House Fly (Diptera: Muscidae): Biology, Pest Status, Current Management Prospects, and Research Needs

Abstract

The house fly, Musca domestica L. (Diptera: Muscidae), is a global pest of humans and animals that carries scores of pathogens and costs up to $1 billion per year in the United States alone. Information is reviewed on recognition, distribution, biology, dispersal, and associations with microbes. Particular challenges of managing flies in different animal systems are discussed for swine, poultry, dairy cattle, beef feedlot, and equine operations. Effective fly management requires diligent monitoring and integration of cultural control, especially manure management, with mechanical control, traps, conservation or augmentative biological control, and judicious use of insecticides. House fly is notorious for developing insecticide resistance and its resistance status is summarized as of August 2020. Several critical research needs are identified. Monitoring systems and nuisance/action thresholds need improvement. Faster-killing strains and better formulations are needed to integrate pathogens into Integrated Pest management (IPM) programs. The use of parasitoids remains an inexact science with many questions remaining about species selection and release rates. New attractants are needed for use in traps and attract-and-infect/kill strategies. Screening of new active ingredients for toxicity should continue, including a rigorous assessment of essential oils and other botanicals. Rising global temperatures may affect the balance of the fly with natural enemies. An understanding of the fly microbiome may reveal unknown vulnerabilities, and much remains to be learned about how flies acquire, retain, and transmit human and animal pathogens. System-specific research is also needed to tailor fly IPM programs to individual animal systems, especially in organic and free-range animal production.

Musca domestica, Housefly, filth flies

House flies have been pests of humans and animals since antiquity. Recent research by Gogarten et al. (2019) suggests that the relationship between humans and flies may predate recorded history for many millennia. These authors reported that muscid and calliphorid flies are closely associated with social groups of highly mobile nonhuman primates in the tropical forests of Ivory Coast. The flies appear to move with the primates and are rarely found outside of their immediate vicinity. Moreover, the flies carry and presumably spread pathogens that cause disease in the associated animals. The authors conclude that ‘attraction of flies might represent a previously underappreciated cost to forming social groups’. The association between humans and flies was undoubtedly strengthened once humans began to form more permanent settlements with domesticated animals and the concomitant manure accumulations. House flies were spread by humans as they radiated across the planet and are found on every continent except Antarctica. Their long synanthropy has allowed house flies to adapt to a wide variety of environmental conditions and food resources. This adaptability has resulted in the evolution of a formidable adversary that can avoid, adapt to, and evolve resistance to our best efforts to bring it under control. The objective of this article was to review the biology, pest status, and current management prospects for this important pest and to suggest areas where future research is needed.

Recognition and Distribution

Adult house flies have reddish eyes, sponging mouthparts, are 3–8 mm in length and can be recognized by the presence of four dark stripes on the dorsum of the thorax and the pronounced upward bend in the fourth longitudinal wing vein (vein M1+2) (Figs. 1 and 2). The basal portion of the abdomen is usually yellowish, especially along the sides, with males typically showing greater lateral yellowing than females. A dark longitudinal band runs along the median dorsal region of the anterior abdominal segments. The eyes of females are much more widely separated than those of males (Fig. 3). The key provided in Crosskey and Lane (1993) is useful for distinguishing Musca domestica from related muscoid flies. Eggs are deposited in batches in moist substrates, are creamy white, and approximately 1 mm long (Fig. 4). Mature house fly larvae can be recognized by the ‘D’-shaped peritreme surrounding sinusoidally shaped spiracular slits (Fig. 5). Pupae range in color from light red to dark brown, are 4–7 mm long, and have hardened spiracles with the same characteristics as the larvae (Fig. 6). House flies are found most commonly and abundantly at animal production facilities, but also occur in urban settings where the larvae develop in a wide range of decaying vegetable materials, feces, and household garbage. The common denominator among suitable developmental substrates is an abundant, viable microbial community.

Figure 1.

Female house fly. Arrows indicate distinct upward bend in the fourth longitudinal wing vein. Photo by Matt Aubuchon.

Figure 2.

Side view of female house fly head showing sponging mouthparts. Photo by Matt Aubuchon.

Figure 3.

House flies (5–8 mm) have a dull gray thorax with 4 thoracic stripes. The female (right) has widely separated eyes and a lightly golden checkered abdomen. The male’s eyes are positioned close to each other and has a darkly checkered abdomen. Photo by Matt Bertone.

Figure 4.

House fly eggs are small (~1 mm), white, and deposited on or just under the surface of moist substrates. Photo by Steve Denning.

Figure 5.

The creamy white third instar (5–8 mm) of the house fly with the blackened, internal mouth-hooks in the front and the distinctive D-shaped spiracles with sinusoidal slits at the rear. Graphic courtesy of Fallon Fowler using photos by Matt Bertone and Steve Denning.

Figure 6.

The reddish-brown puparium (4-7mm) of the house fly with hardened spiracles at the rear. Photo by Matt Bertone.

General Biology

House fly biology, life history, and management have been reviewed often and extensively. Early researchers developed a sound understanding of the fly’s biology over 100 yr ago (Howard 1911, Graham-Smith 1913, Hewitt 1914a), and the volume by West (1951) remains an essential work for its excellent anatomical illustrations and overview of critical life history events. Greenberg (1971, 1973) assembled the known literature on the association of flies and disease organisms into a 2-volume set. Since then, there have been several reviews with different emphases on house fly biology, physiology, management, or ecological associations (Keiding 1986, Axtell and Arends 1990, Axtell 1999, Hogsette and Farkas 2000, Malik et al. 2007). Published proceedings of two national workshops on research and extension needs for veterinary entomology included house fly information in chapters addressing fly concerns in different animal production systems (Anonymous 1976, Geden and Hogsette 2001). The following is intended as an overview of basic house fly life history.

Feeding

Adult house flies have sponging mouthparts and therefore must consume either liquid food or regurgitate crop contents onto solid foods to soften them prior to ingestion. ‘Bubbling’ is a common phenomenon, in which flies exude a droplet of regurgitate. Bubbling is thought to eliminate excess water (Hendrichs et al. 1992, Stoffolano 2019), thus concentrating the nutrients in the ingested food and reducing the weight that flies must carry during flight. Bubbling may also play a role in thermoregulation (Gomes et al. 2018). The droplets are sometimes deposited in the environment, leading to accumulations of both regurgitation and fecal spots on surfaces where flies rest (Fig. 7). The relative amount of regurgitation spots varies with the quality of the food ingested. Flies that have fed on nutrient-dense foods such as blood, liquid milk, and high-concentration sugar solutions produce fewer regurgitation spots than flies fed on items that are of lower nutrient density (CJG, unpublished data). Deposition of regurgitation and fecal spots play a role in the transmission of pathogens and can be used in monitoring programs as an index of fly abundance.

Figure 7.

Fly fecal and regurgitation spots accumulate where flies rest in the environment. Photos by Erika Machtinger.

The crop, a ventral diverticulated structure of the foregut, provides the fly with a highly expandable storage organ that allows it to consume large quantities of high-quality liquid food when such foods are encountered (reviewed in Stoffolano and Haselton 2013, Stoffolano 2019). Crop contents are then either passed to the midgut for digestion or passed forward for release from the proboscis as regurgitate. Food sometimes bypasses the crop and goes directly into the midgut, especially when ingested food is low in moisture content. As food enters the midgut it is surrounded by the peritrophic matrix (PM). The PM is synthesized by midgut cells and is composed of a matrix of chitin, glycoproteins, peritrophins, and other components that provide a barrier that prevents ingested pathogens from infecting the fly by denying them contact with the microvillar membrane of the midgut epithelium. Nutrients pass through the PM and are absorbed in the midgut, after which any unprocessed food enters the hindgut for water absorption and eventual deposition as feces. Passage of food through the entire digestive system can be completed in less than 6 h.

Reproduction

Female flies generally mate once, usually within 36 h of eclosion. A putative sex pheromone, (Z)-9-tricosene, was identified as a cuticular hydrocarbon constituent of female house flies (Carlson et al. 1971). This material is attractive to flies and is often incorporated into baits but is not present in all populations of flies collected in the field (Darbro et al. 2005, Butler et al. 2009). Amounts of (Z)-9-tricosene are higher in laboratory-reared flies than in wild flies (Noorman and Den Otter 2001, Darbro et al. 2005) and increase with fly age (Butler et al. 2013). Flies under laboratory conditions and given highly nutritious food are ready to deposit their first egg batch within 3–5 d after emergence and can lay 100–150 eggs per female per gonotrophic cycle. Because flies under such conditions can produce up to six batches of eggs, an individual female has the potential to lay 900 eggs in her lifetime (West 1951). Fletcher et al. (1990) observed that flies laid a maximum 709 eggs per female at 30°C under laboratory conditions with adequate food. Protein is required for egg maturation, but flies can develop eggs even when allowed brief, intermittent bouts of feeding on a protein source (Adams and Gerst 1991). Under field conditions, where fly longevity is shorter and food resources often limiting, rates of egg deposition are undoubtedly much lower. Lysyk (1991a) observed that flies given poultry feed and manure deposited only about 40 eggs per female over a 9- to 12-day lifespan, and Krafsur (1985) estimated that females on an Iowa dairy farm produced an average of 94 eggs over an approximate lifespan of 5.4 d. Oviposition occurs in groups and is stimulated by semiochemicals from the ovaries of gravid females (Jiang et al. 2002) and can be further modulated by bacterial symbionts deposited along with the eggs (Lam et al. 2007).

Longevity

Adult female flies held under optimal laboratory conditions can survive up to 6 wk (Fletcher et al. 1990, Haselton et al. 2004). Field estimates of fly longevity, based on mark-release-recapture methods, are much shorter than estimates from laboratory studies. The lifespans of wild flies have been estimated at 1–6 d and 3–19 d for flies on poultry (Lysyk and Axtell 1986c, Lysyk 1991a) and dairy farms (Krafsur et al. 1985, Kristiansen and Skovmand 1985, Butler et al. 2013), respectively, and 2–7 d for flies collected from a refuse dump (Imai 1984).

These results suggest that some production systems favor fly longevity over others. The short lifespans of flies reported from landfills and poultry farms may reflect poor food resources at such sites, whereas dairies offer a wider variety of high-nutrient foods such as milk, occasional blood from animal wounds, and pelleted feeds given to young animals.

Immature Development

Development time is determined by temperature and the quality of the larval substrate. Eggs generally hatch 6–12 h after deposition. Larsen and Thomsen (1940) found that development from egg to adult was completed in less than 7 d at 33°C. Lysyk and Axtell (1987) reported that the average time from oviposition to pupation ranged from 26.8 d at 16°C to 5.2 d at 35°C, and that the average time to adult emergence ranged from 43.1 to 8.8 d. Larvae move to dry areas to pupate and the duration of the pupal stage is 3–6 d under summer conditions. It is worth noting that development time experiments based on laboratory colonies may result in substantial underestimates. Early generations of new colonies from field collections have longer larval development times and higher pupal weights, and are much more variable in development times than colonies that have adapted to laboratory conditions (CJG, unpublished data).

The quality of the larval substrate also modulates development time. Larvae that are reared in suboptimal substrates often complete development but require longer larval stadia. Fly larvae reared in horse manure can take 50% longer to develop than larvae reared in poultry manure (Khan et al. 2012). Hogsette (1996) found that house fly larval development at 26.7°C required 22 d when they were reared in moist sand containing small amounts of dairy cattle manure. House fly larvae also can develop in a wide range of decaying plant substrates, including crop residues and culls after harvesting such as corn, carrots, onions, and snow peas (Cook et al. 2011). Regardless of the substrate, fly larvae must consume live microorganisms to complete their development (discussed below under fly-microbe associations).

