Insecticide Resistance Evolution in the Filth Fly Pupal Parasitoid

INSECTICIDE RESISTANCE EVOLUTION IN THE FILTH FLY PUPAL PARASITOID,

SPALANGIA CAMERONI (HYMENOPTERA: PTEROMALIDAE), USING LABORATORY

SELECTIONS

A Thesis

Presented to the faculty of the Department of Biological Sciences

California State University, Sacramento

Submitted in partial satisfaction of the requirements for the degree of

MASTER OF SCIENCE

Biological Sciences

(Ecology, Evolution, and Conservation)

Vincent Francis Maiquez

SPRING 2020

Vincent Francis Maiquez

INSECTICIDE RESISTANCE EVOLUTION IN THE FILTH FLY PUPAL PARASITOID,

SPALANGIA CAMERONI (HYMENOPTERA: PTEROMALIDAE), USING LABORATORY

SELECTIONS

A Thesis

Vincent Francis Maiquez

Approved by:

______, Committee Chair Dr. Jimmy Bruce Pitzer, Jr.

______, Second Reader Dr. Brett Holland

______, Third Reader Dr. Timothy Davidson

______Date

iii

Student: Vincent Francis Maiquez

I certify that this student has met the requirements for format contained in the University format manual, and this thesis is suitable for electronic submission to the library and credit is to be awarded for the thesis.

______, Graduate Coordinator ______Dr. James Baxter Date

Department of Biological Sciences

Abstract

INSECTICIDE RESISTANCE EVOLUTION IN THE FILTH FLY PUPAL PARASITOID,

SPALANGIA CAMERONI (HYMENOPTERA: PTEROMALIDAE), USING LABORATORY

SELECTIONS

Vincent Francis Maiquez

Arthropod pests cause economic losses to the agricultural industry through production losses and management costs. Among these pests, filth flies are prevalent in the animal industry, developing in decaying organic matter and can serve as mechanical vectors of disease-causing pathogens. In addition, most filth flies are considered nuisance pests, as their activities often disrupt the day-to-day activities of humans and animals. Filth flies are difficult to manage due to non-specific host-location preferences, their propensity to disperse great distances, and insecticide resistance evolution. However, insecticides continue to be the primary management method utilized by agricultural producers to reduce the impacts of these pests. Although commercially- available insecticides can be effective and fast-acting, they also can detrimentally impact alternative methods such as biological control organisms. Spalangia cameroni Perkins are beneficial parasitic wasps, which often are utilized as biological control organisms to manage filth fly populations such as house flies, Musca domestica L. and stable flies, Stomoxys calcitrans

(L.). Because these wasps search for filth fly pupae as hosts in areas likely treated with insecticides, they may be experiencing non-target insecticide selection effects. However, research regarding resistance evolution in parasitic wasps, including S. cameroni, is limited. This study was conducted to 1) determine the potential of S. cameroni to evolve resistance to permethrin, an insecticide often used to manage filth fly populations, under laboratory conditions and 2)

determine and compare permethrin susceptibility among several field-collected and long- established S. cameroni strains. Insecticide resistance evolution in this non-target organism has far-reaching implications for potential effects in other agricultural pest management systems. No tendency for increased or decreased susceptibility was observed for the selected strain. However, susceptibility was significantly reduced for the selected strain when compared to that of its unselected parent colony two years later. This was likely due to an absence of insecticidal selective pressures while maintained or an inability to evolve due to fixation of resistance-related genes. Results from geographic comparisons indicated that with the exception of the selected strain, all other strains exhibited similar susceptibility to permethrin as a susceptible strain. A second comparison indicated that an Insectary strain was significantly more susceptible than two field-collected strains. The implications of these findings include potential differences in parasitoid field efficacy and fitness due to differential insecticide susceptibility, and brings to light the potential of unrecognized non-target insecticide exposure effects in other pest systems.

______, Committee Chair Dr. Jimmy Bruce Pitzer, Jr.

______Date

ACKNOWLEDGEMENTS

I would like to thank my committee for their continued support and critique of my thesis. I thank my family for their encouragement and financial stability. I thank the Veterinary

Entomology laboratory volunteers, especially Jessica Navarro and Angela Mihalas for their thoughtfulness and hard work throughout my graduate studies. Last, and certainly not least, I would like to thank my graduate advisor, Dr. Jimmy Bruce Pitzer, Jr. His guidance and work ethic have been the backbone for my higher educational studies. Without his example I would not have come so far, so to him I say thanks, for the free parking.

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TABLE OF CONTENTS Page

Acknowledgements ...... vii

List of Tables ...... ix

Chapter

1. INTRODUCTION ………………………………………………………… ...... 1

Filth flies ...... 1

Chemical Insecticides ...... 4

Unstable Resistance ...... 6

Parasitoid Wasps ...... 7

2. MATERIALS AND METHODS ...... 10

Parasitoid Strains ...... 10

Serial Dilutions ...... 11

Insecticide Bioassay Test Vials ...... 11

Resistant Lethal Concentration Determination ...... 12

Insecticide Resistance Selection Procedure ...... 13

Multiple Strain Lethal Concentration Determination ...... 14

Statistical Analysis ...... 14

3. RESULTS ...... 15

4. DISCUSSION ...... 20

5. CONCLUSION ...... 24

Literature Cited ...... 25

viii

LIST OF TABLES Tables Page

1. The results of 10 permethrin selection procedures with corresponding LC50 values

for the Resistant Spalangia cameroni strain ...... ………………………………. 17

2. Corresponding LC50 values for the Resistant-, Parental 2016-, Parental 2018-,

Insectary-, Susceptible-, California-, Minnesota-, and Nebraska Spalangia cameroni

strains……………………………….… ...... ……………………………. 18

3. Permethrin resistance ratios for the Resistant-, Parental 2016-, Parental 2018-,

Insectary-, Susceptible-, California-, Minnesota-, and Nebraska Spalangia cameroni

strains…………………… ...... ………….…………………………………. 19

Introduction

Filth Flies

Filth flies such as house flies, Musca domestica L., and stable flies, Stomoxys calcitrans

(L.), are cosmopolitan medical- and veterinary pests that cause considerable economic damage to the animal industry world-wide. Although it has been difficult to quantify the economic losses attributed to their activities, house flies are known mechanical vectors of disease-causing pathogens to humans and livestock. Among these pathogens, Escherichia coli, the causal agent of hemorrhagic diarrhea, is known to be transferred by house flies from point sources to surfaces with which humans are likely to interact (Birkemoe et al. 2009, Talley et al. 2009). Alternatively, economic losses due to stable fly blood-feeding activities are approximately $2.21 billion USD annually in North America (Taylor et al. 2012). This economic impact is due to reductions in weight gain and milk production in beef- and dairy cattle, respectively, as well as associated increases in cattle feed costs to offset these losses. House fly and stable fly management has included methods such as varying trap configurations, livestock producer education, manure management, and synthetic and organic repellants (Morgan et al. 1988, Zhu et al. 2009,

Ramkumar and Shivakumar 2015). However, the detrimental effects filth flies have on livestock animals have led producers to rely heavily on insecticides, making it the primary management measure to mitigate fly populations generated at animal facilities (Olafson et al. 2011, Ramkumar and Shivakumar 2015).

