throughout Iowa
(Rudeen et al. 2013, Clifton et al. 2015), which included factors of production (organic vs.
conventional), location (field vs. margin), and crop (corn vs. soybean), our phylogenetic analysis
revealed that all isolates were M. robertsii (Fig. 5). Rehner and Kepler (2017) recently described
the phylogeography of the M. anisopliae complex and found that M. robertsii is predominant in
North America compared to other regions, and this could reflect historical differences in species origin or ecological adaptations. Previous studies have shown that different species of
Metarhizium may not have random distributions but rather exhibit some specificity for the rhizosphere of certain plants (Wyrebek et al. 2011). Behie et al. (2015) obtained hundreds of
Metarhizium isolates from of the roots of grasses, forbs and sedges in Canada and identified 95% of the Metarhizium isolates as M. robertsii. We did not sample rhizosphere soil from corn or soybean roots in the Clifton et al. (2015) study but still obtained M. robertsii isolates in the bulk soil adjacent to crop rows. Despite the apparent prevalence of M. robertsii in North America, other Metarhizium spp. occur in the region and it is possible that our soil sampling methods missed these less common species (Rehner and Kepler 2017).
Metarhizium robertsii is known to infect a variety of insect hosts (Rehner and Kepler
2017) and there was a near 50:50 mix of Coleopteran hosts or Lepidopteran hosts among our
Iowa isolates. We tested the frequency of haplotypes and found a significant effect with insect hosts and location (Table 2). Other studies on EPF isolates belonging to the same species used molecular tools and observed distinct groups based on host insect range (Maurer et al. 1997,
Farques et al. 2002, Rezende et al. 2015). Otherwise, our analyses on M. robertsii haplotypes requires more isolates to know whether the method of crop production (organic vs. conventional) and the crop grown in a particular field (corn vs. soybean) would affect the prevalence of
96
particular M. robertsii haplotypes. Hernández-Domínguez and Guzmán-Franco (2017) collected
Metarhizium isolates from one sugarcane field and found that the frequency of specific haplotypes had significantly varied by sampling dates.
In terms of management of A. glycines, it remains unclear whether microorganisms like
EPF could help to protect soybean cultivars from aphids. Lopez et al. (2014) were able to suppress A. gossypii populations in the lab and field with B. bassiana in cotton, suggesting that some fungal endophytes could be useful tools in management of aphids, but the same study did not include a strain of Metarhizium. The recent review by McKinnon et al. (2017), suggests that
B. bassiana endophytes tend to have positive impacts on plant development and negative impacts on insect pests, but a similar review on Metarhizium endophytes is lacking. Regardless of trends, the tritrophic interactions among host plants, insects and fungal entomopathogens can be very specific and complex. Metarhizium brunneum F52 was used in these experiments but we did not identify any of our field-collected isolates as M. brunneum. Other species of Metarhizium also might establish as soybean endophytes and their potential impacts on plant development and A.
glycines could be different. Because M. robertsii can be found in soybean fields and because of
its wide distribution in North America (Behie et al. 2015, Rehner and Kepler 2017), future
studies should consider the ecological roles of M. robertsii in row crop agriculture and IPM.
Future studies also should determine the potential impacts of fungal endophytes on crops with
host-plant resistance genes, which are already considered as a key tool in IPM (Pedigo and Rice
2009), especially for pests like A. glycines (Ragsdale et al. 2011).
97
Acknowledgments
Bob Gunnarson, Sarah Rueger, Greg VanNostrand, Erika Rodbell and Shelby Pritchard
assisted with the setup of experiments in the lab. Gregory Tylka, Adam Varenhorst and Mike
McCarville provided advice on experimental designs and helpful comments on early versions of the manuscript. This work was partially supported by the College of Agriculture and Life
Sciences at Iowa State University.
98
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Tables
Table 1: Mixed-model analysis of variance for cumulative aphid days (CAD) and root mass in the experiment with soybean aphid (Aphis glycines).
Dataset Fixed effect d.f. F-value P-value CAD Treatment 3, 15 8.68 0.0014 Dry root mass Treatment 3, 15 1.11 0.3819 Random effect d.f. χ2 P-value CAD Experiment run 1 342.9 <0.0001 Experiment run × treatment 1 2.7 0.0502 Dry root mass Experiment run 1 67.1 <0.0001 Experiment run × treatment 1 1.8 0.0899
Table 2: Log-linear model predicting the frequency of Metarhizium haplotypes.
Source d.f. χ2 P Intercept 1 92.80 <0.0001 Hosta 1 9.82 0.0017 Locationb 1 5.39 0.0203 Host × Location 1 0.48 0.4862
a Host: Coleoptera vs. Lepidoptera b Location: field vs. margin
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Figures
Figure 1: Cumulative aphid days for soybean aphids (Aphis glycines) on soybean plants. Bar heights are sample means and error bars are the standard error of the mean. Letters above the bars represent significant differences between means.