Dispersal

On agricultural installations where house fly breeding typically occurs, most flies remain on or near the facility (Lysyk and Axtell 1986b). They are, however, capable of moving longer distances and can create problems when they disperse from the production site to residential areas, schools, and businesses (Thomas and Skoda 1993, Lole 2005, Winpisinger et al. 2005). Dispersal distances of >12 km have been reported (Bishopp and Laake 1921, Quarterman et al. 1954, Greenberg 1973), and Yates et al. (1952) collected a fly 32 km from the release point. Such reports of long-distance movements should be viewed with caution, however, since the vast majority of flies remain close to where they emerge. Dispersal distances vary depending on food availability and/or the number of development sites encountered by flies as they move through their environment (Schoof et al. 1952, Schoof and Siverly 1954a,b, Schoof 1959). Habitat structure can provide corridors for longer distance travel (Fried et al. 2005). In summary, most house fly movement from agricultural facilities is likely a series of short disjointed circuitous flights culminating in the aggregation of house flies at some more distant attractive site (Schoof and Siverly 1954a,b) rather than a directional dispersal to discover and colonize new habitats. It is notoriously difficult to determine the source populations of flies when problems and complaints arise on nonfarm sites, often resulting in disputes between animal operations, neighbors, and local health officials.

Genetics

The house fly has five pairs of autosomes plus an X and Y chromosome. Sex is determined by the Mdmd gene (Sharma et al. 2017), which can be found on any of the chromosomes and can be present in more than one copy (Hamm et al. 2015, Son et al. 2019). Y is the presumed ancestral chromosome for Mdmd. The house fly genome was sequenced in 2014 (Scott et al. 2014) and revealed an abundance of recognition and effector components of the immune system, consistent with its close association to pathogens. Initial studies found a lack of crossing-over in male house flies (McDonald 1971), consistent with what is observed in most Diptera (Gethmann 1988). Subsequently, it was found that crossover frequencies in males vary, depending on the genes examined and the populations used. Reported values range between 0–0.53% (Hamm et al. 2005, Hamm and Scott 2008), 0.03–0.11% (Sullivan 1961), 9.3–31% (Lester et al. 1979), and 7–28% (Feldmeyer et al. 2010). Greater male recombination rates tend to be associated with males that have Mdmd on an autosome.

Fly–Microbe Interactions

Larval Associations

Larvae feed on live microorganisms within the substrate. Gerberich (1948) and Greenberg (1954) first noted that house fly larvae must ingest living microorganisms to complete development and suggested that this was due to B vitamins or other nutritional factors provided by gut-inhabiting microbes. When grain-based fly larval diets were first developed, it was recommended that they be inoculated with a cocktail of Escherichia coli, Sarcina spp., and Lactobacillus spp. to promote larval development (Spiller 1964). Some early studies suggested that house fly larvae could be grown under axenic conditions (Brookes and Frankel 1958, Monroe 1962), but the methods may not have been sufficiently sterile to rule out colonization by microbes. House fly larvae grown on blood agar without bacteria do not develop past the first instar, but can complete development on monoxenic plates with E. coli, Klebsiella pneumoniae, Staphylococcus spp., and Streptococcus bovis (Schmidtmann and Martin 1992). Similarly, Watson et al. (1993) successfully grew house fly larvae on sterile egg yolk and blood agar media inoculated with E. coli, whereas almost no adult flies were produced on sterile media. Bacteria isolated from the gut of fly larvae in soiled turkey litter, including the Gram-positive facultative anaerobic species Staphylococcus lentus, Streptococcus sanguinis, Lactococcus garviae, Yersinia pseudotuberculosis, and Bacillus coagulans all supported house fly development in monoxenic trypticase soy egg yolk agar; larval development was poor when the bacteria were ones recovered from larvae found in fermented corn silage (Zurek et al. 2000). The authors concluded that ‘house fly larvae likely benefit from complex metabolic interactions within a diverse bacterial community in a natural environment leading to a rapid degradation of organic material as well as a great build-up of bacterial mass’. The house fly larval microbial community is complex and variable, and horizontal transmission of microbes among larvae in feeding sites ensures a continuous local supply of organisms (Zhao et al. 2017).

Acquisition, Retention, and Transmission of Microbes by Adult Flies

Adult house flies routinely acquire microbes such as protists, viruses and bacteria during their persistent associations with microbe-rich environments. Over 200 different species of microbes have been isolated from wild-caught house flies (Nayduch and Burrus 2017), and a single house fly can carry up to 100 different pathogenic microbes (Greenberg 1973). Flies acquire microbes on their surfaces via contact with or by directly feeding upon refuse, animal waste, wounds and exudate (West 1951, Nayduch and Burrus 2017). Pathogens on flies’ surfaces are transferred to animals by physical contact or by being dislodged by fly grooming (Yap et al. 2008, Jacques et al. 2017). Some bacteria such as E. coli O157:H7 survive and proliferate for several days on mouthparts before transmission (Kobayashi et al. 1999).

When flies ingest bacteria, the location and persistence of the bacteria within the digestive tract impacts transmission potential. Ingested bacteria harbored in the crop and midgut are either digested and destroyed or survive to be excreted and, possibly, transmitted (Nayduch and Burrus 2017, Stoffolano 2019). Bacterial persistence, propagation, excretion, and transmission can vary by pathogen type (Nayduch et al. 2002, Joyner et al. 2013, Nayduch et al. 2013, Fleming et al. 2014). Even fly sex can impact the acquisition, persistence, proliferation, and excretion of pathogens (Thomson et al. 2017, Nayduch et al. 2018).

Transmission of pathogens by flies to human food items is of growing concern in light of several high-profile recalls of contaminated leafy greens (Talley et al. 2009). Transmission of disease-causing E. coli from house flies to spinach leaves was shown by Wasala et al. (2013). Viable cells of E. coli O157:H7 were detected 8 d after contaminated flies contacted the spinach. Because E. coli cells cannot survive 8 d, the flies must have transmitted viable cells (Wasala et al. 2013). Transmission by house flies has been shown for a number of bacterial pathogens under a variety of test conditions, including contamination of the environment and human food items (Tables 1 and 2).

Table 1.

Studies demonstrating transmission of pathogenic bacteria by house flies

Pathogen	Extent of dissemination	References
Aeromonas caviae	Contaminate environment	Nayduch et al. 2002
Aeromonas hydrophila	Viable in excreta	McGaughey and Nayduch 2009
Campylobacter jejuni	Viable in excreta	Gill et al. 2017
	Contaminate environment	Shane et al. 1985
Corynebacterium pseudotuberculosis	Viable in excreta	Braverman et al. 1999
Enterococcus faecalis	Contaminate environment	Doud and Zurek 2012
Escherichia coli O157:H7	Viable in excreta	Sasaki et al. 2000, Fleming et al. 2014
	Contaminate environment	Wasala et al. 2013
Pseudomonas aeruginosa	Viable in excreta	Joyner et al. 2013
Salmonella Typhimurium	Viable in excreta	Chifanzwa and Nayduch 2018
Salmonella Schottmullerris	Viable in excreta	Hawley et al. 1951
Salmonella Enteritidis	Transmit to hens	Holt et al. 2007
Shigella dysenteriae	Viable in excreta	Hawley et al. 1951
Staphylococcus aureus	Viable in excreta	Nayduch et al. 2013
Yersinia pseudotuberculosis	Contaminate environment	Zurek et al. 2001

Table 2.

Food-borne pathogens transmitted to or from food by house flies

Pathogen	Food type	References
Aeromonas caviae	Various	Nayduch et al. 2002
enterococci	Cooked hamburger patty	Macovei et al. 2008
Escherichia coli O157:H7	Spinach	Wasala et al. 2013
	Lettuce	Pace et al. 2017
Antimicrobial-resistant E. coli	Various	Fukuda et al. 2019
Salmonella enteric	Mexican drink	Greenberg 1964
	Lettuce	Pace et al. 2017
	Canteloupe	Thomson et al. 2020

Only includes studies that have specifically described food-borne pathogens and food sources and demonstrated house fly impact on food safety.

Flies potentially facilitate the evolution and dispersal of new pathogenic and resistant microbial strains (Petridis et al. 2006, Onwugamba et al. 2018). For example, lateral gene transfer between bacteria may occur in the fly gut (midgut + crop), including the horizontal movement of resistance alleles among pathogens and nonpathogens (Akhtar et al. 2009, Zurek and Ghosh 2014, Onwugamba et al. 2018, Poudel et al. 2020). The horizontal transfer of a plasmid harboring cephalosporin and tetracycline resistances between donor and recipient bacteria in the house fly gut has been demonstrated (Fukuda et al. 2016). The plasmid was not only transferred between donor and recipient E. coli strains, but also to other bacterial taxa that were present as ‘normal flora’ (Achromobacter spp. and Pseudomonas fluorescens). The horizontal transfer of plasmids carrying tetracycline resistance among enterococci has also been demonstrated within the house fly gut (Akhtar et al. 2009). Furthermore, flies may facilitate the introduction and dispersal of genes affecting virulence (Zurek and Ghosh 2014). Because adult flies are highly mobile, they substantially contribute to the dissemination of these novel pathogens across habitats and ecological niches (Zurek and Ghosh 2014, Onwugamba et al. 2018, Poudel et al. 2019, Sobur et al. 2019).

Dissemination of Pathogens by Adult Flies

Adult flies bridge unsanitary and sanitary environments, disseminating bacteria from their source to animal facilities, food, water, and nearby humans (Nayduch and Burrus 2017) (Fig. 8). Fly populations flourish in livestock facilities and the threat to human and livestock health is intensified when poor sanitation and insufficient manure management allows flies to have unrestricted access to pathogen sources such as waste and excrement. House fly dispersal between farms and nearby residential and urban centers facilitates bacterial transmission to humans and therefore poses a public health risk. As discussed previously, flies can disperse several kilometers from their larval habitats (e.g., farms). This increases the potential to spread manure-acquired bacteria, including antimicrobial resistant (AMR) strains, to surrounding locales (Alam and Zurek 2004, Winpisinger et al. 2005, Burrus 2010, Baldacchino et al. 2018, Neupane et al. 2020). Chakrabarti et al. (2010) estimated that flies could facilitate movement of antibiotic-resistant bacteria as far as 100 km from a cattle feedlot. Flies may spread AMR E. coli among animal production sites that are geographically separated (Usui et al. 2015), which may contribute to multiple AMR E. coli strains occurring within single facilities (Sola-Gines et al. 2015). Other specific examples of fly transmission include dispersal of methicillin-resistant Staphylococcus aureus to urban communities (Schaumburg et al. 2016), transmission of AMR serotypes of various Salmonella spp. within and among swine farms (Wang et al. 2011), transmission of Corynebacterium pseudotuberculosis, a causative agent of bovine mastitis (Yeruham et al. 1996), and possibly dissemination of Campylobacter (Hald et al. 2007) or Salmonella spp. (Olsen and Hammack 2000) at poultry facilities.

Figure 8.

Flies aid in transmission of animal pathogens when they visit animal manure and food resources. Photos by Chris Geden.

House Fly Impacts in Human and Animal Systems

Economic Losses

It is difficult to determine the present economic scope of the house fly problem. After adjusting for inflation, a 2001 estimate for the annual cost of insecticides for fly control in the poultry industry is about $30 million today (Geden et al. 2001). Estimates for the dairy and swine industries, after adjusting for inflation, are $135 million and $35 million, respectively (USDA 1978, Campbell 1993). Taken together, the cost of insecticide use for house flies in these three commodities can therefore be estimated at about $200 million. A 1976 estimate for the total cost of house fly damage and control is approximately $450 million today (Anonymous 1976).

Even after adjusting for inflation, these figures are undoubtedly low. One reason for this is the high cost of legal settlements against producers in recent decades. Changing demographics have resulted in growing friction between once-isolated agriculture facilities and residential developments. House flies were one of the complaints in the recent $50,000,000 million settlement against Smithfield Foods for nuisance complaints regarding hog farms (Brown 2018). Societal and public health pressures make it necessary to keep fly population levels lower than were acceptable when flies were seen as being a natural outcome of living in rural settings. Moreover, the relatively higher cost of modern insecticides has made fly control more expensive than in past years. Until more solid economic data become available, we estimate that house flies account for $500 million to $1 billion per year in insecticide costs and total economic losses, respectively.