Filth flies are known vectors of pathogens that affect humans and animals.

Approximately 73,000 cases of E. coli O157:H7, a strain known to cause hemorrhagic diarrhea, are documented annually in the United States (Mead et al. 1999). In September 2006, an E. coli

O157:H7 outbreak caused illness in over 200 individuals across the U.S. through the consumption of contaminated, pre-packaged spinach (CDC 2007). The Centers for Disease Control and

Prevention, in association with the Food and Drug Administration and the State of California

Department of Health Services Food and Drug Branch, implicated a spinach field, which was located in close proximity to a livestock production facility housing grass-fed cattle as the bacterial source.

In a related study conducted by Talley et al. (2009), several cattle production facilities were sampled in southern California to determine the diversity of fly species at these sites. Filth flies were observed in two of the four sites sampled, leading to a study regarding the transmission capabilities of field-collected house flies under laboratory conditions. To accomplish this, house flies first were allowed to interact with cattle manure, which had been sterilized and then contaminated with a modified toxin-negative strain of E. coli O157:H7. This bacterium was modified to express green fluorescent protein (GFP) so it could be visually identified on spinach plants that were later exposed to house flies from the manure contamination procedure. Spinach leaves were collected and observed using fluorescent microscopy to locate GFP-expressing bacteria. Results from the study indicated that the bacteria were mechanically-vectored by house flies from the treated cattle manure to the spinach leaves. This research provided a plausible explanation for the contamination that led to the E. coli outbreak in September 2006, whereby filth flies likely contracted the suspect bacteria from a nearby cattle facility and transferred it to the spinach fields.

Synanthropic flies, such as house flies, demonstrate ideal biological characteristics that support the transmission of human and animal pathogens between surfaces (Graczyk et al. 2001).

These include structures such as fine hairs and bristles, and the pulvillus (which assists in adhering the tarsus to surfaces), physiological processes such as frequent fecal deposition and regurgitation, behavioral characteristics such as frequent grooming, as well as the natural production of an electrostatic charge. All of these characteristics contribute to the ability of house

3 flies to collect, store, and release accumulated pathogens up to 3 km from contact points

(Murvosh and Thaggard 1966, Graczyk et al. 2001).

Antimicrobial resistance adds complexity to issues associated with filth fly pathogen transmission. A review by Onwugamba et al. (2018) highlighted the ability for filth flies to contract, develop, and disseminate microbes, particularly due to hospitable conditions provided by the fly crop, which can support the growth of several enteric pathogens. Bacteria stored in the crop reproduce until digested or regurgitated by flies during subsequent feedings. Although definitive evidence supporting the mechanical transfer of antimicrobial resistant bacterial pathogens by filth flies is limited due to the complexity of studying this interaction, it is possible that they are transferred similarly as are enteric bacteria to humans and animals (Onwugamba et al. 2018).

Stable flies are temporary, blood-feeding ectoparasites of cattle that generally land on either the foreleg or hindleg, puncture the epidermis with their mouth parts, and create a hematoma as a feeding site (Dougherty et al. 1995). Preference for these feeding sites is likely due to the presence of fewer hairs, blood vessels that are more closely located to the skin surface, as well as a reduced ability for the animal to defend these areas with deterrent behaviors such as tail swishes and head throws (Dougherty et al. 1995). Stable fly blood-feeding is demonstrated by both sexes; females use bloodmeal proteins for egg production, while males require bloodmeals for sexual reproductive system maturity (Isard et al. 2001). A complete bloodmeal feeding can require up to 4 min, but interrupted and periodic feeding behaviors are characteristic of these flies

(Isard et al. 2001).

Stable flies similarly affect humans by causing economic losses due to reductions in activity at recreational areas, as well as preventive measure costs associated with protecting other domesticated animals such as horses and dogs (Hogsette et al. 1987). Although not typical, areas

4 near the Great Lakes in the northern U.S. and the beaches of West Florida are known developmental sites for stable fly larvae that can produce substantial populations of this pest

(Hogsette et al. 1987, Isard et al. 2001). They also can be the result of meteorological events, such as low and high pressure changes between the inland and offshore areas that drive the insects to beaches where pressure zones converge, resulting in their subsequent feeding on humans and domesticated animals (Isard et al. 2001). In New Jersey and Florida, residents and visitors utilizing beach front resorts or parks as recreational sites, experience stable fly attacks that cause these tourists to end their stay prematurely, detrimentally affecting the local tourism industry

(Hansens 1951).

Large and small equine facility owners located near large bodies of water may experience an increase in stable fly activity due to meteorological pressure changes (Hogsette et al. 1987).

Much like other affected animals such as cattle, horses experiencing stable fly activity show signs of distress by excessive stomping, shoulder twitches, and head- and tail flicks (Muzari et al.

2010). These behaviors lead owners to provide additional care for their animals by applying insecticides, installing a variety of fly traps, and providing physical barriers against stably fly feeding activities (Mottet et al. 2018). The stable fly also is a serious pest of dogs, feeding chronically from ear tips resulting in permanent disfigurement, which is especially prevalent in penned animals (Hogsette et al. 1987, Yeruham and Brauerman 1995). Repeated attacks lead to necrotizing dermatitis, a bacterial infection of the soft tissues, leading some owners to amputate ears in pups as a preventive measure (Yeruham and Brauerman 1995).

Chemical Insecticides

Their tenacity, abundant numbers, and associated costs have resulted in the implementation of many techniques and devices to manage filth flies efficiently and reliably over the last 40 years. Historically, insecticides have been the most common management method

(Skovgård 2004). Of the available chemical insecticide classes, carbamates, organochlorines, organophosphates and pyrethroids are the most common. However, the pyrethroids, such as permethrin, account for approximately 25% of the total insecticide used world-wide due to its low mammalian toxicity, and reliability as a fast-acting toxicant (Williamson et al. 1995, Ramkumar and Shivakumar 2015).

The heavy, world-wide use of permethrin has led to selection effects manifested as resistance evolution in most pest species for which it is intended. This phenomenon has been the result of a variety of mutations in the two types of sodium channel genes that are present in insects, DSC1 and para. Pyrethroid-based insecticides bind to the para-type sodium channels within the insect nervous system causing varying responses such as the repetitive discharge of the sodium-gated channels (Type I) and nerve depolarization and blocking (Type II) depending on the pyrethroid type (Soderlund and Bloomquist 1989, Williamson et al. 1995, Silver et al. 2014).

Extensive use of this insecticide has led to resistance evolution in many insect species, due especially to changes in alleles associated with the para-type sodium channels. These changes have resulted in a reduced sensitivity to pyrethroid insecticides, which has been termed knockdown resistance or kdr (Williamson et al. 1995).