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Figure 2: Dry root mass of soybean plants. Bar heights are sample means and error bars are the standard error of the mean.
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Figure 3: Mean proportion of plants with endophyte recovery in (A) soybean stems and (B) soybean leaves. Error bars represent standard error of the means. Bars are separated by the time points of 14, 21 or 28 days after planting. The shading in bars represents treatments of Beauveria bassiana GHA alone or Metarhizium brunneum F52 alone.
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Figure 4: Mean proportion of plants with endophyte recovery in (A) soybean stems and (B) soybean leaves in the 1: 1 blend treatment of Beauveria bassiana GHA and Metarhizium brunneum F52. Error bars represent standard error of the means. Bars are separated by the time points of 14, 21 or 28 days after planting. The shading in bars represents the fungus that was recovered. Image is included showing endophytic B. bassiana and M. brunneum being recovered from separate stem pieces in the same soybean plant.
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Figure 5: Maximum Likelihood-based phylogenetic relationships among our Metarhizium isolates and related species based on analysis of aligned concatenated β-tubulin and elongation factor 1-alphasequence data. Bootstrap values (1,000 replicates) are indicated besides the nodes. The tree is rooted to Metarhizium acridum. The tree is drawn to scale, with branch lengths measured in the number of substitutions per site. Brackets are placed next to our unique haplotypes (#1-4). The GenBank accession numbers for β-tubulin and elongation factor 1-alpha sequences are provided in Table S1.
Supplemental Tables
Supplementary Table 1: Metarhizium isolates used in the phylogeny study, their location in Iowa, their host before media culture, and the previous study from which they came. Elongation factor 1-alpha (EF-1α) and β–tubulin (Bt) genes were sequenced for all isolates.
Cropping system Isolate ID and location Nearby city Isolation source Order Study Bta EF-1αb
11wax01 Conva corn field Iowa Falls G. mellonella Lepidoptera Clifton et al. (2015) MF326391 MF326374 11wax05 Organic corn margin Kalona G. mellonella Lepidoptera Clifton et al. (2015) MF326392 MF326375 11wax10 Organic soybean field Kalona G. mellonella Lepidoptera Clifton et al. (2015) MF326393 MF326376 11wax11 Organic soybean margin Kalona G. mellonella Lepidoptera Clifton et al. (2015) MF326394 MF326377 11wax13 Organic soybean field Hampton G. mellonella Lepidoptera Clifton et al. (2015) MF326395 MF326378 12meal02 Conv soybean field Iowa Falls T. molitor Coleoptera Clifton et al. (2015) MF326396 MF326379
12meal04 Conv soybean margin Iowa Falls T. molitor Coleoptera Clifton et al. (2015) MF326397 MF326380 111 12meal06 Conv corn field Kalona T. molitor Coleoptera Clifton et al. (2015) MF326398 MF326381 12meal21 Conv soybean field Hampton T. molitor Coleoptera Clifton et al. (2015) MF326399 MF326382 12meal23 Conv soybean field Hampton T. molitor Coleoptera Clifton et al. (2015) MF326400 MF326383 12meal25 Conv soybean field Hampton T. molitor Coleoptera Clifton et al. (2015) MF326401 MF326384 12meal39 Conv soybean field Sutherland T. molitor Coleoptera Clifton et al. (2015) MF326402 MF326385 12meal41 Conv soybean margin Sutherland T. molitor Coleoptera Clifton et al. (2015) MF326403 MF326386 12wax25 Conv soybean field Hampton G. mellonella Lepidoptera Clifton et al. (2015) MF326404 MF326387 12wax40 Organic soybean margin Sioux Center G. mellonella Lepidoptera Clifton et al. (2015) MF326405 MF326388 12wax52 Conv soybean margin Sutherland G. mellonella Lepidoptera Clifton et al. (2015) MF326406 MF326389 Met3A Conv corn field Manchester D. v. virgifera Coleoptera Rudeen et al. (2013) MF326407 MF326390 a Bt: GenBank accession number for β-tubulin sequence data b EF-1α: GenBank accession number for elongation factor 1-alpha sequence data c Conv: conventional farming production
Supplementary Table 2: Reference sequences used for phylogenetic analyses; includes strain codes, taxon name, host, country of collection, and GenBank accession numbers for the subset of isolates picked from Bischoff et al. (2009).