Human Health Concerns

Due to their persistent associations with decomposing substrates, flies carry a rich and diverse bacterial community. House flies are known to carry an ever-growing list of human pathogens that can be transmitted directly to people or indirectly via contamination of food (Sundin 1996, Graczyk et al. 2001, Boulesteix et al. 2005, Rahuma et al. 2005, Macovei and Zurek 2006, Macovei et al. 2008, Graham et al. 2009, Nayduch and Burrus 2017, Khamesipour et al. 2018, Xu et al. 2018). Infected flies often originate from concentrated livestock operations, which generate large numbers of house flies that have access to potential zoonotic and AMR pathogens shed in animal manure. Many different pathogenic bacteria have been isolated from livestock-associated flies, including AMR strains and both human and livestock pathogens (Table 3). Other pathogens such as viruses, protists, and helminth eggs are also associated with wild house flies, albeit only transiently and in less abundance; they are reviewed elsewhere (e.g., Graczyk et al. 2001, Nayduch and Burrus 2017).

Table 3.

Animal and human pathogens found in flies collected at different animal systems

Bacteria	Animal system							References
	Dairy cattle	Beef cattle	Cattle (unspecified)	Swine	Poultry	Equine	Other
Acinetobacter spp.					✓			Nazni et al. 2005
Acinetobacter baumannii					✓			Nazni et al. 2005
Aeromonas spp.a	✓		✓	✓	✓			Nayduch et al. 2001; Ommi et al. 2015a
Aeromonas caviae	✓			✓				Nayduch et al. 2001
Bacillus spp.	✓				✓			Nazni et al. 2005; Bahrndorff et al. 2017
Campylobacter spp.a	✓		✓		✓			Szalanski et al. 2004; Brazil et al. 2007; Choo et al. 2011; Royden et al. 2016; Ommi et al. 2016a
Campylobacter coli				✓	✓			Rosef and Kapperud 1983; Hald et al. 2008
Campylobacter fetus subsp. jejuni				✓	✓			Rosef and Kapperud 1983
Campylobacter jejuni				✓	✓			Hald et al. 2008; Förster et al. 2009a,b
Citrobacter spp.a			✓		✓	✓	✓	Moissant et al. 2004, Neupane et al. 2020a
Clostridium spp.	✓							Bahrndorff et al. 2017
C. perfringens					✓			Dhillon et al. 2004
Coccobacillus spp.					✓			Nazni et al. 2005
Corynebacterium spp.	✓							Hernandez 2012
Corynebacterium pseudotuberculosis	✓							Braverman et al. 1999
Edwardsiella spp.					✓			Shukla et al. 2013
Enterobacter spp.			✓		✓	✓	✓	Moissant et al. 2004; Nazni et al. 2005; Neupane et al. 2020a
Enterobacter sakazakiia	✓							Buma et al. 1999a
Enterococcus spp.					✓			Graham et al. 2009a
Enterococcus casseliflavusa				✓				Ahmad et al. 2011a
Enterococcus faecalisa				✓	✓			Ahmad et al. 2011a
Enterococcus faeciuma				✓	✓			Ahmad et al. 2011a
Enterococcus hirae				✓				Ahmad et al. 2011
Escherichia spp.	✓			✓	✓			Nazni et al. 2005; Nmorsi et al. 2007; Förster et al. 2009a,b
Escherichia colia	✓	✓	✓	✓	✓	✓	✓	Echeverria et al. 1983; Oo et al. 1989; Iwasa et al. 1999; Agui 2001; Alam and Zurek 2004; Moissant et al. 2004; Szalanski et al. 2004; Brazil et al. 2007; Nmorsi et al. 2007; Literak et al. 2009,a; Usui et al. 2013,a; Usui et al. 2015,a; Blaak et al. 2014,a; Solà-Ginés et al. 2015,a; Schaumberg 2016,a; Bahrndorff et al. 2017; Puri-Guri et al. 2017; Neupane et al. 2020a
E. coli 0157:H7 (EHEC)a	✓		✓	✓	✓			Buma et al. 1999,a; Iwasa et al. 1999; Szalanski et al. 2004; Burrus 2010; Förster et al. 2009a,b; Burrus et al. 2017
Histophilus somni		✓						Neupane et al. 2019
Klebsiella spp.a	✓				✓			Nazni et al. 2005; Nmorsi et al. 2007; Neupane et al. 2020a
Klebsiella pneumoniaea	✓		✓		✓			Nmorsi et al. 2007; Ranjbar et al. 2016,a; Neupane et al. 2020a
Lactobacillus spp.	✓							Hernandez 2012
Listeria spp.			✓		✓	✓	✓	Hernandez 2012
Mannheimia haemolytica		✓						Neupane et al. 2019
Micrococcus spp.					✓			Nazni et al. 2005
Pasteurella multocida		✓						Neupane et al. 2019
Proteus spp.a	✓				✓			Nazni et al. 2005; Nmorsi et al. 2007; Neupane et al. 2020a
Proteus mirabilis	✓							Nmorsi et al. 2007
Providencia spp.a					✓			Shukla et al. 2013; Neupane et al. 2020a
Pseudomonas spp.a			✓		✓			Hemmatinezhad et al. 2015a
Pseudomonas fluorescensa	✓							Buma et al. 1999a
Salmonella spp.a	✓	✓	✓	✓	✓			Oo et al. 1989; Olsen and Hammack 2000; Nmorsi et al. 2007; Choo et al. 2011; Wang et al. 2011,a; Xu et al. 2018a
Salmonella Heidelberg					✓			Olsen and Hammack 2000
Salmonella infantis					✓			Olsen and Hammack 2000
Salmonella typhi	✓							Nmorsi et al. 2007
Salmonella typhimurium	✓							Nmorsi et al. 2007
Serratia spp.a	✓							Nmorsi et al. 2007; Neupane et al. 2020a
Serratia marcescensa	✓							Buma et al. 1999,a; Neupane et al. 2020a
Shigella spp.			✓	✓	✓		✓	Echeverria et al. 1983; Oo et al. 1989; Shukla et al. 2013
Staphylococcus spp.a	✓		✓		✓	✓	✓	Nazni et al. 2005; Nmorsi et al. 2007; Graham et al. 2009,a; Hernandez 2012; Bahrndorff et al. 2017
Staphylococcus aureusa	✓		✓	✓				Nmorsi et al. 2007; Schaumberg et al. 2016a
Stenotrophomonas maltophilaa				✓	✓			Fukuda et al. 2017a
Streptococcus spp.	✓				✓			Nazni et al. 2005; Nmorsi et al. 2007
Streptococcus faecalis	✓							Nmorsi et al. 2007
Streptococcus pyogenes	✓							Nmorsi et al. 2007
Vibrio spp.			✓	✓	✓	✓	✓	Echeverria et al. 1983; Hernandez 2012
Vibrio cholera			✓	✓	✓			Oo et al. 1989

^aAntimicrobial resistance found in bacterial isolates from house flies.

Flies pose a risk to humans living near sources of human pathogenic microbes including livestock and poultry operations, landfills, and wastewater management facilities (Iwasa et al. 1999, Zurek and Ghosh 2014, Schaumberg et al. 2016). Several studies have suggested flies as a source of human pathogenic bacteria in animal operations, including E. coli O157:H7 (Rahn et al. 1997, Hancock et al. 1998, Iwasa et al. 1999, Alam and Zurek 2004, Szalanski et al. 2004), Camplylobacter spp. (Szalanski et al. 2004, Ekdahl et al. 2005), and Salmonella spp. (Mian et al. 2002). A recent survey of house flies from dairies and feedlots in Georgia identified Salmonella spp., including strains with AMR, in 11% (185/1650) of flies and pathogen incidence varied widely across sites (0–78%) (Xu et al. 2018). Single-drug AMR also was reported in some of these isolates, and 28% of the Salmonella isolates were multidrug resistant. Flies carrying enteropathogenic E. coli which originated from a nearby animal farm were posited as the source of a colitis outbreak at a children’s school in Japan (Moriya et al. 1999). House flies collected from wastewater management facilities carried antibiotic resistant Enterococcus faecalis and a few flies were found to carry these resistant bacteria in a recreational vehicle park, a fast food restaurant, and an apartment complex close to some of the treatment facilities (Doud et al. 2014).

Flies also pose a risk when they have access to human garbage and feces in situations where basic sanitation practices are poor. House flies were connected to outbreaks of typhoid fever from Salmonella typhi during the Spanish-American war in military camps (Cirillo 2006) and a dysentery outbreak in a U.S. army camp (Kuhns and Anderson 1944). The proximity of food and water sources to feces in addition to the presence of higher than average fly densities contributed to a 15% increase in the risk of diarrhea in humans in parts of India (Collinet-Adler et al. 2015). In Bangladesh, flies were implicated in contaminating food with enteropathogenic E. coli that originated from nearby human feces (Doza et al. 2018). The bazaar fly, Musca sorbens Wiedemann, a close relative of house fly with a strong affinity for human eye secretions, is an important vector of the causative agent of trachoma (Chlamydia trachomatis) in many developing countries (Emerson et al. 2000); M. domestica appears to be a competent vector as well (Stoffolano, personal communication). Although there is no current direct evidence of house flies being able to transmit the Ebola or SARS-CoV-2 viruses, fly control is recommended as a sanitation precaution against mechanical transmission (Haddow et al. 2017, Dehghani and Kassiri 2020).

One way to demonstrate the role of house flies in transmitting pathogens is by showing that lack of fly control is linked to diarrheal disease prevalence. Shortly after the discovery of DDT, a study in Texas found that the use of this insecticide for fly control reduced the prevalence of diarrheal diseases in children under the age of 5 (Watt and Lindsay 1948). The prevalence of Shigella infections was significantly lower only during the time when an effective fly control program was carried out in select towns of rural Georgia (Lindsay et al. 1953). McCabe and Haines (1957) showed that over the 18 yr after reconstructing outhouses in Boston to reduce house fly breeding in human feces, the frequency of Shigella infections in children decreased and the diarrheal disease rate for Boston was cut in half. In Bangladesh, a surge of house fly populations in the spring was correlated with the subsequent increase in Shigella diarrhea among children two months later (Farag et al. 2013). Similarly, the incidence of diarrheal disease was lower in the children from towns in Pakistan where insecticides were used to control flies (Chavasse et al. 1999). Fly control at an Israeli military base resulted in an 85% drop in clinic visits for shigellosis (Cohen et al. 1991).

Animal Health Concerns

Filth flies, including house flies, are key players in food security (e.g., animal health) and both pre- and postharvest food safety across a variety of livestock commodities (Mian et al. 2002, Dhillon et al. 2004, Winpisinger et al. 2005, Holt et al. 2007, Hald et al. 2004). The list of pathogenic microbes found in association with flies, especially those associated with livestock, is vast and continues to grow (Keiding 1986, Scott et al. 2014, Nayduch and Burrus 2017, Khamesipour et al. 2018). House flies are mechanical vectors of scores of pathogens responsible for animal diseases. Many of the bacterial species listed in Table 3 are animal pathogens.

Livestock diseases can have differing impacts depending on typology and severity. Diseases impacting the food supply can have profound effects on the entire livestock industry as outbreaks of some diseases can lead to dramatic supply reductions, partial, or full stoppages of varying duration with trading partners, as well as a hesitation to consume meat products from regions that are found to have livestock disease outbreaks. A less severe scenario includes a spread of disease throughout the herd, causing diminished productivity or an increase in mortality and morbidity. Commodity specific pathogens are discussed below.

Challenges for Different Animal Systems

Animal production systems are continuing a trend of greater intensification to maximize production while minimizing costs that started in the previous century (Machtinger et al. 2020). One of the hallmarks of this intensification is that increased animal density results in greater production of animal waste and a greater need for (and often storage of) feed per unit of area (Gerry 2018, Clay et al. 2020)). The abundance of suitable fly larval development habitat, particularly animal manure, is the most important determinant of house fly presence and populations sizes in such systems. Manure is typically collected and stored at the facility, providing an uninterrupted input of fly-development substrates into the system. The mixture of manure with animal feed and bedding is another feature of intensive confined animal production, and these mixtures provide ideal fly development sites. Control of house flies under these conditions is difficult as it requires substantial effort to manage manure and other developmental substrates. Swine, poultry, dairy cattle, beef feedlots, and equine systems have many elements in common, but each poses particular challenges. Some of these are outlined below, as well as examples of disease concerns in each system where house flies play a documented or suspected role.