Among the numerous pathways resulting in insecticide resistance evolution, metabolism by cytochrome P450s also plays an important role (Riveron et al. 2013, Li et al. 2015). For example, the Halloween genes that encode the cytochrome P450 enzymes producing the hormone ecdysone, which is used by insects to signal molting and metamorphosis, are considered key genes for detoxification, and are conserved throughout the Arthropoda (Rewitz et al. 2007).

Additionally, research has demonstrated that mutations of neuron channel targets can prohibit functional binding of particular insecticide components (Matowo et al. 2010). A study conducted by Riveron et al. (2013) identified that the P450 genes CYP6P9a and CYP6P9b were responsible

6 for pyrethroid resistance in field populations of the mosquito, Anopholes funestus Giles, and verified them using the pyrethroid deltamethrin in bioassays on transgenic Drosophila melanogaster Morgan.

Unstable Resistance

While resistance evolution in insect pests has been widely studied, there are gaps in knowledge surrounding decreasing resistance in colonies maintained in laboratories. This phenomenon has been demonstrated by the tarnished plant bug, Lygus lineolaris Beauvois, the sheep body louse, Bovicola ovis Schrank, and the tobacco budworm, Heliothis virescens (F.). A study by Snodgrass (1996) focused on the interaction between field-collected L. lineolaris, a susceptible laboratory strain, and resistance levels from inter-mixed strains. It was concluded that permethrin resistance in the field-collected strain decreased by approximately 90% after nine generations in colony. Additionally, resistant and susceptible L. lineolaris strains were crossbred and maintained with no insecticide exposure for five generations, with the exception being one permethrin selection during the second generation. Snodgrass (1996) identified a 62.3% decline in permethrin resistance between the third and fifth generations. However, this was attributed to the production of additional, unselected generations between selection procedures that were required to generate population sizes adequate for experimentation. Snodgrass (1996) surmised that the lower LC25 used as the selective pressure likely reduced resistance levels in the colony.

A retrospective study conducted by Levot (2012) investigated data collected from several pyrethroid susceptibility assays conducted on the sheep louse, B. ovis, collected in Rowena,

Australia from 1992 to 2010. The study utilized LC50 values obtained from previous pyrethroid bioassays and were compared to a reference susceptible strain collected from Peak Hill, Australia.

The data demonstrated fluctuating resistance levels between 1992 and 2010, with higher LC50 values occurring after pyrethroid application years. From 1996 to 2003 no pyrethroid treatments

were applied. In 2008, treatments resumed until 2010, causing increases in the LC50 levels observed. Levot (2012) attributed the periodic increases in LC50 values to minimal reproductive costs in the resistant population, or a reduced number of susceptible individuals.

A study by Plapp (1981) investigated field populations of the tobacco budworm, H. virescens, by comparing their respective LC50 levels for permethrin, fenvalerate, cypermethrin, as well as combinations of those insecticides. While maintained in colony for five generations, two

LC50 determination assays were conducted. The author noted a substantial decrease in resistance between the first and second assays, which resulted in susceptibility comparable to that of a susceptible strain. It was likely that removal of the insecticidal pressure resulted in these increases in susceptibility.

Parasitoid Wasps

Resistance to insecticide use continues to grow among the public. Alternative management methods such as the use of biological control organisms are becoming increasingly- favored, as they represent more environmentally-conscious options (Skovgård 2004). Pupal parasitoids, such as Spalangia cameroni Perkins, Spalangia endius Walker, and Muscidifurax raptor Kogan and Legner, are commercially available as biological control organisms used to manage filth fly populations (Birkemoe et al. 2004, 2009, Skovgård 2004, Birkemoe and

Øyrehagen 2010, Pitzer et al. 2011, Machtinger et al. 2015). Parasitoid wasps likely encounter insecticides throughout their adult lifespan as they search for a suitable location that will likely harbor hosts they can use for feeding or oviposition (Gerling and Legner 1968). Species such as

S. cameroni, are strongly attracted to substrates that contain host odors, particularly those associated with fly larvae (Machtinger et al. 2015). Once a suitable habitat is located, a systematic search begins as S. cameroni navigates substrates such as calf bedding, silage, and hay piles at depths as far as 10 cm to locate hosts (Smith and Rutz 1991). Spalangia cameroni females spend

8 up to 130 min drumming and tapping the surface of a fly pupa before inserting their ovipositor to deposit an egg on the developing fly (Gerling and Legner 1968). Maturation times vary between the sexes, with males and females completing larval development at 32- and 35 days post- oviposition, respectively (Vinson 1976). Immediately following adult emergence, parasitoids begin their search for a suitable host environment and in doing so, will likely suffer exposures to applied insecticides intended for filth flies.

Spalangia spp. are known to seek and locate hosts that are deeper in substrates than other parasitoid species (Vinson 1976). Imidacloprid is a widely-used photo-stable neonicotinoid that is an effective filth fly insecticide used in several commercially-available granular bait formulations, which can persist in animal manure and affect both filth flies and their parasitoids

(Burgess and King 2016, Burgess et al. 2019). Additionally, this insecticide has a half-life of

<150-days in any soil type as deep as 30.5 cm (Mullins 1993). A study by Burgess and King

(2016) demonstrated interactions between filth fly insecticides and two parasitoid wasp species.

In their study, potential changes in S. endius behavior was assessed after imidacloprid exposure.

The results indicated that although S. endius was attracted to the insecticide, it did not experience any initial mortality effects. However, a decreased ability to parasitize house fly pupae was observed. In contrast, when hosts were buried in substrate, a shift towards control-level parasitism was achieved. Buried hosts provided an environment where S. endius was forced to search through substrates and in doing so, likely removed imidacloprid residues. Because S. cameroni shares similar behavioral characteristics with S. endius, particularly in searching for filth fly hosts, it too may be experiencing similar effects from granular filth fly insecticide bait applications.

A potential disadvantage of using biological organisms, is that some consumers may concurrently apply insecticides as a complementary method to manage filth fly populations.

Parasitoids likely frequent similar areas as adult filth flies, areas which also serve as application sites for insecticides and therefore, likely result in the unintentional reduction in parasitoid populations (Burgess & King 2015). Although treating substrates with insecticides is thought to be beneficial, there are potential future repercussions from these practices. In a study by Burgess et al. (2019), imidacloprid persistence in dairy cattle manure and its potential effects on development and effectiveness of S. endius as a biological control were assessed. It was determined that parasitoid management effectiveness was reduced overall, with house fly emergence rates of 95.8, 95.9, 10.6, and 35.9% for untreated manure with no parasitoids, treated manure with no parasitoids, untreated manure with parasitoids, and treated manure with parasitoids, respectively. Although fly emergence increased compared to the control, it was lower than in a previous study with unburied hosts and imidacloprid-treated parasitoids (Burgess and

King 2016).