GenBank accession numbers Voucher # Taxon Isolation source Country EF-1α Beta-tub 727 Metarhizium robertsii Orthoptera Brazil DQ463994 EU248816 4739 Metarhizium robertsii Soil Australia EU248848 EU248928 7501 Metarhizium robertsii Coleoptera Australia EU248849 EU248929 4342 Metarhizium pingshaense Coleoptera Solomon Islands EU248851 EU248821 CBS 257.90 Metarhizium pingshaense Coleoptera China EU248850 EU248820 3210 Metarhizium pingshaense Coleoptera India EQ463995 EU248819 7929 Metarhizium pingshaense Isoptera Australia EU248847 EU248815 7450 Metarhizium anisopliae Coleoptera Australia EU248852 EU248823 112 7487 Metarhizium anisopliae Orthoptera Eritrea DQ463996 EU248822 2107 Metarhizium brunneum Coleoptera USA EU248855 EU248826 4179 Metarhizium brunneum Soil Australia EU248854 EU248825 4152 Metarhizium brunneum Soil Australia EU248853 EU248824 7505 Metarhizium majus Coleoptera Australia EU248870 EU248842 4566 Metarhizium majus Coleoptera Australia EU248869 EU248841 2808 Metarhizium majus Coleoptera Philippines EU248871 EU248843 1914 Metarhizium majus Coleoptera Philippines EU248868 EU248840 1946 Metarhizium majus Coleoptera Philippines EU248867 EU248839 1015 Metarhizium majus Lepidoptera Japan EU248866 EU248838 7502 Metarhizium guizhouense - Australia EU248861 EU248833
Supplementary Table 2 continued
GenBank accession numbers Voucher # Taxon Isolation source Country EF-1α Beta-tub 4321 Metarhizium guizhouense Soil Australia EU248860 EU248832 7507 Metarhizium guizhouense Soil Kiribati EU248858 EU248831 CBS 258.90 Metarhizium guizhouense Lepidoptera China EU248862 EU248834 6238 Metarhizium guizhouense Lepidoptera China EU248857 EU248830 5714 Metarhizium guizhouense - - EU248856 EU248829 7488 Metarhizium lepidiotae Coleoptera Australia EU248865 EU248837 7412 Metarhizium lepidiotae Coleoptera Australia EU248864 EU248836 4628 Metarhizium lepidiotae Soil Australia EU248863 EU248835
7486 Metarhizium acridum Orthoptera Niger EU248845 EU248813 113
Supplemental Figures
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Supplemental Figure 1: Sites of transition mutations in the genes amplified for our Metarhizium isolates that made up the four haplotypes. Corresponding mutation sites are included for the reference isolates of M. robertsii from Bischoff et al. (2009).
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CHAPTER 4. GENERAL CONCLUSIONS
The objectives of this dissertation were to evaluate pest management tools on two soybean pests. In two separate experiments, our findings suggest that host-plant resistance is a consistent tool for managing soybean aphid and soybean cyst nematode. The goal of Chapter 2 was to measure the effects of resistant cultivars and pesticidal seed treatments on pest populations and soybean yield. Soybean aphids responded negatively to resistant cultivars and the insecticide thiamethoxam only reduced aphids on one of the susceptible cultivars. Soybean cyst nematodes had reduced reproduction on resistant cultivars, and also identified all of the nematode populations in our fields as a virulent biotype. The nematicide abamectin did not affect soybean cyst nematode. Pesticidal seed treatment inconsistently improved yields. The prophylactic use of these chemicals could add unnecessary costs to soybean production.
The goals of Chapter 3 were to determine whether fungal entomopathogens could establish as endophytes in soybean and their potential effects on soybean aphids if plant immunity were altered. Previous research suggests that fungal entomopathogens could induce plant defenses and reduce pest injury, and thus we predicted that soybean aphids would reproduce more slowly on inoculated plants. Beauveria bassiana had no effect on aphids but still established as an endophyte; Metarhizium brunneum actually increased aphid numbers.
Metarhizium brunneum had low prevalence as an endophyte, suggesting that plants may have responded with defense mechanisms used for plant pathogens and subsequent cross-talk of defense pathways made plants more susceptible to aphid infestation. It is unclear whether these fungi could be exploited for preventive pest management, but our identification of Metarhizium robertsii in soils throughout Iowa suggests that this species might already serve ecological roles in cropping systems that should be studied further
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APPENDIX A. EFFECTS OF HOST-PLANT RESISTANCE AND
PESTICIDAL SEED TREATMENT ON SOYBEAN APHID AND
SOYBEAN CYST NEMATODE ON SOYBEAN
Preface
This experiment was intended to be a separate chapter or serve as a complementary experiment to Chapter 2, with similar objectives that addressed the impacts of host-plant
resistance and seed-applied pesticides on pest populations. The work was ultimately removed
from the manuscript and the body of the dissertation because of errors that affected data
collection. The protocol of the experiment closely followed the water bath experiments
described in the greenhouse experiment in Chapter 2. The experiment used soil containing H.
glycines eggs, except that plants were artificially infested with A. glycines for part of the
experiment. We later discovered that the A. glycines colony used for the experiment may have
been contaminated with a different biotype or incidentally transferred from the incorrect colony.