Swine

Swine production has moved toward fewer farm numbers and increased concentrations of animals over the past few decades (USDA 2017) (Fig. 9). These large numbers of swine produce waste and decaying organic matter that may serve as development sites for house flies. While understanding production and legal risks associated with house flies has increased over the past decades, information on the muscoid fauna in swine facilities in the United States is limited. Burns and Nipper (1960) reported that house flies were the most important insect problem associated with pig parlors in Louisiana, whereas house flies accounted for only 11.8% of filth flies in confined hog facilities in Texas (Robertson and Sanders 1979).

Figure 9.

Swine in the U.S. are usually held in high-density enclosed facilities. Manure is collected into liquid storage systems of various types, but barn design sometimes leaves corners and other places that are difficult to clean. Source: USDA NCRS, photo by Bob Nichols.

Porcine reproductive and respiratory syndrome (PRRS) virus, one of the most economically significant pathogens in the swine industry (Holck and Polson 2003), can be transmitted by house flies (Otake et al. 2003). PRRS is estimated to cost the U.S. swine industry approximately $560 million per year (Neumann et al. 2005). House flies transmit PRRS among animals within a facility and can move PRRS from one facility to another (Schurrer et al. 2004, Pitkin et al. 2008). House flies may also play a role in between-farm movement of porcine epidemic diarrhea virus (PEDV) (Masiuk et al. 2018, Allison et al. 2019, 2020). PEDV first emerged in the United States in 2013, and by May 2014 it had been found in 29 states (Schulz and Tonsor 2015). PEDV is most serious in neonatal piglets where morbidity and mortality can be 80 to 100%, with mortality increasing with age. House flies have also been implicated in the transmission of hog cholera (Dorset et al.1919), transmissible gastroenteritis virus TGEV (Saif and Wesley 1999), and bacterial pathogens including Lawsonia intracellularis (Dee et al. 2004, Förster et al. 2007, McOrist et al. 2011), Streptococcus suis (Enright et al. 1987, Staats et al. 1997), and Mycobacterium (Fischer et al. 2001). House flies can also transmit E. coli among swine causing neonatal and post-weaning diarrhea, which are important causes of death in suckling and weaned pigs respectively (Fairbrother and Gyles 2012). Salmonella spp. can be transmitted by house flies in swine (Wang et al. 2011) causing salmonellosis, one of the top 10 most common diseases in weaning and grower/finisher pigs, costing pork producers an estimated $100 million annually (Knetter et al. 2015).

While direct economic losses in swine resulting from house fly infestations have not been documented, it has been estimated that over $20 million is spent annually on house fly control by pork producers in the North Central states (Campbell 1993). High numbers of house flies developing in swine facilities may subject producers to nuisance litigation. Legal cases citing fly nuisance have increased in recent years as residential homes have increased in historically rural areas. In 2019, over $470 million was awarded to plaintiffs in North Carolina citing, in part, high fly numbers originating at local hog facilities (Wall Street Journal 2018).

Sanitation practices to eliminate or minimize fly breeding materials is the most important and effective approach to house fly management in and around swine facilities. While there are several management strategies for waste in hog facilities, housing on concrete slatted floors over a slurry pit is common (USDA 2017) (Fig. 8). Alternatively, under-floor, sloping drainage channels take liquid waste to large liquid lagoons several times a day. Generally, the high moisture content of waste from these management techniques prohibit house fly development. However, both the concrete pens and waste channels are difficult to clean thoroughly and, if not well designed, can leave areas for house fly development. House flies are managed in swine facilities using methods developed for dairy, beef cattle, and poultry facilities; the efficacy of these methods in swine systems is largely unknown

Poultry

From the standpoint of fly production, poultry facilities can be divided into those where: 1) birds are permitted to contact and forage through their own feces; and 2) birds are separated above feces which accumulates undisturbed beneath caged birds or birds held on slatted floors. Turkeys and broiler chickens are generally housed in wide single-story buildings (‘grow-out houses’) where birds can move freely across a floor covered with wood shavings or other litter material (Axtell 1986). Fly production in such facilities is low because birds scratch through the soiled litter, disturbing and drying feces, and consuming fly immatures and other insects. In contrast, layer hens are typically held suspended above the ground in wire cages or aviaries, with bird feces and spilled feed accumulating in piles or rows beneath the birds and often resulting in much greater fly densities relative to other housing designs (Axtell 1986) (Fig. 10). Even cage-free facilities typical for breeder birds can allow for manure to accumulate in nest boxes and under floor slats where the manure may remain undisturbed for long periods which can promote fly development. Typically, house flies are the most abundant pest fly in layer poultry facilities. (Lysyk and Axtell 1986a, Stafford et al. 1988, Hogsette 1993).

Figure 10.

Manure in high-rise caged layer houses either collects below the birds or is removed by automated belt systems to liquid storage or removal off-site. Large fans for circulation also aid in manure drying. Photos by Erika Machtinger.

High fly populations are a risk to confined poultry, primarily due to their potential to transmit pathogens among birds within a confined setting (Shane et al. 1985). Necrotic enteritis (NE) is a disease found worldwide wherever chickens are farmed (McDevitt et al. 2006). This economically significant disease, caused by the bacterium Clostridium perfringens, results in lesions in the chicken’s intestine and can lead to flock mortality of 1% per day (clinical NE). The estimated cost of NE is $2.5 billion per year in the United States (Wade and Keyburn 2015). Dhillon et al. (2004) found C. perfringens in house flies on a poultry farm with both high fly populations and NE disease incidence. Avian pathogenic strains of E. coli are also a concern in laying hens, where they cause avian colibacillosis, salpingitis/peritonitis/salpingoperitonitis (SPS) and E. coli peritonitis syndrome (EPS). EPS is estimated to kill about 6% of the hens at egg facilities annually, with an annual cost to the facility of about $1.15 M (Zoetis 2018). Salmonella spp. infections are common in poultry and result in acute and chronic diseases that consume large economic investments for monitoring and control (Gast and Porter 2020). Flies not only disseminate Salmonella among animals, but also can serve as a reservoir for the bacteria in the environment (Mian et al. 2002). Naive chickens became infected with Salmonella enteritis serovar Enteritidis when they ingested flies collected from a facility with infected hens (Holt et al. 2007). House flies may also serve as mechanical vectors of viral pathogens such as turkey coronavirus (Calibeo-Hayes et al. 2003), exotic Newcastle disease (Chakrabarti et al. 2007, Chakrabarti et al. 2008), and avian influenza (Habibi et al. 2018).

Poultry husbandry practices in intensive production systems maximize egg production at lowest cost, but these practices can create ideal conditions for house fly development as a result of very high bird densities and rapid accumulation of bird feces leaving too little time for feces to dry to prevent fly development. Conventional caged layer farms account for over 54% of layer poultry facilities in the United States with the majority of these (93.5%) being farms with over 100,000 birds (USDA 2014). Manure waste is produced at 113 g per bird per day (North and Bell 1990), which translate to 4,129 metric tons per year for a 100,000-bird house. In many poultry facilities, birds are housed in an artificial environment with temperature and humidity managed in a narrow range that is quite suitable for fly production.

There has been an increase in high-rise poultry housing from 39.7% of facilities in 1999 to 61.7% in 2013. Modern caged-layer facilities often have automated belt systems that remove manure daily, where it can be either placed in storage areas or taken off-site, but such systems comprise less than 20% of facilities (USDA 2014). Composting of stockpiled manure prevents larval development and has the additional benefit of killing pathogens (Macklin et al. 2008). Only 16.5% of farms store manure outside (USDA 2014).

Control of house flies is best achieved in intensive poultry systems by increased efforts toward management of feces and litter to reduce their availability for fly production or their suitability for fly development, primarily by using strict moisture management. When fly outbreaks occur, most egg producers use baits and traps (77.8%) and residual sprays for fly management (59.4%) (USDA 2014). Over 30% use larvicides and 49.6% use space sprays or foggers. The number of farms using biological control agents (mostly parasitoids) increased from 14% in 1999 to 31% in 2013. This increase may be due to increased organic production, which now represents 27.6% of layer production in the United States (USDA 2014).

Dairy

Milk production since the 1980s has increased more than 59% worldwide to 843 million tons in 2018 (FAO 2020). In developed countries, dairy farms are growing larger (USDA 2018) and are increasingly mechanized with cow management and feed carefully controlled to increase milk production per animal. As animal density increases, so do quantities of house fly developmental substrates.

House flies can transmit pathogens that are of concern to dairy production, typically through mechanical methods. Although the house fly has been implicated in transmission or transportation of microbes that cause human illness, many of these organisms are not pathogenic in cattle. Salmonella spp. cause substantial problems for dairy health. Salmonellosis in cattle most commonly affects colostrum and deficient calves can exhibit fever, diarrhea, rapid dehydration and death within 24–48 h (Wray and Wray 2000). Both clinical outbreaks and subclinical infections of Salmonella can drain profit from the dairy operation by contributing to declines in milk production, abortions, losses from antibiotic contaminated milk, increased culling, increased labor for management of sick animals, reduced feed efficiency, the inability to sell animals originating from an ‘infected’ herds, and death (Holschbach and Peek 2018). Aeromonas bacteria are associated with diarrheal diseases in livestock and humans. The role of the house flies is unclear. Flies can harbor several Aeromonas species, but transmission studies are lacking (Nayduch et al. 2001). Within-herd mechanical transmission of Corynebacterium pseudotuberculosis by house flies is suspected in Israeli dairies (Yeruham et al. 1996, Braverman et al. 1999). Yeruham et al. (2003) reported morbidity and animal culling exceeding 6 and 16%, respectively, and milk loss of greater than 6% in heavily affected herds. House flies have been implicated in transmission of Cryptosporidium parvum and Giardia lamblia, both of which cause zoonotic diarrheal diseases in humans and livestock and have been recovered from house flies on dairy farms (Doiz et al. 2000, Clavel et al. 2002). House flies collected from dairies can be infected with Klebsiella pneumoniae, an important cause of mastitis, however, animal management practices are considered the primary driver of disease incidence (Nmorsi et al. 2007). Staphylococcus aureus, a cause of mastitis in dairy cattle, has been recovered from house flies, but flies are not thought to be a major vector of these bacteria. Recently house flies collected from a dairy facility have been shown to carry antimicrobial- and multi-drug-resistant coliforms, indicating that they are not only a source of pathogenic bacteria but also harbor commensals and their resistance genes (Neupane et al. 2020).

Housing of dairy cattle can be varied. As of 2014, conventional dairies made up 58.8% of farms in the United States and conventional with grazing access farms were 26.5% of all dairies. Organic dairy farming made up 74% of farms. Most farms either used tie stall or stanchion (38.9%) or free stall without outdoor access (20%) as the primary housing method (USDA 2016). Calves housed on bedding are important sources of fly production. The use of individual hutches to house calves is helpful for disease mitigation, but creates substantial fly problems if hutches are not moved frequently and provisioned with fresh bedding (Fig. 11). Bedding choice is typically straw or hay (47.1%) or sawdust (34.1%), which are very supportive of fly larval development (Schmidtmann 1991).

Figure 11.

Calf housing is often a major contributor to fly production on dairy operations. Photo on left by Chris Geden; photo on right from USDA NCRS by Scott Bauer.

In intensive dairy production systems, substantial quantities of cattle feces collected from animal housing areas are often stored on-site. Flushing and scraping often are used to collect manure from lactating cow housing areas. Although practices may vary by region, most dairies surveyed in California use mechanical or gravity separation to sort solid manure from liquid manure (Meyer et al. 2011). Similar to swine, recycled water from liquid manure storage ponds or tanks is used to flush manure, while solids are most frequently piled and remain uncovered (80.1%) or composted (26.3%) (Meyer et al. 2011). Both solid and liquid manure are often land-applied at some point after removal.

Intensive systems must keep substantial quantities of animal feed on site, including hay, straw, grains, and fermenting feed additives (Gerry 2018). Including fruit and nut waste in feed is common, much of which is fermented either deliberately or due to placement of dry feed in a location where it is wetted by rainfall, sprinklers, or runoff from pens. Fermenting feed stocks can provide suitable development substrates for house flies (Fig 12).

Figure 12.

Stored animal feed can provide fly development sites when it becomes wet. Photo by Chris Geden.