Several studies have explored the interactions between Spalangia spp. and a variety of insecticides (Scott et al 1988, Burgess and King 2015, 2016, Burgess et al. 2019), but the evolutionary capabilities and possible application of resistant parasitoids is an area not typical of resistance studies. If pupal parasitoids are affected by insecticides through non-target pathways, the potential effects in other non-target insects may be similar. Therefore, the objective of this study was to 1) determine the potential for a common filth fly parasitoid, S. cameroni, to evolve insecticide resistance to the commonly-used insecticide, permethrin, using laboratory selections and 2) to determine susceptibility profiles for strains collected from California, Florida,

Minnesota, Nebraska, and Nevada to be used in a comparative analysis against a susceptible strain to highlight the effects of non-target chemical insecticide exposure and their far-reaching implications for other insects, beneficial or pestiferous.

Materials and Methods

Parasitoid Strains

Parasitoid wasps used in this study were maintained in 1-L plastic, screw-top cylindrical containers (Nalge Nunc International, Corp., Rochester, NY) fitted with three, evenly spaced 2.54 cm diameter circular brass screen disks (80 mesh) for air flow. All parasitoid rearing containers were held in a Percival incubator (Percival Scientific, Inc., Perry, Iowa) at 26  1 C with a photoperiod of 12:12 h (Light:Dark) and provided approximately 4,000 to 5,000 house fly pupae every seven days as a food- and host source, which allowed for the maintenance of approximately

1,200 adults per strain. A total of six S. cameroni strains (~1,200 parasitoids per strain) were received from Dr. Christopher Geden at the United States Department of Agriculture,

Agricultural Research Service, Center for Medical and Veterinary Entomology in Gainesville, FL in 2016. When collected from field populations, S. cameroni strains were colonized from no less than 300 female parasitoids and were maintained in similar conditions as previously described.

The Susceptible strain, which was collected from a dairy farm near Bell, FL in 2000, served as a baseline for all permethrin susceptibility comparisons because it is the longest-lived colony known to be reared in the absence of any insecticidal selective pressures. The Parental strain, which was collected from a dairy farm near Bell, FL in 2012, was used to generate the permethrin

Resistant strain (with this newly produced Resistant colony representing a seventh strain by the end of the study) using laboratory selections. It was chosen as the Parental strain because it had been most recently colonized and maintained for a period long enough to consistently produce quantities required for this study. Additionally, the Parental strain originated from a site known to use permethrin for filth fly management, suggesting it would be an appropriate strain to demonstrate potential insecticide evolution. The California (collected from a dairy farm near

Moreno Valley, CA in 2015), Minnesota (single colony derived from collections from several

11 dairy farms in MN in 2015), and Nebraska (collected from a dairy farm near Firth, NE in 2014) strains were field strains used to compare permethrin susceptibility profiles of parasitoids collected from different regions in the U.S. The Insectary strain was purchased from Spalding

Laboratories in Reno, NV in 2017 and also was used in permethrin susceptibility comparisons.

These field strains were used in comparison assays due to the limited availability of parasitoid strains maintained in the U.S., and because they were the most recently collected field populations known. This allowed for the most accurate assessment regarding current permethrin susceptibility profiles of S. cameroni populations in the U.S. to date.

Serial Dilutions

All permethrin serial dilutions used for this study were made from newly-created stock solutions having an initial concentration of 1,000 µg/mL. To achieve this concentration, a 0.021 g weight of technical grade permethrin (95%, Lot Number: PMN940T5656B, Central Life

Sciences, Dallas, TX) was dissolved in 20 mL of laboratory grade acetone (≥99.5%, Fisher

Scientific). To determine the susceptibility profile of the Parental strain, a previously-determined

2 concentration that would cause 50% mortality in the population (LC50) of 0.148649 µg/cm for female S. cameroni was used (Scott et al. 1988). Permethrin dilutions of 10x and 1/10x this concentration were made in an attempt to encompass a concentration response range from 0 to

100% mortality. This method allowed for the preparation of an entire serial dilution set that would provide data appropriate for statistical analysis.

Insecticide Bioassay Test Vials

Glass scintillation vials (20 mL) (VWR International) were used to determine all permethrin susceptibility profiles in this study. The internal dimensions of the vials were 7.85 x

4.4 x 2.4 cm (circumference, height, diameter), which was used to calculate the surface area of

39.06 cm2. A 1-mL aliquot of each permethrin serial dilution was applied to the inside of its

12 respective, previously-labeled glass scintillation vial and placed on an electric hot dog roller for at least 60 min. This process ensured a uniform insecticide distribution, as well as complete evaporation and drying of the acetone solvent.

Resistant Lethal Concentration Determination

To assess the potential for resistance evolution by S. cameroni using laboratory selections with permethrin, procedures similar to those described by Pitzer et al. (2010) were followed.

Briefly, an appropriate concentration range (0, 9.375, 1.875x10, 3.750x10, 7.500x10, and

1.500x102 g permethrin/mL) was serially-diluted from a previously-prepared stock solution of

1.000 x103 g permethrin/mL and applied to 20 mL glass scintillation vials. Three treatment vials were prepared for each serially-diluted concentration, as well as the acetone-only controls. Ten,

6-10 day-old female parasitoids from the Parental parasitoid strain (F0 of the new Resistant strain) were placed into each vial (n=30 for each treatment/control) for a 4-h exposure period. Parasitoid mortality was assessed at 5 min (handling mortality <1%) and 4-h post-exposure. After the 4-h post-exposure mortality assessment, all parasitoids were moved to clean, 120-ml plastic soufflé cups (Dart Container Corporation, Mason, MI) fitted with a 2.54 cm-long dental wick saturated with a 10% sucrose solution for an additional 44-h (48-h post-exposure mortality assessment).

Any surviving parasitoids were euthanized by being placed in a freezer. This represented one experimental replication. Three experimental replications were performed concurrently for each strain to generate individual strain susceptibility profile determinations (n=90 for each treatment/control). The data from these three replications were pooled prior to Probit analysis (Li et al. 2015, Marcombe et al. 2012, Matowo et al. 2010, Pitzer et al. 2010, Yanola et al. 2010,

Scott 1998) using IBM SPSS Statistics software (IBM SPSS Statistics for Windows, Version 26.0.

2019). The results of the Probit analysis for the Resistant strain allowed for the identification of a generation-specific target LC50 value, which was determined for the resultant offspring produced

13 by adults of each individual selection procedure, and was applied to the entire Resistant strain during the insecticide resistance selection procedures.

Insecticide Resistance Selection Procedures

The Probit-estimated LC50 was used to treat the remaining parasitoids in the Resistant (F0) strain. Prior to these generation-wide selection procedures, all house fly pupae previously exposed to parasitoids for feeding and oviposition were placed individually into 48-well cell culture plates (Corning, Inc.) to prevent mating upon adult emergence. All newly emerged parasitoids were separated by sex from the culture plates and placed into 120-ml plastic soufflé cups with a 2.54 cm-long sucrose solution-saturated dental wick until they are used in the selection procedure. When all parasitoids were 6-10 days old, they were placed into treated glass scintillation vials and monitored as described previously. At 48 h post-exposure however, all surviving male and female parasitoids were combined into one parasitoid-rearing container to allow for mating, and subsequent oviposition into house fly pupae that were provided ad-libitum.