Follow-up assays confirmed that the A. glycines colony was virulent on Rag1 cultivars, and this explained the observation of high numbers of aphids on our Rag1 plants. In one run of the experiment, we measured low numbers of H. glycines females on most of the plants. During a rainstorm that followed this experimental run, we noticed that outside rainfall was leaking through the greenhouse roof vents and dripping over the greenhouse bench. This unintentional watering of the greenhouse benches might have flooded the buckets and stalled the reproduction
of H. glycines on the roots, which do not survive well in saturated soils. These outcomes led to
the termination of the experiment after a few runs, but we still observed how seed-applied
thiamethoxam can suppress A. glycines on young plants.
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1. Introduction
Soybean cyst nematode, Heterodera glycines Ichinohe (Tylenchida: Heteroderidae), and soybean aphid, Aphis glycines Matsumura (Hemiptera: Aphididae), are invasive pests that share soybean as a host plant (Niblack et al. 2006, Tilmon et al. 2011). Heterodera glycines has been present in North America for more than 60 years and continues to be one of the leading suppressors of soybean yield (Wrather et al. 2010). Aphis glycines has been present in North
America since 2000 and can reduce soybean yield by as much as 40% (Ragsdale et al. 2007).
Since the discovery of H. glycines in North America in 1954, management of this pest has relied on soybean germplasm with genetic resistance (Winstead 1955, Niblack 2005). Some of the main sources of H. glycines-resistant germplasm include Peking, PI88788, and PI90763, with PI88788 being the most-widely used source of resistance (Shannon et al. 2004, McCarville et al. 2017). Host-plant resistance also can be used to manage A. glycines (McCarville et al.
2013). The first soybean cultivars with A. glycines resistance used a single resistance gene,
Rag1, and were available in 2010 (Mardorf et al. 2010). Aphid-resistant cultivars are not completely devoid of aphid populations, but those aphids typically have lower reproduction on resistant plants (McCarville et al. 2014b). Virulent biotypes of H. glycines, which are often designated by different HG types according to the H. glycines HG type test (Niblack et al. 2002), have been observed in Iowa, Illinois, Minnesota, Missouri and Ontario (Cary and Diers 2016).
These virulent biotypes can diminish the benefits of resistant cultivars and those that are virulent to PI88788 are more widespread in North America (McCarville et al. 2017). Virulent biotypes of A. glycines also occur and they can reproduce on Rag cultivars just as well as they can on susceptible cultivars. Aphis glycines populations identified as biotype-1 are considered non- virulent and reproduce poorly on soybean cultivars with Rag genes compared to aphid- susceptible cultivars (Michel et al. 2011).
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Insecticides and nematicides applied as seed treatments are another option to manage A. glycines and H. glycines. The most widely-used class of insecticidal seed treatments on the market are the neonicotinoids, including clothianidin, imidacloprid, and thiamethoxam (Douglas and Tooker 2015). Some labels for neonicotinoid seed treatments include A. glycines as a target pest. Neonicotinoid seed treatments can slow the reproduction and kill A. glycines populations that colonize soybean plants within the first 55 days after planting (McCarville and O’Neal
2013). The nematicide abamectin is used in some seed treatment products, but it is not systemic and interferes with the nervous system of nematodes that come into contact with the chemical.
The hypotheses of this greenhouse experiment were:
1) Host-plant resistance will reduce numbers of A. glycines and H. glycines
2) Abamectin and thiamethoxam seed treatment will reduce numbers of A. glycines and H. glycines
2. Materials and Methods
2.1 Greenhouse experiment
We performed 28-day greenhouse experiments modified from the Standard Cyst
Evalution-2008 (SCE-08) protocol to measure treatment effects on a single generation of H. glycines reproduction (Niblack et al. 2009). We also measured treatment effects on A. glycines populations that were put on plants halfway through the experiment. The experimental design followed McCarville et al. (2014a). Heterodera glycines typically completes one generation every 25 to 32 days at 27 to 30ºC, thus the 30-day experiment estimates the amount of H. glycines reproduction in one generation (Alston and Schmitt 1988).