Cattle Feedlots

Beef production and consumption worldwide is increasing slowly with beef consumption increasing primarily in developing countries (USDA 2020). Beef production is limited by declining rangeland availability in most countries due to encroachment of other land uses and degradation of available rangelands. Further increases in beef production are likely to result from increasing animal density on available lands with animals provided supplemental feeds where forage is no longer sufficient (Bruinsma 2003). Modern cattle feedlots, where cattle do not have access to pasture and are fed entirely on supplemental feed, are an extreme example of beef cattle intensification (Fig. 13). The size and animal density on U.S. feedlots continues to grow, with 40% of fed beef cattle produced on facilities with over 32,000 animals (Economics Research Service 2020). In April 2020, there were approximately 12 million cattle in U.S. feedlots (National Agriculture Statistics Service 2020), mostly housed at densities of 2–46 m² of pen space per animal (Ingvartsen and Andersen 2009, Euken et al. 2015).

Figure 13.

Many modern cattle feedlots pose particular problems for fly management because of their large size and high animal densities. Source USDA NCRS, photo by Jeff Vanuga.

As with dairy cattle, numerous human pathogens are associated with flies from these locations, however, most are commensal or nonpathogenic to the cattle. Large populations of flies highly correlated with the potential for beef cattle to shed Salmonella spp., indicating fly-facilitated movement of bacteria among the herd (Vanselow et al. 2007). Neupane et al. (2019) examined the potential role of house flies in transmission of bovine respiratory disease (BRD). Flies harbored three primary BRD pathogens of concern; Mannheimia haemolytica, Pasteurella multocida, and Histophilus somni were recovered from flies collected near a pen of cattle exhibiting BRD. Although this implicates flies as a reservoir for these microbes, the role of the house fly in the epidemiology of BRD requires further examination.

Feedlots offer a wide range of potential house fly development substrates of manure and animal feed. Meyer and Petersen (1983) found that most house fly development occurred in sites that were protected from trampling by animals, especially along fence lines and drainage ditches. Skoda et al. (1993, 1996) found house fly larvae to be most abundant at the interface between the feed apron and soil, and other studies in Nebraska (Gilbertson and Campbell 1986) and Australia (Hogsette et al. 2012) have noted that house fly larvae are clustered along fence lines and in other protected sites (Fig. 14). Little breeding appears to occur in manure mounds within pens, apparently because of disturbance and trampling by the animals (Gilbertson and Campbell 1986). Protected larval development sites along pen margins are prone to drying, and house fly populations increase after rain events (Talley et al. 2002, Hogsette et al. 2012, Urech et al. 2012). Godwin et al. (2018) modeled house fly populations on Australian feedlots and found that rainfall events (85–90 mm/week) resulted in elevated adult house fly populations for 5 weeks afterward.

Figure 14.

Most fly larval development in feedlots occurs in protected sites such as fence lines and gaps between and beneath feeders, where spilled feed gets mixed with manure. Photos by Chris Geden.

The large size of many modern feedlots makes it difficult to implement farm-wide fly management programs and monitor their success. Sanitation practices such as improved drainage and manure removal from under fence lines and other preferred development sites are important but challenging to implement on very large operations (Clymer 1974, Thomas et al. 1996, Talley et al. 2002). Parasitoid releases may be helpful (Petersen et al. 1995, Floate 2003), but the number needed to be effective on a large feedlot may be prohibitively expensive. Parasitoid use on feedlots is discussed further below in the section on Biological Control.

Equine

House flies can have a negative impact on the welfare of horses. While house flies do not bite, their persistent presence on the body, eyes, mouth, and nose of horses increase stress and their potential as disease vectors. Pigeon fever, a highly contagious condition causing internal or external abscesses or limb infection called ulcerative lymphangitis, is caused by the bacteria Corynebacterium pseudotuberculosis. House flies can transmit C. pseudotuberculosis to horses (Barba et al. 2015). Pigeon fever was first reported in a 2015 horse health survey with 0.7% of equine farms reporting cases (approximately 6,887 farms) (USDA 2016). House flies also are biological vectors of Habronema spp. nematodes that can cause digestive disorders, diarrhea, progressive weight loss, ulcers, colic, and skin lesions (Amado et al. 2014, Pugh et al. 2014). Equine sarcoids, caused by bovine papilliomavirus type 1 (BVP-1), is one of the most common skin tumors in horses and other equids. Epidemiological data and spontaneous development of sarcoids without direct contact with affected individuals suggest flies may play an important role as mechanical vectors of BVP and BVP-1 has been isolated from house flies (Finlay et al. 2009).

House fly control challenges on equine facilities are generally related to the diversity of husbandry practices, uses, and ownership. Equid numbers on a facility may range from just one to >100 individuals. High-value animals used for show or racing may receive very little time in pasture, or more commonly a few hours to half a day. Conversely, pleasure or ranch horses may be pastured individually or in groups with little stall time. Bedding material used in stalls varies from straw, wood shavings, sawdust, or even newer products containing paper, peanut hulls, or hemp. Forage and feeding may be primarily from pasture or can be supplemented with hay fed in small or large flakes, or in large hay round bales as with cattle. Manure management practices vary with availability of land and local or regional regulations. Most facilities use manure pits or piles. Piles can be removed at regular intervals, left to accumulate, or applied to pastures or crop fields (USDA 2016).

Along with region and local conditions, management choices can influence house fly presence. House flies may develop in manure accumulation areas, pasture sheds or animal aggregation areas, waste hay in fields, or in stalls. In choice tests, house flies developed better in equine manure alone and manure mixed with pine shavings than in manure mixed with hay or straw (Machtinger et al. 2014), suggesting bedding choice influences fly development as it does in calf hutches (Schmidtmann 1991). In well-managed facilities, high house fly numbers may be a result of emigration from local or neighboring livestock or poultry facilities.

To reduce house fly presence on horses, many owners and managers rely on commercial fly repellents. In 2015, 76% of equine operations used on-animal repellents (USDA 2016), and 36.8% reported applying insecticides in or near equine housing areas, usually in the form of automatic misters. However, most commercial fly repellents and residual spray products are pyrethroid-based. These products generally have low concentrations of pyrethroid because of equine dermal sensitivity to higher levels (Stevens et al. 1988). High levels of pyrethroid resistance in house fly populations throughout the country have reduced the residual activity of many of these products to less than a few hours (Tuorinsky and Machtinger 2020) and thus they may not offer the protection needed to reduce annoyance or pathogen transmission. Recently, product formulations including fatty acids, geraniol, and other alternatives have been shown to induce more behavioral inhibition of house flies in laboratory settings and have longer residual activity (Tuorinsky and Machtinger 2020).

Monitoring and Management

House fly’s short development time, high fecundity, mobility, ability to exploit a myriad of developmental substrates, adaptability, and notorious propensity for developing resistance to new insecticides combine to make it challenging to manage. Maintenance of fly populations below acceptable levels requires a diligent IPM approach that takes full advantage of the many opportunities to combine monitoring with cultural, mechanical, biological, and chemical control approaches. Examples of available and potential tools for each of these elements is presented in the following sections.

Monitoring

Methods to monitor house fly activity have been described and evaluated for poultry facilities (Anderson and Poorbaugh 1964, Axtell 1970, Burg and Axtell 1984, Beck and Turner 1985, Lysyk and Axtell 1986), dairies (Pickens et al. 1972, Pickens and Miller 1987, Gerry et al. 2011), and beef cattle (Urech et al. 2004). General reviews of house fly monitoring methods and applications of those methods are also available (Lysyk and Moon 1994; Gerry 2020).

In 1945, Harvey Scudder was assigned by the U.S. Public Health Service to assess the efficacy of DDT for house fly control. To do this, he developed what is now known as the Scudder grid sampling method (Scudder 1947, 1996). The grid, a framed series of parallel wooden slats, is thrown to the ground where flies are concentrated and counts are made after the flies have settled again, usually 30 s later (Fig. 15). When multiple counts are made at several locations at a field site, the method can provide a simple and inexpensive instantaneous index of fly abundance at a location (Dhillon and Chalet 1985, Murvosh and Thaggard 1966). Population changes can also be monitored by repeated sampling over time (Madwar and Zahar 1951). Because house fly activity varies considerably with time of day and changing environmental conditions (Parker 1962, Zahn and Gerry 2020), grid counts should be recorded at a consistent time of day and with similar environmental conditions when sites are to be visited on multiple occasions.

Figure 15.

The Scudder grid can be useful for making instantaneous fly counts when only one species is present. Photo by Chris Geden.

Sticky traps of various kinds also can be used to monitor house flies. The conventional fly ribbon (Fig. 16) has been used for many years and provides more reliable estimates of fly abundance than grid counts (Anderson and Poorbaugh 1964, Raybould 1966, Pickens et al. 1972, Nurita et al. 2008). Ribbons also have the advantage of providing time-lapse rather than instantaneous counts and allow data collection on other species when present. Disadvantages of sticky ribbons are dust accumulations that compromise the adhesive and underestimations at high fly densities when tapes reach their carrying capacity of flies before they are collected (Rutz and Axtell 1981, Kaufmann et al. 2001a, Gerry et al. 2011). A variation on stationary tapes is the ‘walking sticky tape’ method, where a person carries the tape and walks at a consistent pace. This method has the advantage of providing an immediate estimate of fly abundance and has mostly been used to survey the upstairs of high-rise poultry houses (Turner and Ruszler 1989, Hinton and Moon 2003, Kaufman et al. 2005b). Peel-off sticky cards are a convenient alternative to ribbons that are easy to place and transport (Hogsette et al. 1993, Bell et al. 2019).

Figure 16.

Sticky tapes and cards are commonly used for monitoring house flies, especially when species identification is needed. Photos by Erika Machtinger.

Burg and Axtell (1984) described the use of baited jug traps for monitoring indoor populations of flies. These traps were standard 1-gallon milk jugs with holes cut in their sides that were baited with a dry sugar-insecticide bait and suspended from ceilings (Fig. 17). Flies entered the traps, fed, and died in place. The trap was used for several years (Lysyk and Axtell 1985, 1986a; Beck and Turner 1985; Stafford et al. 1988), but fly resistance to the toxicant in the bait (methomyl) rendered it impractical until new fast-killing insecticides became available. The method may be useful in locations where neonicotinoids are still effective. However, the use of this method for long-term monitoring (over years) must be carefully considered as increasing insecticide resistance in the fly population will reduce trap counts unrelated to fly density or activity.

Figure 17.

Baited jug trap used for monitoring house flies in indoor settings. This method is only effective when flies are sufficiently susceptible to the bait toxicant to die immediately after feeding. Photo by Chris Geden.

Spot cards (Fig. 18) are the most widely used monitoring method. First described by Axtell (1970), spot cards are standard 7.62 × 12.70 cm (3 × 5 inch) white cards that are attached to building structures and left in place to accumulate fly fecal and regurgitation spots. Cards are typically left in place for a week. A number of studies have compared spot cards with other sampling methods, and attempts have been made to use the cards to estimate absolute fly numbers (Beck and Turner 1985; Lysyk and Axtell 1985, 1986; Gerry et al. 2011). Although card counts can vary widely depending on temperature and card location and the fly species cannot be distinguished, they are easy and inexpensive to use and can be stored for years. They are particularly effective for monitoring relative abundance of indoor fly populations over time. In total, 100 spots per card per week has long been cited as an action or nuisance threshold (Axtell 1970), but this threshold is not based on scientific evidence.

Figure 18.

Spot cards are highly effective for monitoring relative fly activity over time if only one species is present. Cards with printed grid lines can help when making spot counts. Counting spots is challenging when fly populations are high. Photos by Erika Machtinger.

Most of the work discussed above has concentrated on relatively enclosed systems such as poultry houses and dairy barns in cooler climatic zones. These systems have geometric consistency and constraints that are helpful when implementing a monitoring plan. Monitoring is more complicated when a large portion of the fly population occurs outdoors such as on equine farms, feedlots, and large dairy farms in warmer parts of the world. Gerry et al. (2011) reported that spot cards were the most effective sampling method at dairies in Southern California, where frequent rains are not a problem. They also described a software application (FlySpotter) that automates the tedious process of counting the numbers of spots on cards (available at https://www.veterinaryentomology.org/flyspotter-house-fly-monitoring). Where frequent rains preclude the practical use of spot cards, sticky traps and liquid jar traps can be effective monitoring tools for outdoor fly populations. (Geden 2005, 2006; Gerry et al. 2011; Urech et al. 2012; Aziz et al. 2016). Baited plastic strips with toxicant can also be placed in sheltered locations for monitoring (Geden 2005), but only if flies are sufficiently susceptible to die close to the strips.