The resulting offspring were used for subsequent selection procedures, which included a new

LC50 assessment for the current generation. This process allowed for a continuously-adaptive selective pressure at the 50% mortality target. Due to low survivorship post-selection, an additional generation was produced between selection procedures when needed, which allowed for an adequate population to conduct a minimum of two LC50 determination replicates, as well as a generation-wide selection procedure. This protocol was repeated for a total of 10 selection procedures, spanning a total of 15 generations; a protocol which has been adequate for similar studies of different insect species (Marcombe et al. 2012, Matowo et al. 2010, Pitzer et al. 2010,

Yanola et al. 2010, Scott 1998). After the final selection procedure, the Resistant strain was provided hosts, and a final LC50 value was determined using offspring produced in the next generation.

Multiple Strain Lethal Concentration Determination

For all seven strains used in this study, a corresponding LC50 determination procedure was conducted following the final selection on the Resistant strain as previously described.

Briefly, each LC50 was provided by Probit analysis using mortality data collected from three experimental replications comprised of three trials (three vials containing 10, 6-10 day-old female

S. cameroni parasitoids) per concentration. The permethrin concentration range used for all strains and assays in this portion of the study was 0.000, 4.688, 9.375, 1.875x10, 3.750x10,

7.500x10, and 1.500x102 g permethrin/mL.

Statistical Analysis

All statistical procedures were performed using IBM SPSS Statistics 26 (IBM SPSS

Statistics for Windows, Version 26.0. 2019). Pooled mortality data was subjected to Probit analysis, which provided susceptibility profiles (LC values). The Probit procedure, which is a logistic regression used to analyze the relationship between a stimulus and its corresponding quantal response, was used to identify strain and generation-specific LC50 values. Corresponding

Resistance Ratios (RR) were calculated by dividing the LC50 value of the strain in question by a reference strain, which provided a relative comparison between them. Relative median potency

(RMP) analysis was used to determine the significance of the RR. This analysis calculates differences between two different or similar toxicants by dividing the median LC50 of a known toxicant with that of an unknown. Significant differences in the RR values between parasitoid strains were determined when the confidence interval (CI) generated from the comparison did not overlap with the value “1” (Benelli et al. 2015, Gaire et al. 2019).

Results

Resistance Evolution

The primary goal of this study was to determine if S. cameroni was capable of resistance evolution due to exposure to the insecticide active ingredient, permethrin. The results from each

LC50 determination procedure provided a concentration that was used as the selective pressure for the next selection procedure, which ranged from 15.232 to 29.146 µg permethrin/mL (Table 1).

In general, no tendency for an increase or decrease in LC50 concentration was observed during the course of the selection procedures. Both the LC50 values for the Parental strain determined in

2016 and again in 2018 were compared to that of the Resistant strain (Table 3). The Insectary,

Susceptible, California, Minnesota, and Nebraska strains also were compared to the Resistant strain. The final LC50 values for the Resistant, Parental 2016, and Parental 2018, Insectary,

Susceptible, California, Minnesota, and Nebraska strains were 23.086, 21.816, 12.012, 11.879,

15.211, 11.986, 14.449, and 15.056 µg permethrin/mL, respectively (Table 2). This resulted in

RR calculations comparing the Resistant strain with the Parental 2016, Parental 2018, Insectary,

Susceptible, California, Minnesota, and Nebraska strains (Table 3). The Parental 2018, Insectary,

Susceptible, California, Minnesota, and Nebraska strains were significantly more susceptible to permethrin than the Resistant strain, but there was no difference when the Resistant strain was compared to the Parental 2016 strain.

Geographic Insecticide Resistance

The LC50 values for the Resistant, Parental 2016, Parental 2018, Insectary, California,

Minnesota, and Nebraska strains were compared to the Susceptible strain (Table 3). The results indicated that only the Resistant strain was significantly more resistant when compared to the

Susceptible strain. All other strains were not significantly different from the Susceptible strain.

Based on the LC50 values for the Susceptible and Insectary strains, it appeared that the Insectary strain was more susceptible to permethrin than the Susceptible strain. Therefore, a second, similar comparison was made, whereby all strains were compared to the Insectary strain (Table 3). From this comparison the Resistant, Parental 2016, Minnesota, and Nebraska strains were significantly more resistant than the Insectary strain, whereas the Parental 2018, Susceptible, and California strains were not.

Table 1. The results of 10 permethrin selection procedures with corresponding LC50 values for the

Resistant Spalangia cameroni strain.

a b, c d Selection F LC50 (95% CI) n Slope (95% CI)

1 4 29.146 (25.237 – 33.586) 530 4.421 (3.336 – 5.507)

2 6 26.712 (23.173 – 30.727) 240 4.613 (3.498 – 5.727)

3 7 18.669 (17.231 – 20.240) 460 6.914 (5.392 – 8.437)

4 9 26.991 (23.945 – 30.307) 510 4.712 (3.907 – 5.517)

5 10 18.048 (15.598 – 20.793) 230 4.577 (3.335 – 5.820)

6 11 22.285 (20.628 – 24.059) 540 6.598 (5.290 – 7.906)

7 13 20.626 (17.478 – 24.058) 530 3.277 (2.740 – 3.814)

8 15 15.232 (13.565 – 17.095) 230 6.989 (4.849 – 9.128)

9 17 18.817 (16.547 – 21.450) 240 5.488 (3.983 – 6.994)

10 19 17.095 (13.453 – 20.824) 230 2.634 (1.932 – 3.337) aSpalangia cameroni Resistant generation selected bValues represent micrograms of permethrin per milliliter of acetone applied to 20 mL glass scintillation vials. c Values were back calculated from log10 concentration values provided by Probit analysis in IBM SPSS

Statistics (SPSS). dTotal number of female parasitoid wasps evaluated.

Table 2. Corresponding LC50 values for the Resistant-, Parental 2016-, Parental 2018-, Insectary-,

Susceptible-, California-, Minnesota-, and Nebraska Spalangia cameroni strains.

a, b c Parasitoid Strain LC50 (95% CI) n Slope (95% CI)

Resistant 23.086 (20.423 – 26.115) 530 4.864 (4.038 – 5.691)

Parental 2016 21.816 (18.188 – 26.206) 360 3.775 (3.061 – 4.490)

Parental 2018 12.012 (11.161 – 12.935) 540 7.285 (5.830 – 8.741)

Insectary 11.879 (10.470 – 13.457) 510 4.841 (4.021 – 5.661)

Susceptible 15.211 (11.348 – 20.274) 540 2.802 (2.375 – 3.230)

California 11.986 (10.519 – 13.696) 540 4.300 (3.611 – 4.989)

Minnesota 14.449 (12.874 – 16.245) 540 5.574 (4.592 – 6.556)

Nebraska 15.056 (13.223 – 17.139) 530 5.272 (4.344 – 6.201) aValues represent micrograms of permethrin per milliliter of acetone applied to 20 mL glass scintillation vials. b Values were back calculated from log10 concentration values provided by Probit analysis in IBM SPSS

Statistics (SPSS). cTotal number of female parasitoid wasps evaluated.