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We used 12 treatments from a fully-crossed factorial design with four soybean cultivars
and three seed treatments. We used four soybean varieties with relative maturities ranging from
2.1 to 2.5, as indicated by the first two numbers in the following variety names: 1) the cultivar
susceptible to both pests was S24-K2, 2) the A. glycines-resistant (Rag1) cultivar was S25-F2, 3)
the H. glycines-resistant (PI88788) cultivar was S23-P8, and 4) the cultivar with both Rag1 and
PI88788 genes was S21-Q3 (Syngenta AG, Greensboro, North Carolina, U.S.A.). The seed
treatments used in this study were 1) ApronMaxx®, which contained the fungicides mefenoxam
(0.0113 mg AI seed-1) and fludioxonil (0.0038 mg AI seed-1), 2) Avicta Complete®, which
contained the same fungicides in ApronMaxx® at the same rates plus the nematicide abamectin
(0.15 mg AI seed-1) and the neonicotinoid insecticide thiamethoxam (0.0907 mg AI seed-1), and
3) seeds that were left untreated (Syngenta AG, Greensboro, North Carolina, U.S.A.).
The soil used for the containers contained a population of H. glycines with the HG type 0
(zero) that is considered avirulent to H. glycines-resistant cultivars. We used the same soil source from Muscatine, Iowa containing a H. glycines population that is described in McCarville et al. (2014a). At the time of preparing soil for the containers, the field soil was diluted with the appropriate amount of construction sand to adjust the H. glycines egg densities to ca. 10 eggs 1 cc soil-1. Single soybean plants were grown in 125-mL cone-shaped containers (Stuewe & Sons,
Tangent, OR), containing 100 mL field soil.
One container for each treatment was placed in a random arrangement inside a 7.5 L
sealed plastic bucket that was filled with construction sand to match the level of the soil inside
each container (Fig. 1). In each experiment, eight replications (i.e., buckets) were established,
with each bucket containing one container per treatment, for a total of 12 containers per bucket.
We performed two experimental runs in April 2015 and June 2015 and total of 192 plants were
120 used in the experiments (2 runs × 8 replications × 12 treatments = 192 pants). Buckets were kept in a temperature-controlled water bath (2.44 m × 1.22 m × 0.30 m; L × W × D) that stabilizes the soil temperature between 27 and 30ºC. The greenhouse used for these experiments contained three water baths. A water bath containing buckets and plants was housed on a greenhouse bench such that the tops of buckets were situated ca. 60 cm below 400 W high-pressure sodium growth lamps (16:8 (L:D)). Plants were watered as needed. At the end of the experiment, containers were removed from the buckets and roots were washed with a pressurized faucet sprayer to dislodge H. glycines females (cysts) on a 600-µm–pore sieve placed above a 250-µm- pore sieve. The cysts were then washed from the 250-µm-pore sieve into individual 100 mL beakers and later counted using direct microscope observation (Leica S6 E, Leica Microsystems,
Wetzlar, Germany).
The same plants grown in the water bath experiments were artificially infested with A. glycines nymphs. The A. glycines nymphs came from a colony maintained at Iowa State
University that was considered to be biotype-1, and the colony was initiated from a population in
Illinois (Kim et al. 2008). Ten A. glycines nymphs were placed on each plant 11 d after seeds were planted. Nymphs were placed on the underside of the middle leaf in the first soybean trifoliate using a fine-hair paintbrush. After infestation, buckets containing the plants were covered with mesh nets to prevent aphid escape (Fig. 1). However, individual plants within a bucket were not covered with their own nets, so it is possible for aphids to move from plant to plant inside of a bucket and choose other plants to colonize. At 7, 11, and 14 d after infestation
(or 18, 22, 25 d in the experiment), mesh nets were carefully removed to count the numbers of A. glycines on individual plants.
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2.2 Data Analysis
Data on the number of aphids per plant per data collection period were converted to
cumulative aphid days (CAD) per plant following Ruppel (1983). Unless otherwise stated, we
analyzed data with a mixed-model analysis of variance (ANOVA) using the PROC MIXED
statement in SAS 9.4. Random effects were tested using a log-likelihood ratio statistic (-2 RES
log likelihood) based on a one-tailed χ2 test assuming one degree of freedom (Littell et al. 1996).
Random factors were removed from the model to increase the statistical power when these
factors were not significant at a level of α < 0.25 (Quinn and Keough 2002).
The number of H. glycines females (cysts) and CAD per soybean plant were analyzed
with a mixed-model ANOVA that included the fixed factors of cultivar, seed treatment, and the
interaction of cultivar × seed treatment. The model included the random factor of experimental
run. When a significant interaction was present, pairwise comparisons were made using the
PDIFF statement in PROC MIXED. Pairwise comparisons were based on least-square means
with a significance level of P < 0.05 after using the Bonferroni adjustment for multiple comparisons.