Cultural Control

Manure management is the cornerstone of any fly control program. Manure removal schedules are particularly critical. Mature larvae begin searching for pupation sites after 4–6 d under warm conditions, and often pupate away from the manure in protected and dry areas that are not cleaned out. Manure removal in facilities other than layer poultry therefore needs to be done at least twice a week to be effective during summer months. In caged-layer poultry systems, leaving residues of old manure during house cleanouts would be expected to encourage colonization of fresh droppings by fly predators, but recolonization by predators is slow under alternate row removal schedules (Peck and Anderson 1070, Geden and Stoffolano 1988; Mullens et al. 1996b, 2001). Leaving a pad of old manure may promote drying of fresh deposits and predator colonization (Hinton and Moon 2003), but Mullens et al. (1996a) observed no benefit from this practice in open-sided houses in California. Frequent removal of solid manure to liquid storage systems also prevents fly larval development, but such systems can produce troublesome populations of biting midges (Culicoides) and Culex mosquitoes.

When manure is removed, house fly larvae that are present in that manure can complete development after it is applied to cropland, even when it is incorporated into or covered with soil. Watson et al. (1998) estimated 89% of larvae in infested poultry manure were killed by mechanical damage during application with a manure spreader, and that only 25% of larvae produced adult flies when they were covered with 30 cm of soil. In spite of this attrition, the authors estimated that over 300,000 flies could emerge from a field on which manure from a poultry house had been applied. Tahir and Ahmad (2013) observed some fly emergence when pupae were buried under 30 cm of heavy clay soil, and Cook et al. (2020) found that house fly emergence from soil was unaffected by severe compaction. Composting is an alternative that can render manure unsuitable for fly larval development (Moon et al. 2001, Abu-Rayyan et al. 2010).

Moisture mitigation is another important component of cultural control. Because house fly larval development is optimal in substrates with 50–75% moisture (Fatchurochim et al. 1989), tactics that aid in drying out fly larval habitats are beneficial for managing fly populations. Water management practices include maintaining waterers, addressing drainage problems, and improving airflow in enclosed systems through the use of tunnel or turbo ventilation. Harrowing manure in animal pens can also aid in drying.

Mechanical and Physical Control

Mechanical and physical control options kill pests directly or make the environment unsuitable or inaccessible for pest flies. Screens or air curtains can reduce the entry of flies into sensitive areas such as milk and egg rooms (Mathis et al. 1970, Carlson et al. 2006), and high-speed fans in poultry houses can affect fly distribution in manure pits (Geden et al. 1999). Additives like hydrated lime, Ca(OH)₂, acetic and boric acid (Lachance et al. 2017) and sodium bisulfate (Sweeny et al. 2000, Calvo et al. 2010) have been used to dry manure and to make the pH of manure less suitable for fly development.

Ultraviolet light has long been recognized as a house fly attractant (Pickens et al. 1969, Pickens and Thimijan 1986). Electrocuting light traps sometimes have been reported to kill enormous numbers of flies in poultry houses (Rutz et al. 1988, Pickens et al. 1994, Hogsette 2019), but there are no data indicating that they are effective at reducing populations. UV traps are useful against small fly populations in sensitive areas such as dining areas and grocery stores (Lillie and Goddard 1987). Traps with glue boards are preferred to electrocuting models, as electrocution can result in dissemination of fly-borne pathogens into the local air space (Tesch and Goodman 1995, Urban and Broce 2000).

Sticky ribbons, sheets and cards can be useful but also are limited by the number of flies they can collect before becoming saturated and by the adhesive becoming ineffective from dust. Giant sticky ribbons are available in rolls that greatly expand the available surface area of adhesive traps. Kaufman et al. (2001a) reported that such large ribbons collected over 9 million flies during a 10-wk period in a poultry house and resulted in a measurable reduction in fly abundance in the upstairs of the house. In a subsequent study (Kaufman et al. 2005a), large sticky ribbons collected 900,000 house flies in a calf barn over 10 wk.

Odor-Based Traps and Baits

Attractant-based traps are a mainstay of fly management. There is a vast literature on house fly attractant strategies. Harper (1872) received a patent for the first inverted cone trap, and Howard (1911) described a bait made with fish heads, watermelon rinds, corncobs, and ice cream. Much of the research on fly attractants has focused on identifying components of food odors that can be incorporated into lures (Garrett 1965, Frishman and Matthysse 1966, Mayer 1971, Mulla et al. 1978). Early efforts with baits relied on natural products such as fermented egg slurries (Willson and Mulla 1973) or combinations of items such as molasses, milk, yeast, grain, blood, and banana extract (Pickens et al. 1973, Pickens and Miller 1987). Brown et al. (1961) tested a range of defined chemical attractant candidates and found that combinations were superior to any individual component tested alone. Mulla et al. (1977) reported that blends of trimethylamine, ammonia, indole, and linoleic acid were as attractive to house flies as natural food baits. More recently, Hung et al. (2015) found that house flies were attracted to honeydew-contaminated plant material and that associated fungi may play a role in the attraction. Specific compounds associated with honeydew contaminated plants that were attractive to house flies were (Z)-3-hexenyl acetate and benzaldehyde (Hung et al. 2019). Tang et al. (2016) examined components of fermenting wheat bran and found that a blend of ethyl palmitate, ethyl linoleate, methyl linoleate, and linoleic acid was attractive to gravid females.

In addition to feeding attractants, flies are attracted to (Z)-9-tricosene (muscalure) (Carlson et al. 1971, Carlson and Beroza 1973). The most common commercial feeding-attractant in use today, the Farnam Fly Attractant (now branded as the Starbar Fly Trap Attractant by Central Life Sciences), derived by modifying ratios of trimethylamine and indole and adding (Z)-9-tricosene. This attractant is used in granular sugar baits with a toxicant and in liquid jar traps without a toxicant such as the Terminator and Captivator brands. A wide variety of attractant-baited traps without toxicants are now available on the market, most of which are variations on the ‘jar’ or ‘jug’ design that lure flies into a container containing liquids in which the dead flies accumulate (Geden et al. 2009). Because these types of traps rapidly become ‘saturated’ when fly populations are high (Fig. 19), require frequent maintenance, and are malodorous, they are most appropriate for outdoor environments with light to moderate fly populations.

Figure 19.

Jar traps with liquid attractants and no toxicant are a mainstay of fly management but can fill with flies quickly, require frequent servicing, and are malodorous. Photos by Chris Geden (left) and Erika Machtinger (right).

Biological Control

Many methods have been researched for biological control of house flies including fungal, bacterial, or viral pathogens as well as predators, parasitoid wasps, and parasitic nematodes. Many of these methods have been explored extensively, while others are relatively new or have yet to be developed fully. There are specific benefits and challenges associated with each method that should be considered prior to incorporation in an IPM plan.

Entomopathogenic Fungi

Entomopathogenic fungi are naturally occurring fungi that kill insects and closely related arthropod hosts. They enter the host by invading the cuticle or being ingested, where they quickly proliferate, disseminating hyphal bodies and releasing various toxins. After the death of the host the fungus may sporulate on the exterior of the host cadaver if the humidity is high enough (Charnley 1989, Samson et al. 1988). Conidia are then dispersed to other uninfected individuals and the cycle continues (Kurtti and Keyhani 2008). The primary entomopathogenic fungi that have been evaluated for house fly control are Metarhizium brunneum (=anisopliae), Beavueria bassiana (Fig. 20), and Entomophthora muscae.

Figure 20.

House fly cadavers with Beauveria bassiana conidial blooms. Photo by Chris Geden.

Metarhizium brunneum Petch and B. bassiana Balsamo-Crivelli, while taxonomically distinct, are often tested and treated similarly in the literature. These are soil-dwelling fungal pathogens that are naturally found in filth fly populations. It is important to note that the M. anisopliae lineage was revised in 2009 (Bischoff et al. 2009). Many isolates of M. brunneum have been researched for biocontrol with some strains being developed as mycopesticides, however many of the articles detailing this research were published prior to 2009, when these isolates were classified as M. anisopliae. Here we refer to all M. anisopliae/brunneum isolates as M. anisopliae Sensu lato (s.l.).

The potential of B. bassiana as a biological control agent against pest flies was suggested by Dresner (1950). Steinkraus et al. (1990) first reported B. bassiana infections at a low prevalence (~1%) in house flies from New York state dairy farms. Since then, an abundance of literature has documented the potential efficacy of these fungal species to infect and kill immature and adult filth flies under laboratory (Hall et al. 1972, Kuramoto and Shimazu 1992, Barson et al. 1994, Geden et al. 1995, Watson et al. 1995, Angel-Sahagún et al. 2005, Lecuona et al. 2005, Kaufman et al. 2008, Weeks et al. 2017), and field conditions (Watson et al. 1996, Kaufman et al. 2005b, Cova and Scorza-Dagert 2006).

Beauveria bassiana and M. anisopliae s.l. have been tested against flies of various life stages, ages, and physiological characteristics. While house fly age does not seem to affect pathogenicity of B. bassiana or M. anisopliae s.l. (Rizzo 1977, Kaufman et al. 2008), female flies seem to be more susceptible than males (Rockstein and Lieberman, 1958, Lecuona et al. 2005, Anderson et al. 2013, Acharya et al. 2015a), and infection also reduces reproductive fitness (Acharya et al. 2015a). Horizontal transfer of conidia from males to females may be an avenue of dissemination of fungi in an otherwise adverse environment for fungal dispersal (Cárcamo et al. 2015).

Entomopathogenic fungi effects on immature house flies have been variable. Machtinger et al. (2016a) found decreased eclosion rates of house fly eggs treated with some commercial formulations of B. bassiana and M. anisopliae s.l. Exposure of larvae to B. bassiana has had little effect in some studies (Geden et al. 1995, Lecuona et al. 2005) and caused higher mortality in others (Steinkraus et al. 1990, Barson et al. 1994, Darwish and Zayed 2002). Young larvae are more susceptible than older larvae (White et al. 2020), possibly due to cuticular properties of developing larvae (Boucias and Pendland 1991). Immature flies may be more susceptible to M. anisopliae s.l. than B. bassiana (Bernardi et al. 2006, Fernandes et al. 2013). The responses of immature filth flies to fungal infection may also depend on the virulence of the fungal isolate, assay method, fly colony variability, conidial doses, substrate used, and culture methods. Larval resistance to fungal infection may be related to the strong immunological defenses that flies need to cope with the diversity of microorganisms found in their developmental environment (Nayduch and Joyner 2013). House fly larvae provide few suitable attachment sites for conidia and their constant movement through the larval substrate provides ample opportunity for conidia to be dislodged.

Various application methods for B. bassiana and M. anisopliae s.l. have been examined in laboratory and limited field settings, including topical applications (Sharififard et al. 2011), baits (Geden et al. 1995, Watson et al. 1995, Renn et al. 1999, Lecuona et al. 2005, Hong and Hai 2012, Mishra et al. 2013, Machtinger et al. 2016c), residual sprays (Watson et al. 1995, Blanford et al. 2012, Acharya et al. 2015b), and dusts (Geden et al. 1995, Watson et al. 1995). Aqueous and dust formulations can be applied directly on target flies or as a spray into the environment. Use of baits also has been suggested to slow insecticide resistance, reduce control costs, and reduce the effects on nontarget organisms by targeting fungal exposure to flies (Vega et al. 1995, Zimmer et al. 2010). However, there have been no published field studies assessing the efficacy of baits containing M. annisoliae s.l. or B. bassiana for control of house flies.