Table 3. Permethrin resistance ratios for the Resistant-, Parental 2016-, Parental 2018-, Insectary-,

Susceptible-, California-, Minnesota-, and Nebraska Spalangia cameroni strains.

a b c Parasitoid Strain RR (95% CI) RR (95% CI) RR (95% CI)

Resistant 1.000 1.507 (1.078 – 2.316)* 1.943 (1.538 – 2.610)*

Parental 2016 1.059 (0.852 – 1.328) 1.438 (0.995 – 2.255) 1.840 (1.393 – 2.639)*

Parental 2018 1.917 (1.511 – 2.628)* 1.270 (0.925 – 1.833) 1.010 (0.865 – 1.179)

Insectary 1.943 (1.538 – 2.610)* 0.769 (0.532 – 1.064) 1.000

Susceptible 1.507 (1.078 – 2.316)* 1.000 1.300 (0.940 – 1.881)

California 1.930 (1.513 – 2.629)* 0.787 (0.546 – 1.081) 1.008 (0.841 – 1.207)

Minnesota 1.598 (1.315 – 2.026)* 0.952 (0.694 – 1.298) 1.216 (1.038 – 1.449)*

Nebraska 1.533 (1.259 – 1.952)* 0.985 (0.717 – 1.356) 1.267 (1.065 – 1.542)*

a Resistance ratios are calculated by dividing the LC50 of the Resistant strain by the LC50 of another strain.

b Resistance ratios are calculated by dividing the LC50 of a strain by the LC50 of the Susceptible strain.

c Resistance ratios are calculated by dividing the LC50 of a strain by the LC50 of the Insectary strain.

*Values were determined by Relative Median Potency (RMP) analysis and were significantly different if

the provided 95% CI range does not overlap with the value “1”.

Discussion

Resistance Evolution

The results of this study suggest S. cameroni exhibit selective effects due to permethrin exposures under laboratory conditions. However, the results from the selection procedures did not align with the prediction that the LC50 value would increase dramatically. Instead the LC50 values fluctuated between selected generations. Periodically, an unselected interim generation between selections was used to increase the population size, which may have restricted resistance evolution. This phenomenon has been observed in other field-collected insect pest species that were maintained without an insecticidal selective pressure (Plapp 1981, Snodgrass 1996).

Alternatively, the lack of resistance evolution may be attributed to the Parental strain previously being exposed to permethrin resulting in substantial resistance evolution in the field population.

Therefore, pre-existing, low frequency, genetic variation for permethrin resistance may have gone to fixation prior to collection of the population. This could reduce the rate of adaptive evolution under the conditions used in this experiment, because new resistance mutations would be rare.

This is a plausible explanation for the unresponsive LC50 value between the initial assessment of the Parental strain in 2016, and that for the Resistant strain in 2018.

After the final selection procedure, the Resistant strain was significantly more resistant than the Parental strain in 2018, which suggests that the Resistant strain had evolved insecticide resistance (Table 3). To confirm this, the Resistant strain was compared to the Parental 2016 strain to ensure the Parental strain had remained similarly resistant while maintained in colony.

The RR calculated for this comparison suggested that the two strains were not significantly different from one another, likely meaning that the Resistant strain had maintained its resistance over time while that of the Parental strain had decreased until its second evaluation in 2018

(Table 3). It is possible that a lack of an insecticide selective pressure from an insecticide is the

21 cause for this reduction in resistance over time (Plapp 1981, Snodgrass 1996, and Levot 2012).

The results of the inter-strain comparison generated RR values that indicated the Resistant strain was significantly more resistant than the Insectary, Susceptible, California, Minnesota, and

Nebraska strains.

Geographic Insecticide Resistance

The second goal for this study was to determine susceptibility profiles of S. cameroni strains from varying geographic locations and to compare these profiles with a known Susceptible strain to determine potential differences in susceptibility (Table 3). The field-collected strains,

California, Minnesota, and Nebraska did not differ significantly from the Susceptible strain.

When the Resistant and Parental 2016 and 2018 strains were compared, the difference in RR values between both Parental strains was likely due to the Parental strain remaining unexposed in colony for two years. With the exception of the California strain, it is possible that the change in significance observed also occurred in the field-collected strains, which had been maintained unexposed in colony for several years. The LC50 values for the Resistant and Insectary strains also were compared to the Susceptible strain (Table 3). Only the Resistant strain was significantly more resistant than the Susceptible strain, which is due either to successful population selections leading to resistance evolution, or that the selection procedures maintained a level of resistance already present in the Parental 2016 strain.

The Insectary strain had the lowest LC50 value of all other strains, which suggested that it was more susceptible than the Susceptible strain (Table 2). This difference warranted further investigation, using the Insectary strain as a baseline comparison for all other parasitoid strains.

The results of a subsequent Probit and accompanying RMP analysis indicated that all but three, the Parental 2018, Susceptible, and California strains, were significantly more resistant than the

Insectary strain (Table 3). This suggested that the Insectary strain had been maintained in colony

22 long enough to reduce any acquired resistance to more susceptible levels. When the Resistant strain was compared to the Insectary and Susceptible strains, both were significantly more susceptible than the Resistant strain (Table 3). This suggested that the Insectary and Susceptible strains were equally susceptible to permethrin.

Due to the findings regarding the Insectary strain, it is likely that commercially available

S. cameroni from this insectary, and possibly others, may be as vulnerable to permethrin as a susceptible strain. If changes in resistance, such as those in this study, are occurring in insectary populations, then it is in the interest of parasitoid consumers to consider that insectary-reared S. cameroni may not be as insecticide-tolerant as their field-strain counterparts. Not only can mass- releases of less susceptible parasitoids result in reduced filth fly management, but it can also potentially increase susceptibility in field populations.

Far-reaching implications of this study include non-target effects, such as unintended insecticide resistance evolution, that can potentially be experienced by other beneficial and pestiferous insects. Insecticide use is widespread and includes application formulations such as pour-on, granular baits, and space sprays. Some of these application formulations, such as granular baits, can persist and remain effective at an application site for at least 168 h post- application, depending on the environment and anthropological activities in the area (Burgess and

King 2016, Murillo et al. 2018, Burgess et al. 2019). With insecticidal chemicals that have the potential to remain lethal days after application, it is possible that other beneficial and pestiferous insects are exposed and evolve insecticide resistance. Although it can be potentially advantageous for these beneficial insects to evolve insecticide resistance, they may not be affected on the same scale or rate as the intended targets. In field populations, S. cameroni is capable of burrowing through substrates to locate hosts, and through this action, can remove contacted insecticide residues from themselves, effectively reducing the amount they absorb. Rearing resistant

23 biological controls has the potential to be beneficial, not only to consumers but to insectaries, and programs for developing resistant parasitoid wasps should be reviewed. These programs would likely benefit field strains and have the potential for parasitoids to be utilized more readily alongside certain chemical insecticides. Therefore, future study regarding the performance of resistant filth fly parasitoids is warranted.