3. Results and Discussion
We observed a significant effect of cultivar on the numbers of H. glycines females per plant, but seed treatment was not significant (Table 1). The H. glycines-resistant cultivar had significantly fewer H. glycines females than the A. glycines-resistant cultivar (t178 = 3.08, P =
0.0144), and the same was true for the A. glycines- and H. glycines-resistant cultivar (t178 = 3.30,
P = 0.0070) (Fig. 2). The cultivar susceptible to both pests had significantly more H. glycines females compared to the A. glycines- and H. glycines-resistant cultivar (t178 = 2.79, P = 0.0347), but the same comparison to the H. glycines-resistant cultivar was not significant (Fig. 2).
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For CAD there was a significant effect of cultivar, seed treatment, and the interaction of
cultivar with seed treatment (Table 1, Fig. 3). The factor of cultivar also was significant, as we
observed the cultivar with resistance to both pests having the lowest CAD, and it was
significantly lower than the CAD on the cultivar with susceptibility to both pests (t178 = 3.31, P =
0.0069). With all cultivars combined, the seed treatment containing the insecticide
thiamethoxam had significantly lower CAD than untreated seeds (t178 = 8.93, P < 0.0001) and
the fungicides seed treatment (t178 = 7.05, P < 0.0001). For the soybean cultivar that was susceptible to both pests and for the A. glycines-resistant cultivar, plants grown from seeds
treated with the insecticide thiamethoxam had significantly lower CAD than untreated soybeans
(t178 = 6.98, P < 0.0001; and t178 = 5.04, P < 0.0001, respectively) and those treated with fungicide (t178 = 3.73, P = 0.0171; and t178 = 4.51, P = 0.0008, respectively) (Fig. 3). However,
for the H. glycines-resistant cultivar and the cultivar resistant to both pests there were no
significant differences in CAD among plants with our without seed treatment (Fig. 3).
Our results with H. glycines confirm our hypothesis that resistant cultivars support lower
reproduction of an avirulent nematode population. However, the differences in the numbers of
H. glycines females between the resistant and susceptible cultivars were not as great as the
differences observed in other greenhouse experiments on H. glycines (see Table S3 in Chapter
2). One potential explanation for this outcome is that the H. glycines-resistant cultivars (S23-P8
and S21-Q3) are only moderately resistant and do not suppress nematode reproduction as well as
other lines. Some other greenhouse trials with these cultivars have shown high numbers of H.
glycines females on the S23-P8 cultivar (unpublished data) (Tylka 2013). We also observed that
abamectin had no effect on H. glycines. Abamectin can suppress root-knot nematodes
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(Meloidogyne spp.) in cotton and vegetable crops, but so far there is little evidence that abamectin works well for cyst-forming nematodes like H. glycines (Monfort et al. 2006).
Although the cultivar with resistance to both pests had lower CAD than the cultivar that was susceptible to both pests, the difference in CAD was not as great as the difference we observed in field plot experiments that counted A. glycines populations. Having used the putative biotype-1 colony for infestation, we expected the resistant cultivars with the Rag1 gene to have much lower numbers of aphids. We observed significantly low numbers of A. glycines on Rag1 cultivars in the field and other greenhouse experiments that used the biotype-1 colony before these experiments were performed. After the completion of the greenhouse experiment,
we performed some whole-plant assays on the A. glycines colony and found that their
reproduction Rag1 was similar to its reproduction on susceptible cultivars, and thus the colony could be considered a biotype-2 population at the time of the experiments (unpublished data).
We do not know how the colony was altered, but one hypothesis is that some virulent biotype-2 aphids were inadvertently introduced into the colony before we performed the experiments.
Another potential explanation is that Rag1 plants were mistakenly used for rearing the colony at some time before our experiments, and subsequently pressured the A. glycines colony to
reproduce on Rag1 soybean.
The low numbers of aphids on the plants containing the thiamethoxam seed treatment was anticipated because the soybean plants were <55 d old, and thus thiamethoxam still had systemic activity in the plants (McCarville and O’Neal 2013). Because the plants were not separated within the buckets covered with mesh nets, we do not know if the aphids fed on the plants with thiamethoxam and then died, or if the aphids attempted feeding on those plants
before moving to untreated plants.
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In summary, we observed that the insecticide thiamethoxam can effectively reduce A. glycines populations on young soybean plants, but the nematicide abamectin in the same seed
treatment had no effect on H. glycines. Insecticidal seed treatments may help with early-season
protection of soybean, but otherwise they have shown inconsistent results with season-long
suppression of A. glycines and preservation of yield (McCornack and Ragsdale 2006, Magalhaes
et al. 2009, Seagraves and Lundgren 2012, Krupke et al. 2017). The H. glycines-resistant
cultivars had fewer females than susceptible cultivars, but our cultivars may not reflect the
efficacy of other resistant cultivars for H. glycines management. Previous studies on A. glycines-
resistant cultivars have demonstrated their ability to suppress aphid populations (O’Neal and
Johnson 2010, Mardorf et al. 2010), but in this experiment we happened to use a virulent
biotype. Furthermore, virulent biotypes of A. glycines do occur in the field and there are now
soybean cultivars with the Rag1/Rag2 pyramid of resistance genes that can suppress A. glycines more than Rag1 or Rag2 alone (McCarville et al. 2014b).