Significant differences in virulence have been noted among isolates of B. bassiana and M. anisopliae s.l. Fungal adaptation to host or host to fungal enzymes may influence virulence (St. Leger et al. 1986). Isolate differences have been noted in house fly adult and larval mortality (Mwamburi et al. 2010, Sharififard et al. 2011, Mishra and Malik 2012). When isolates adapt to their current host they may lose plasticity for invading other arthropods. Thus, the search for fungi to be used as control agents against filth flies should include isolates from the target insect (Poprawski et al. 1985, Steinkraus et al. 1991). Potential synergism between entomopathogenic fungi and bacterial pathogens or insecticides (Mwamburi et al. 2009, Sharififard et al. 2011, Farooq and Freed 2016, Johnson et al. 2018) may provide additional opportunities to increase virulence of fungi.

Virulence of B. bassiana and M. anisopliae s.l. to house flies may depend on environmental conditions. Temperature and relative humidity affect germination of conidia or virulence (Sivasankaran et al. 1998, Arthurs and Thomas 2001, Sharififard et al. 2012, Mishra et al. 2015). Ultraviolet light exposure, dust and particulate debris, and ammonia from livestock facilities can affect efficacy of M. anisopliae s.l. and B. bassiana (Leland and Behle 2005, Watson et al. 1995, Acharya et al. 2015b). These are serious considerations for development and deployment of M. anisopliae s.l. and B. bassiana in livestock, poultry, or equine facilities, but fungi may be protected by appropriate formulations (Morley-Davis et al. 1996, Alves et al. 1998, Reis et al. 2008).

Relatively few field studies have been conducted evaluating entomopathogenic fungi as filth fly control agents. An integrated management program including a spray formulation of B. bassiana in conjunction with pupal parasitoid releases was successful in caged-layer poultry in New York state (Kaufman et al. 2005b). The number of fly larvae recovered from the treated facilities was less than half the number recovered from pyrethrin treated facilities, moreover there was a greater recovery of beneficial insects. Watson et al. (1996) observed reduced adult house fly numbers after spray applications of B. bassiana in calf hutches. Applications of M. anisopliae s.l. reduced populations of house flies in treated poultry sheds two-fold compared to the control sheds (Fernandes et al. 2013). Conversely, weekly applications of B. bassiana were inadequate to achieve effective larval control in poultry houses in South Africa (Mwamburi et al. 2009).

The Entomophthora muscae (Cohn) Fresen (Entomophthorales: Entomophthoraceae) species complex, which includes E. muscae and E. schizophorae Keller and Wilding (Pinnock and Mullens 2007), is a group of fungi that have generally larger and more fragile conidia, faster germination times, a stronger tendency to produce conspicuous epizootics, and a narrower host range than the group that includes M. anisopliae s.l. and B. bassiana (Pell et al. 2001). Transmission of E. muscae occurs during a characteristic behavior change that causes house flies to anchor themselves to an elevated structure in the environment (Krasnoff et al. 1995). Following the death of the fly, large and sticky conidia are discharged into the environment where they can be encountered by healthy flies (Mullens and Rodriguez 1985, Six and Mullens 1996a, Kalsbeek et al. 2001a). Male flies, attracted to swollen, infected cadavers, can transfer conidia to healthy females (Watson and Petersen 1993, Zurek et al. 2002) and the infection renders females sterile.

Natural epizootic infections of E. muscae are well documented and primarily occur in the fall when temperatures are moderate and populations of flies are still high (Mullens et al. 1987a, Watson and Petersen 1993, Steinkraus et al. 1993, Six and Mullens 1996b). Because of the temperature sensitivity of E. muscae, flies are able to induce “behavioral fever,” whereby infected individuals locate areas of higher temperature to elevate their body temperature and kill the infection (Watson et al. 1993, Kalsbeek et al. 2001b). To what extent natural populations perform this behavior is not known.

Use of E. muscae for house fly control has significant challenges. The conidia of E. muscae are fragile and cannot be held for future use like M. anisopliae s.l. and B. bassiana (Kalsbeek et al. 2001a). Methods for mass production of E. muscae have been developed (Kramer and Steinkraus 1981, Mullens 1986), but the introduction of the fungus to the target population requires using either live infected flies or cadavers of recently infected flies. Attempts to control house fly populations with E. muscae in the field have had limited success to date (Steinkraus et al. 1993, Geden et al. 1993, Six and Mullens 1996b).

Entomopathogenic Bacteria

Bacillus thuringiensis Berliner (Bt) is an aerobic, Gram-positive, spore-forming bacterium (Mwamburi et al. 2009). Bacillus thuringiensis can be isolated from many environmental sources such as soil, insects, and coniferous and deciduous leaves (Schnepf et al. 1998). Formulations of Bt have been used for decades as biopesticides for many agricultural pests.

Early work demonstrated that feeding animals (cattle and poultry) Bt could control larvae developing in the resulting manure (Burns et al. 1961, Miller et al. 1971). Larval control was also obtained by mixing Bt into larval development sites (Rupes et al. 1987). However, fly resistance to the exotoxins produced by the tested Bt strains quickly made these techniques unsuitable for control (Harvey and Howell 1965, Wilson and Burns 1968). Additional vertebrate safety concerns were raised, which led to the prohibition of exotoxin-producing Bt in the United States (McClintock et al. 1995, Tsai et al. 2003).

Subsequent evaluations of exotoxin-free Bt were disappointing. However, several strains of Bt were identified that were effective against house fly larvae and adults (Indrasith et al. 1992, Hodgman et al. 1993, Johnson et al. 1998, Zhong et al. 2000) and it was discovered that the δ-endotoxin Cry1B was found in all the Bt strains that were active against house flies. The endotoxins are safe for humans, other vertebrates, plants, and are biodegradable (de Barjac 1978, Adang et al. 2014).

Current research has focused on Bt for house fly control in topical applications and as a feed additive. While application of Bt to larval development sites has resulted in some mortality of flies, greater success has been from use of Bt as an animal feed additive. Promising control when fed to poultry (Labib and Rady 2001; Mwamburi et al. 2009, 2011; Merdan 2012) and horses (Martins 2013) has been observed. There is promise in the use of Bt as a biological control method for house flies in a variety of animal production systems.

Salivary Gland Hypertrophy Virus

House fly salivary gland hypertrophy virus (MdSGHV) is a house fly-specific virus that replicates in salivary glands, generally resulting in obvious hypertrophy of the glands and sterility of female hosts (Lietze et al. 2011b) (Fig. 21), although virus particles can be found in other tissues (Lietze et al. 2011a, Kariithi et al. 2017). House flies are infected with MdSGHV globally, with prevalence rates usually of less than 2% (Geden et al. 2008, 2011a; Prompiboon et al. 2010; Lietze et al. 2011b; Lietze et al. 2013).

Figure 21.

A healthy fly (A) and a fly infected with salivary gland hypertrophy virus (B), showing underdeveloped ovaries (Ov) and overdeveloped salivary glands (Sg) in the infected fly (Mg = midgut). Photo by Lyle Buss, University of Florida (Florida, USA).

Infection is thought to occur via food sources shared between infected and healthy flies (Lietze et al. 2009, 2013); thus, infective baits have been considered for fly control purposes. However, feeding on infected material results in limited infection of flies in the laboratory (Geden et al. 2008, 2011a; Lietze et al. 2009; Lietze et al. 2011c), likely because house fly susceptibility to the virus is only during a brief window after adult eclosion, when they would not normally be feeding. The PM is an effective barrier to oral infection, as susceptibility in older flies can be restored if they are given drugs that disrupt the PM (Boucias et al. 2015). An alternative approach to infective baits is the use of a homogenate of infected flies sprayed in areas of house fly activity (Geden et al. 2011). However, the efficacy of this approach has not been demonstrated in the field.

Predators

Three families of mites, Macrochelidae, Uropodidae, and Parasitidae, are known to prey upon fly eggs or larvae and naturally occur in poultry manure (Axtell 1961, 1963a). One of the most common predatory mites is Macrocheles muscaedomesticae (Scopoli) (Fig. 22). Some experiments have demonstrated upwards of 20 house fly eggs consumed by M. muscaedomesticae per mite per day (Geden and Axtell 1988, Geden et al. 1988). Reductions in house fly numbers in cattle manure and poultry manure have been demonstrated under simulated field conditions with the use of M. muscaedomesticae (Axtell 1986). Reviews of the literature and an assessment of Macrochelidae as biological control agents have been published (Axtell 1969, Geden 1990). Populations of M. muscaedomesticae are important for the natural suppression of house flies (Axtell 1963b), however this species has not been developed commercially.

Figure 22.

The two most important predators of house fly immatures are the mite Macrocheles muscaedomesticae (left) and the histerid Carcinops pumilio. Photos by Eric Palevsky, Agricultural Research Organization, Israel and Erika Machtinger, respectively.

Many beetles are important predators on fly eggs and larvae, but the major predaceous species is the histerid Carcinops pumilio (Erichson) which often is very abundant in poultry manure; adults and larval stages prey on fly eggs and larvae (Morgan et al. 1983) (Fig. 22). A single beetle adult may consume 13–83 house fly eggs per day and beetle larvae consume 13–26 eggs per day (Geden and Axtell 1988, Geden et al. 1988). The wide distribution of these beetles as well as abundance and high rate of predation make C. pumilio a major predator of house flies. These beetles are typically found in poultry pit houses (Geden 1984) and are less common in other animal facilities. Carcinops pumilio can be purchased from commercial insectaries for release, and populations can be collected with species-specific traps to re-release after pit cleanout.

Other fly species are also predators of immature house flies. Black dump fly (Hydrotea aenescens [Wiedemann]) is a naturally occurring muscid that is most commonly found on poultry and swine facilities. It is known in the older literature as Ophyra aenesens Weideman. Larvae of this species are facultative predators that can develop in substrates without prey (Hogsette and Washington 1995, Farkas and Hogsette 1998, Hogsette et al. 2002) but are also capable of killing 15 house fly larvae per day (Geden et al. 1988). Adult flies prefer dark spaces and are less prone to becoming nuisance pests than house flies (Nolan and Kissam 1987). Black dump flies have been studied as potential biological control agents for house flies, and augmentative releases sometimes have resulted in its dominance over house flies (Nolan and Kissam 1985, Turner and Carter 1990, Turner et al. 1992). They are available from commercial insectaries in the United States and Europe. However, there are concerns about release of this species as their ability to transmit pathogens mechanically may be similar to that of other pest flies (Olsen and Hammack 2000, Szalanski et al. 2004).

Parasitoids

Naturally occurring pupal parasitoids (Hymenoptera: Pteromalidae) have been used for control of house flies on animal facilities for nearly 50 yr. House fly parasitoids occur naturally in all areas where suitable hosts can be found. Naturally occurring parasitoid populations are not sufficient to manage flies at acceptable levels because they are slower to develop than the fly host. However, augmentative releases of parasitoids can be effective at suppressing pest fly populations when coupled with other management methods (Geden et al. 1992, Skovgård and Nachman 2004, McKay et al. 2007). Parasitoids are commercially available, and their use has increased by some commodity groups, likely in part due to the easy availability (USDA 2006, Machtinger et al. 2013). The taxonomy, natural history, and use of parasitoids have been reviewed in Machtinger and Geden (2018) and Machtinger et al. (2015b). Here we will review available species for house fly control programs and use on the target animal commodity groups.

Although a wide diversity of species parasitize house fly pupae, the most commonly collected and commercially available parasitoids include Muscidifurax raptor Girault and Sanders, M. zaraptor Kogan and Legner, M. raptorellus Kogan and Legner, Spalangia cameroni Perkins, S. endius Walker, Nasonia vitripennis (Walker), and Trichomalopsis sarcophagiae (Gahan) (Fig. 23). Most of these species have similar life histories where the female parasitoid locates a suitable fly puparium, drills through the puparium, and deposits either one (solitary species) or multiple eggs (gregarious species) on the surface of the host pupa (Gerling and Legner 1968). The resulting parasitoid larva (or larvae) consumes the pupa and emerges as an adult 2–4 wk later. Development of a parasitoid from egg to adult takes 14–30 d under warm conditions depending on species, sex, host, temperature, environment, and biotypes (Birkemoe et al. 2012).

Figure 23.

Examples of the two most important genera of house fly parasitoids; Spalangia (S. cameroni, left) and Muscidifurax (M. raptor, right). Photos by Erika Machtinger.