Conclusion

With the continued world-wide use of chemical insecticides, non-target exposure events can potentially occur to both intended pest and beneficial insect species due to their association with application sites. The ability for a biological control organism to evolve insecticide resistance is especially relevant in the ongoing arms race between the agricultural industry and its arthropod pests. Future directions for this study include investigating the interaction and potential performance differences between resistant and susceptible parasitoid strains and quantifying the evolution of susceptibility in parasitoid species in the absence of insecticidal selective pressure.

Spalangia cameroni are common beneficial organisms that commonly are encountered where filth fly species develop. The results of this study further our understanding of potential insecticide and parasitoid interaction effects and are a primer for potentially enhancing filth fly parasitoid performance for stakeholders by providing valuable information to commercial insectaries. Increasing filth fly parasitoid performance ultimately, will result in decreasing insecticide use and its effects on humans, animals, and the environment, while increasing animal health and their production of economically-important goods.

Literature Cited

Benelli, G., S. Bedini, G. Flamini, F. Cosci, P. L. Cioni, S. Amira, F. Benchikh, H. Laouer,

G. Di Giuseppe, and B. Conti. 2015. Mediterranean essential oils as effective weapons

against the West Nile vector Culex pipiens and the Echinostoma intermediate host

Physella acuta: what happens around? An acute toxicity survey on non-target mayflies.

Parasitol Res. 114: 1011–1021.

Birkemoe, T., and H. Øyrehagen. 2010. Parasitism of the house fly parasitoid Spalangia

cameroni on Norwegian pig farms: local effect of release method. BioControl. 55: 583–

591.

Birkemoe, T., A. Soleng, and A. Aak. 2009. Biological control of Musca domestica and

Stomoxys calcitrans by mass releases of the parasitoid Spalangia cameroni on two

Norwegian pig farms. BioControl. 54: 425–436.

Birkemoe, T., A. Soleng, and K. W. Riddervold. 2004. Abundance of parasitoid Hymenoptera

on pupae of Musca domestica and Stomoxys calcitrans (Diptera, Muscidae) on pig farms

in Vestfold, Norway. Norw J Entomol. 51: 159–164.

Burgess, E. R., and B. H. King. 2015. Compatibility of the parasitoid wasp Spalangia endius

(Hymenoptera: Pteromalidae) and insecticides against Musca domestica (Diptera:

Muscidae) as evaluated by a new index. J Econ Entomol. 108: 986–992.

Burgess, E. R., and B. H. King. 2016. Behavior and survival of the filth fly parasitoids

Spalangia endius and Urolepis rufipes (Hymenoptera: Pteromalidae) in response to three

granular house fly baits and components. Environ Entomol. 45: 1496–1504.

Burgess, E. R., S. M. Watkins, B. H. King, K. Chantos-Davidson, A. N. Kremer, J. C.

Tournear, J. Morrow, T. J. Hagen, and E. R. Gaillard. 2019. Dissemination of

imidacloprid through dairy cattle manure and its effect on the biological control agent,

Spalangia endius (Hymenoptera: Pteromalidae), and a filth fly host, Musca domestica

(Diptera: Muscidae). J Econ Entomol. 112: 974–980.

(CDC) Centers for Disease Control and Prevention. 2007. Investigation of an Escherichia coli

O157:H7 outbreak associated with Dole pre-packaged spinach.

(https://www.cdc.gov/nceh/ehs/docs/investigation_of_an_e_coli_outbreak_associated_wi

th_dole_pre-packaged_spinach.pdf). CDC, Atlanta, Geogria.

Dougherty, C. T., F. W. Knapp, P. B. Burrus, D. C. Willis, and P. L. Cornelius. 1995.

Behavior of grazing cattle exposed to small populations of stable flies (Stomoxys

calcitrans L.). Appl Anim Behav Sci. 42: 231–248.

Gaire, S., M. E. Scharf, and A. D. Gondhalekar. 2019. Toxicity and neurophysiological

impacts of plant essential oil components on bed bugs (Cimicidae: Hemiptera). Sci Rep.

9: 3961.

Gerling, D., and E. F. Legner. 1968. Developmental history and reproduction of Spalangia

cameroni, parasite of synanthropic flies. Ann Entomol Soc Am. 61: 1436–1443.

Graczyk, T. K., R. Knight, R. H. Gilman, and M. R. Cranfield. 2001. The role of non-biting

flies in the epidemiology of human infectious diseases. Microbes Infect. 3: 231–235.

Hansens, E. J. 1951. The stable fly and its effect on seashore recreational areas in New Jersey. J

Econ Entomol. 44: 482–487.

Hogsette, J. A., J. P. Ruff, and C. J. Jones. 1987. Stable fly biology and control in northwest

Florida. J Agric Entomol. 4: 1–11.

IBM SPSS Statistics for Windows. 2019. Version 26.0. IBM Corp., Armonk, NY.

Isard, S. A., D. A. R. Kristovich, S. H. Gage, C. J. Jones, and N. F. Laird. 2001. Atmospheric

motion systems that influence the redistribution and accumulation of insects on the

beaches of the Great Lakes in North America. Aerobiologia. 17: 275–291.

Levot, G. 2012. Unstable pyrethroid resistance in sheep body lice Bovicola ovis (Shrank),

(Phthiraptera: Trichodectidae) and its implications for lice control on sheep. Vet

Parasitol. 185: 247–278.

Li, T., C. Cao, T. Yang, L. Zhang, L. He, Z. Xi, G. Bian, and N. Liu. 2015. A G-protein-

coupled receptor regulation pathway in cytochrome P450-mediated permethrin-resistance

in mosquitoes, Culex quinquefasciatus. Sci Rep. 5: 17772, doi: 10.1038/srep17772.

Machtinger, E. T., C. J. Geden, P. E. Teal, and N. C. Leppla. 2015. Comparison of host-

seeking behavior of the filth fly pupal parasitoids, Spalangia cameroni and Muscidifurax

raptor (Hymenoptera: Pteromalidae). Environ Entomol. 44: 330–337.

Marcombe, S., R. B. Mathieu, N. Pocquet, M. Riaz, R. Poupardin, S. Sélior, F. Darriet, S.

Reynaud, A. Yébakima, V. Corbel, J. David, and F. Chandre. 2012. Insecticide

resistance in the dengue vector Aedes aegypti from Martinique: distribution, dechanisms

and relations with environmental factors. Plos One. 7: e30989,

doi:10.1371/journal.pone.0030989.