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Magalhaes, L. C., Hunt, T. E., Siegfried, B. L. 2009. Efficacy of neonicotinoid seed treatments to reduce soybean aphid populations under field and controlled conditions in Nebraska. J. Econ. Entomol. 102 (1), 187-195.
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Tables
Table 1: Mixed model of analysis for Heterodera glycines females per plant and CAD in the greenhouse experiment.
Data set Fixed effect d.f. F-value P-value
H. glycines Cultivar 3, 178 5.85 0.0008
Seed treatment 2, 178 1.27 0.2833
Cultivar × seed treatment 6, 178 0.21 0.9722
CAD Cultivar 3, 178 4.55 0.0042
Seed treatment 2, 178 44.20 <0.0001
Cultivar × seed treatment 6, 178 2.38 0.0307
Data set Random effect d.f. χ2 P-value
H. glycines Experimental run 1 29.6 <0.0001
CAD Experimental run 1 24.3 <0.0001
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Figures
Figure 1: Photos of the water bath experiments after planting seeds and the 11 d old plants after A. glycines infestation.
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Figure 2: Average number of H. glycines females per plant after 28 days in the greenhouse experiment. Bars represent combinations of cultivar × seed treatment. F = fungicides; I = insecticide; N = nematicide. Error bars represent standard error of the means. Bar groups are separated by soybean cultivar. SCN = soybean cyst nematode. SBA = soybean aphid. Shading or patterns in the bars represent seed treatment. Letters above the bars represent significant means separation (P < 0.05) by soybean cultivar, using the Bonferroni adjustment for multiple comparisons in ANOVA.
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Figure 3: Average cumulative aphid days after 14 days in the greenhouse experiment. Bars represent combinations of cultivar × seed treatment. Error bars represent standard error of the means. Bar groups are separated by soybean cultivar. SCN = soybean cyst nematode. SBA = soybean aphid. Shading or patterns in the bars represent seed treatment. F = fungicides; I = insecticide; N = nematicide. Capital letters above the bars represent significant means separation (P < 0.05) by soybean cultivar. Lowercase letters above the bars denotes a significant difference between means for seed treatments within a soybean cultivar.
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APPENDIX B. MISCELLANEOUS PROJECTS AND CO-AUTHORED
PUBLICATIONS
Listed below are publications from my time as a PhD student. In all cases, I helped conduct experiments and/or prepare drafts of the manuscripts.
Gassmann, A.J. and Clifton, E.H. 2016. Current and potential applications of biopesticides to manage insect pests of maize. In Lacey, L.A. (Ed.), Microbial Control of Insect and Mite Pests: From Theory to Practice. 1st edition. Elsevier, London.
Abstract Maize is attacked by a range of lepidopteran and coleopteran pests, and these pests feed on plants at every phenological stage. Historically, management of maize pests has focused on conventional insecticides. However, beginning in the 1990s genetically engineered maize that produced insecticidal toxins derived from the bacterium Bacillus thuringiensis was made commercially available for management of pest insects. The first Bt maize produced single Bt toxins that targeted lepidopteran pests but later additional Bt toxins targeted coleopteran pests, specifically corn rootworms Diabrotica spp., and multiple Bt toxins were placed into the same maize hybrids. Planting of Bt maize has provided an effective management option for many key pests of maize but recent cases of pest resistance illustrate the potential limitations of this technology. More diversified management will increase the sustainability of pest management in maize. Past research has demonstrated the potential efficacy of a range of microbial control agents, including bacteria, fungi, nematodes and viruses. Of particular interest are entomopathogenic fungi for management of aboveground pests and entomopathogenic fungi and nematodes for management of belowground pests. One challenge to the implementation of microbial control agents is the need for pest management options that are cost effective and may be applied easily and rapidly over a large area. Potential future applications of microbial control agents in maize may include bolstering populations of entomopathogens in the soil through conservation biological control, developing entomopathogenic fungi for application to maize seeds, and novel Bt toxins for genetically engineered maize.
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Gassmann, A.J., Shrestha, R.B., Jakka, S.R.K., Dunbar, M.W., Clifton, E.H., Paolino, A.R., Ingber, D.A., French, B.W., Masloski, K.E., Doudna, J.W., St. Clair, C.R. 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. Journal of Economic Entomology: 109(4): 1872-1880
Abstract Western corn rootworm, Diabrotica virgifera virgifera LeConte (Coleoptera: Chrysomelidae), is a serious pest of corn in the United States, and recent management of western corn rootworm has included planting of Bt corn. Beginning in 2009, western corn rootworm populations with resistance to Cry3Bb1 corn and mCry3A corn were found in Iowa and elsewhere. To date, western corn rootworm populations have remained susceptible to corn producing Bt toxin Cry34/35Ab1. In this study, we used single-plant bioassays to test field populations of western corn rootworm for resistance to Cry34/35Ab1 corn, Cry3Bb1 corn, and mCry3A corn. Bioassays included nine rootworm populations collected from fields where severe injury to Bt corn had been observed and six control populations that had never been exposed to Bt corn. We found incomplete resistance to Cry34/35Ab1 corn among field populations collected from fields where severe injury to corn producing Cry34/35Ab1, either singly or as a pyramid, had been observed. Additionally, resistance to Cry3Bb1 corn and mCry3A corn was found among the majority of populations tested. These first cases of resistance to Cry34/35Ab1 corn, and the presence of resistance to multiple Bt toxins by western corn rootworm, highlight the potential vulnerability of Bt corn to the evolution of resistance by western corn rootworm. The use of more diversified management practices, in addition to insect resistance management, likely will be essential to sustain the viability of Bt corn for management of western corn rootworm.
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Gassmann, A.J., Petzold-Maxwell, J.L., Clifton, E.H., Dunbar, M.W., Hoffmann, A.H., Ingber, D.A., and Keweshan, R.S. 2014. Field-evolved resistance by western corn rootworm to multiple Bacillus thuringiensis toxins in transgenic maize. Proceedings of the National Academy of Sciences: 111(14), 5141-5146.
Abstract The widespread planting of crops genetically engineered to produce insecticidal toxins derived from the bacterium Bacillus thuringiensis (Bt) places intense selective pressure on pest populations to evolve resistance. Western corn rootworm is a key pest of maize, and in continuous maize fields it is often managed through planting of Bt maize. During 2009 and 2010, fields were identified in Iowa in which western corn rootworm imposed severe injury to maize producing Bt toxin Cry3Bb1. Subsequent bioassays revealed Cry3Bb1 resistance in these populations. Here, we report that, during 2011, injury to Bt maize in the field expanded to include mCry3A maize in addition to Cry3Bb1 maize and that laboratory analysis of western corn rootworm from these fields found resistance to Cry3Bb1 and mCry3A and cross-resistance between these toxins. Resistance to Bt maize has persisted in Iowa, with both the number of Bt fields identified with severe root injury and the ability western corn rootworm populations to survive on Cry3Bb1 maize increasing between 2009 and 2011. Additionally, Bt maize targeting western corn rootworm does not produce a high dose of Bt toxin, and the magnitude of resistance associated with feeding injury was less than that seen in a high-dose Bt crop. These first cases of resistance by western corn rootworm highlight the vulnerability of Bt maize to further evolution of resistance from this pest and, more broadly, point to the potential of insects to develop resistance rapidly when Bt crops do not achieve a high dose of Bt toxin.
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Petzold-Maxwell, J.L., Jaronski, S.T., Clifton, E.H., Dunbar, M.W., Jackson, M.A., and Gassmann, A.J. 2013. Interactions among Bt maize, entomopathogens, and rootworm species (Coleoptera: Chrysomelidae) in the field: effects on survival, yield, and root injury. Journal of Economic Entomology: 106(2), 622-632.
Abstract A 2 yr field study was conducted to determine how a blend of entomopathogens interacted with Bt maize to affect mortality of Diabrotica spp. (Coleoptera: Chrysomelidae), root injury to maize (Zea maize L.) and yield. The blend of entomopathogens included two entomopathogenic nematodes, Steinernema carpocapsae Weiser and Heterorhabditis bacteriophora Poinar, and one entomopathogenic fungus, Metarhizium brunneum (Metschnikoff) Sorokin. Bt maize (event DAS59122–7, which produces Bt toxin Cry34/35Ab1) decreased root injury and survival of western corn rootworm (Diabrotica virgifera virgifera LeConte) and northern corn rootworm (Diabrotica barberi Smith & Lawrence) but did not affect yield. During year 1 of the study, when rootworm abundance was high, entomopathogens in combination with Bt maize led to a significant reduction in root injury. In year 2 of the study, when rootworm abundance was lower, entomopathogens significantly decreased injury to non-Bt maize roots, but had no effect on Bt maize roots. Yield was significantly increased by the addition of entomopathogens to the soil. Entomopathogens did not decrease survival of corn rootworm species. The results suggest that soil-borne entomopathogens can complement Bt maize by protecting roots from feeding injury from corn rootworm when pest abundance is high, and can decrease root injury to non-Bt maize when rootworm abundance is low. In addition, this study also showed that the addition of entomopathogens to soil contributed to an overall increase in yield.