Matowo, J., M. A. Kulkarni, F. W. Mosha, R. M. Oxborough, J. A. Kitau, F. Tenu, and M.

Rowland. 2010. Biochemical basis of permethrin resistance in Anopheles arabiensis

from Lower Moshi, north-eastern Tanzania. Malaria J. 9: 193–202.

Mead, P. S., L. Slutsker, V. Dietz, L. F. McCaig, J. S. Bresee, C. Shapiro, P. M. Griffin, and

R. V. Tauxe. 1999. Food-related illness and death in the United States. Emerg Infect Dis.

5: 607–625.

Morgan, P. B., R. S. Patterson, and G. D. Propp. 1988. Manure management as a means of

controlling filth flies at agricultural installations. Fla Entomol. 71: 44–49.

Mottet, R. S., R. D. Moon, M. R. Hathaway, and K. L. Martinson. 2018. Effectiveness of

stable fly protectants on adult horses. J Equine Vet Sci. 69: 11–15.

Mullins, J. W. 1993. Imidacloprid: a new nitroguanidine insecticide, pp. 183–198. In Pest

Control with Enhanced Environmental Safety, ACS Symposium Series. American

Chemical Society, Washington, DC.

Murillo, A. C., D. Cox, and B. A. Mullens. 2018. Longevity of fly baits exposed to field

conditions. J Econ Entomol. 111: 967–973.

Murvosh, C. M., and C. W. Thaggard. 1966. Ecological studies of the house fly. Ann Entomol

Soc Am. 59: 533–547.

Muzari, M. O., L. F. Skerratt, R. E. Jones, and T. L. Duran. 2010. Alighting and feeding

behavior of tabanid flies on horses, kangaroos and pigs. Vet Parasitol. 170: 104–111.

Olafson, P. U., J. B. Pitzer, P. E. Kaufman. 2011. Identification of a mutation associated with

permethrin resistance in the para-type sodium channel of the stable fly (Diptera:

Muscidae). J Econ Entomol. 104: 250–7.

Onwugamba, F. C., J. R. Fitzgerald, K. Rochon, L. Guardabassi, A. Alabi, S. Kühne, M. P.

Grobusch, and F. Schaumburg. 2018. The role of ‘filth flies’ in the spread of

antimicrobial resistance. Travel Med Infect D. 22: 8–17.

Pitzer, J. B., P. E. Kaufman, S. H. Tenbroeck. 2010. Assessing permethrin resistance in the

stable fly (Diptera: Muscidae) in Florida by using laboratory selections and field

evaluations. J Econ Entomol. 103: 2258–63.

Pitzer, J. B., P. E. Kaufman, C. J. Geden, and J. A. Hogsette. 2011. The Ability of Selected

Pupal Parasitoids (Hymenoptera: Pteromalidae) to Locate Stable Fly Hosts in a Soiled

Equine Bedding Substrate. Environ Entomol. 40: 88–93.

Plapp, F. W. Jr. 1981. Toxicity of synthetic pyrethroids to laboratory and field populations of

tobacco budworm in central Texas. J Econ Entomol. 74: 207–209.

Ramkumar, G., and M. S. Shivakumar. 2015. Laboratory development of permethrin

resistance and cross-resistance pattern of Culex quinquefasciatus to other insecticides.

Parasitol Res. 114: 2553–2560.

Rewitz, K. F., M. B. O’Connor, and L. I. Gilbert. 2007. Molecular evolution of the insect

Halloween family of cytochrome P450s: phylogeny, gene organization and functional

conservation. Insect Biochem Molec. 37: 741–753.

Riveron, J. M., H. Irving, M. Ndula, K. G. Barnes, S. S. Ibrahim, M. J. I. Paine, and C. S.

Wondji. 2013. Directionally selected cytochrome P450 alleles are driving the spread of

pyrethroid resistance in the major malaria vector Anopheles funestus. P Natl Acad Sci-

Biol. 110: 252–257.

Scott, J. G. 1998. Toxicity of spinosad to susceptible and resistant strains of house flies, Musca

domestica. Pesti Sci. 54: 131–133.

Scott, J. G., D. A. Rutz, and J. Walcott. 1988. Comparative toxicity of seven insecticides to

adult Spalangia Cameroni Perkins.J Agric Entomol. 5: 139–145.

Silver, K. S., Y. Du, Y. Nomura, E. E. Oliveira, V. L. Salgado, B. S. Zhorov, and K. Dong.

2014. Voltage-gated sodium channels as insecticide targets. Adv Insect Physiol. 46: 389–

433.

Skovgård, H. 2004. Sustained releases of the pupal parasitoid Spalangia cameroni

(Hymenoptera: Pteromalidae) for control of house flies, Musca domestica and stable flies

Stomoxys calcitrans (Diptera: Muscidae) on dairy farms in Denmark. Biol Control. 30:

288–297.

Snodgrass, G. L. 1996. Insecticide resistance in field populations of the tarnished plant bug

(Heteroptera: Miridae) in cotton in the Mississippi delta. J Econ Entomol. 89: 783–790.

Soderlund, D. M. and J. R. Bloomquist. 1989. Neurotoxic actions of pyrethroid insecticides.

Ann Rev Entomol. 34: 77–96.

Smith, L., and D. A. Rutz. 1991. Microhabitat associations of hymenopterous parasitoids that

attack house fly pupae at dairy farms in central New York. Environ Entomol. 20: 675–

684.

Taylor, D. B., and D. Berkebile. 2006. Comparative efficiency of six stable fly (Diptera:

Muscidae) traps. J Econ Entomol. 99: 1415–1419.

Taylor, D. B., R. D. Moon, and D. R. Mark. 2012. Economic impact of stable flies (Diptera:

Muscidae) on dairy and beef cattle production. J Med Entomol. 49: 198–209.

Vinson, S. B. 1976. Host selection by insect parasitoids. Annu Rev Entomol. 21: 109–133.

Williamson, M. S., D. Martinez-Torres, C. A. Hick, and A. L. Devonshire. 1996.

Identification of mutations in the housefly para-type sodium channel gene associated

with knockdown resistance (kdr) to pyrethroid insecticides. Mol Gen Genet. 252: 51–60.

Yanola, J., P. Somboon, C. Walton, W. Nachaiwieng, and L. Prapanthadara. 2010. A novel

F1552/C1552 point mutation in the Aedes aegypti voltage-gated sodium channel gene

associated with permethrin resistance. Pestic Biochem Phys. 96: 127–131.

Yeruham, I and Y. Braverman. 1995. Skin lesions in dogs, horses and calves caused by the

stable fly Stomoxys calcitrans (L.) (Diptera: Muscidae). Rev Elev Med Vet Pay Trop.

48:347–349.

Zhu, J. J., X. P. Zeng, D. Berkebile, H. J. Du, Y. Tong, and K. Qian. 2009. Efficacy and

safety of catnip (Nepeta cataria) as a novel filth fly repellent. Med Vet Entomol. 23:

209–216.