Evaluations of parasitoid releases on filth fly populations have demonstrated reduction in some situations (Morgan and Patterson 1990, Geden et al. 1992, Petersen and Cawthra 1995, Crespo et al. 1998, 2002, Skovgård and Nachman 2004, Geden and Hogsette 2006), but not in others (Morgan 1980, Meyer et al. 1990b, Andress and Campbell 1994, Weinzierl and Jones 1998, McKay and Galloway 1999, Kaufman et al. 2001c). Biological characteristics of released parasitoid species that may influence effectiveness include parasitoid microhabitat preferences, intra- and interspecific competition (Machtinger and Geden 2015, Taylor et al. 2016), and factors influencing parasitoid abundance and distribution (Skovgård 2004). Colony quality, previously established parasitoid populations, and the lack of understanding of timing and methods of parasitoid release could also have impacted the success of releases (Peterson et al. 1983, Patterson and Rutz 1986, Legner et al. 1990). Nasonia vitrpennis does not appear to be effective in poultry, dairy, cattle feedlot, or swine systems (Stage and Petersen 1981, Meyer et al. 1990b, Andress and Campbell 1994, McKay and Galloway 1999, Kaufman et al. 2001c, Birkemoe et al. 2004).

Parasitoid releases in poultry facilities were some of the earliest evaluations (Legner and Brydon 1966). The relative stability of habitat in poultry facilities and presence of other predators, including beetle and mites, make these facilities well-suited for parasitoid releases (Geden 1990). Reduction in house fly numbers has been achieved with releases of S. endius (Morgan et al. 1975a; Morgan et al. 1975b; Morgan et al. 1981a,b; Morgan and Patterson 1990), M. raptor and M. raptorellus (individually or in tandem) (Rutz and Axtell 1979, Crespo et al. 1998, Kaufman et al. 2002, Al-Ani et al. 2012), and combinations of M. raptorellus and S. cameroni (Geden and Hogsette 2006, McKay et al. 2007).

Dairy facilities often have fly-breeding sites that have some measure of habitat constancy and environmental protection. However, house fly control as a result of parasitoid releases has been more variable. High levels of parasitism have been achieved with releases of S. endius (Morgan and Patterson 1977), S. cameroni (Skovgård 2004; Skovgård and Nachman 2004), M. raptor (Geden et al. 1992), and M. raptor and M. raptorellus combinations (Kaufman et al. 2012). Conversely, M. zaraptor and M. raptor releases had negligible impacts on adult fly populations on dairies (Meyer et al. 1990b, Miller et al. 1993), although numbers of parasitoids released were not as high as in other studies.

In cattle feedlots, fly development sites are spread over large areas and often disturbed and exposed to the environment, making parasitoid-based fly control more challenging. The range of dispersal for pteromalid parasitoids is limited (Pawson and Petersen 1988, Tobin and Pitts 1999, Skovgård 2002, Machtinger et al. 2015c). Releases of S. endius (Morgan 1980), and locally collected M. zaraptor and M. raptorellus (Petersen et al. 1992, 1995; Petersen and Cawthra 1995; Petersen and Curry 1996), S. nigroaenea Curtis (Weinzierl and Jones 1998) and combinations of M. raptorellus and Trichomalopsis sarcophagae Gahan (Floate 2003, Floate et al. 2000) have been effective to some degree in reducing fly populations in confined cattle. However, S. endius releases (Petersen et al. 1983) and mixed releases of M. zaraptor, N. vitripennis, S. endius, and S. nigroaenea (Andress and Campbell 1994) have not demonstrated control. In both these latter cases, species may have been released in areas where they were not well-adapted.

Information on use of parasitoids at other animal facilities is limited. On swine facilities, releases of S. endius and S. cameroni have had some effect (Morgan 1980, Skovgård and Nachman 2004). While Spalangia spp. have been the dominant species recovered from equine farms (Pitzer et al. 2011, Machtinger et al. 2016b), these studies were limited to surveillance.

Parasitic Nematodes

Entomopathogenic nematodes in the families Steinernematidae (Steinernema spp.) and Heterorhabditidae (Heterorhabditis spp.) (Order: Rhabditida) have been reviewed extensively (Kaya and Gaugler 1993, Gaugler 2002, Georgis et al. 2006, Koppenhöfer 2007, Poinar and Grewal 2012, Atwa 2014). These parasites are vectors of bacterial symbionts in the genera Xenorhabdus and Photorhabdus (Enterobacteriales: Enterobacteriaceae), respectively (Forst et al. 1997, Boemare 2002), that account for most of the virulence associated with the nematodes. House fly larvae and adults are highly susceptible to Steinernema feltiae (=carpocapsae) Filipjev (Rhabditida: Steinernematidae) and Heterorhabditis heliothidis (=bacteriophora) (Khan, Brooks and Hirschmann) (Rhabditida: Heterorhabditidae) under laboratory conditions (Geden et al. 1986).

Applications of entomopathogenic nematodes can target immature or adult house flies. However, successful applications of nematodes to control immature house flies may depend on larval development areas. Efficacy in poultry manure is poor (Renn et al. 1985, Georgis et al. 1987, Mullens et al. 1987b, Mullens et al. 1987c, Archana et al. 2017), but may be better in cattle or swine manure (Taylor et al. 1998, Renn 1998). Additives, such as calcium alginate capsules, may allow nematodes to be more effective by reducing unintended mortality (Renn 1995). Like entomopathogenic fungi, species and strain may influence virulence for fly control (Taylor et al. 1998, Wojciechowska et al. 2013), thus the development of superior strains and formulations may have greater impact on fly control. Adult house flies also can be targeted with nematodes (Renn et al. 1999). Delivery method and complementary additives or formulation may be critical to high infectivity (Geden et al. 1986, Renn 1998, Renn and Wright 2000).

Entomopathogenic nematodes are attractive candidates as biological control agents. They often have a wide host range, can be easily mass-produced at low cost, have a long storage life (especially Steinernema spp.), can be selected for desirable traits, are compatible with certain insecticides, are safe for vertebrates, and are easily applied in the field. However, efforts will need to be made to conduct field research to demonstrate utility for house fly control in various livestock and equine facilities, and improvements to delivery systems could support their use in poultry facilities.

Black Soldier Fly: a Special Case

Black soldier fly (Hermetia illucens (L.)) is found in temperate and tropical regions and has received considerable attention for its ability to decompose animal waste (Gold et al. 2018, Lalander et al. 2019) and convert it to an insect source of animal and human food (Wang and Shelomi 2017, Beskin et al. 2018, Tomberlin and Van Huis 2020). Soldier flies are most common in poultry layer facilities, where the activity of their larvae modifies the manure habitat in ways that make it unsuitable for house fly larvae (Furman et al. 1959, Sheppard 1983). The presence of their larvae in manure also inhibits oviposition by house flies (Bradley and Sheppard 1984).

Chemical Control

Many chemicals are used in pest management, but differ in range of action, toxicity, and persistence in the environment. Insecticides are synthetic or natural compounds that act as direct toxins to house flies. Control of house flies is most commonly attempted with conventional insecticides because of their rapid action, low cost (relative to other strategies) and effectiveness (at least for new insecticides). However, resistance can radically reduce the effectiveness of insecticides for fly control. The specific insecticides available for fly control vary by state (Gerry 2020) (https://www.veterinaryentomology.org/vetpestx). A 2007 survey of U.S. dairies indicated numerous insecticides were used for house fly control in the United States [chlorpyrifos, cyfluthrin, cyhalothrin, cypermethrin, dichlorvos, diflubenzuron, fenvalerate, imidacloprid, methomyl, naled, permethrin (some formulations contain piperonyl butoxide (PBO), pyrethrins + PBO, spinosad, and tetrachlorvinphos (some formulations contain dichlorvos)], although there was significant variability in each state. At present, the main active ingredients used in fly-control insecticides are pyrethroids (e.g., permethrin, β-cyfluthrin, λ-cyhalothrin) in space sprays/premise treatments and neonicotinoids (imidacloprid, dinotefuran, and nithiazine) in a variety of baited delivery systems. Table 4 provides a list of the common insecticides that are currently used for house fly control.

Table 4.

Documented house fly resistance to insecticides with different modes of action

Insecticide or insecticide class	Target life stage	Delivery methodsa	Cases of resistance in North Americab
Carbamates	Adult	Spray, bait	Forgash and Hansens 1959, Georghiou et al. 1961, Plapp and Bigley 1961, Georghiou 1962, 1966, Harris et al. 1982, Price and Chapman 1987, Bull and Pryor 1990, Liu and Yue 2000, Scott et al. 2000, Kaufman et al. 2001b, Darbro and Mullens 2004, Scott et al. 2013, Freeman et al. 2019
Organophosphates	Adult	Spray, dust, feed-through, impregnated strip	Fay et al. 1958, Hansens 1958, Labrecque et al. 1958, Forgash and Hansens 1959, Harris and Burns 1959, Wilson et al. 1959, Forgash and Hansens 1960, Hansens 1960, Bigley and Plapp 1961, Labrecque and Wilson 1961, Georghiou 1962, Georghiou and Bowden 1966, Forgash and Hansens 1967, Georghiou 1967, Mathis et al. 1967, Georghiou and Hawley 1971, Georghiou et al. 1972, Harris et al. 1982, Boxler and Campbell 1983, MacDonald et al. 1983, Bloomcamp et al. 1987, Bull and Pryor 1990, Liu and Yue 2000, Scott et al. 2000, Kaufman et al. 2001b, Marcon et al. 2003, Scott et al. 2013, Freeman et al. 2019
Pyrethrins and Pyrethroids	Adult	Spray, fog/aerosol	Forgash and Hansens 1959, Georghiou 1962, DeVries and Georghiou 1980, Harris et al. 1982, MacDonald et al. 1983, Scott and Georghiou 1985, Meyer et al. 1987, Bull and Pryor 1990, Meyer et al. 1990a, Chapman et al. 1993, Liu and Yue 2000, Scott et al. 2000, Kaufman and Rutz 2001, Kaufman et al. 2001b, Marcon et al. 2003, Kaufman et al. 2010, Scott et al. 2013, Freeman et al. 2019
Neonicotinoids	Adult	Bait, spray, impregnated strip	Liu and Yue 2000, Kaufman et al. 2006, Kaufman et al. 2010, Scott et al. 2013
Spinosyns	Adult	Bait, spray	Scott et al. 2000, Deacutis et al. 2007
Pyriproxyfen	Larvae	Spray, fog/aerosol	Cerf and Georghiou 1972, Plapp and Vinson 1973, Cerf and Georghiou 1974, Georghiou et al. 1978, Pimprikar and Georghiou 1979c
Benzoylureas	Larvae	Spray, feed-through	Pimprikar and Georghiou 1979, Shen and Plapp 1990
Cyromazine	Larvae	Feed-through, spray	Pimprikar and Georghiou 1979, Iseki and Georghiou 1986, Scott et al. 2000
Indoxacarb	Adult	Spray	Shono et al. 2004
Diamides	Adult	Bait	None as of April 2020

^aCurrent and past methods. Some approved methods are rarely used. Sprays = premise, animal and/or manure applications (see https://www.veterinaryentomology.org/vetpestx for details)

^bCases of cross-resistance.

^cCases of methoprene resistance.

Insecticide Resistance and Its Consequences

Resistance has developed to all available insecticides used for house fly control and is a global problem (Cao et al. 2006, Scott et al. 2013, Scott 2017). The speed with which a given insect species evolves resistance can vary considerably, and in some cases is remarkably rapid (after a single spray season). House flies have consistently been one of the species in which resistance evolves quickly. Resistance has evolved rapidly and globally to nearly every class of insecticide, including organochlorines, organophosphates, carbamates, pyrethroids, insect growth regulators, neonicotinoids and spinosyns (Keiding 1986, Gerry and Zhang 2009, Scott et al. 2013, Table 4). Recently, facilities have been identified where complete failure of house fly control by pyrethroids was observed (Freeman et al. 2019). This has caused tremendous concerns at those facilities (one was cattle and one was poultry) and has growers and extension agents scrambling for alternatives. Thus, the evolution of resistance is pitted against the ingenuity of humans to invent new and safe insecticides. There are few alternative options for house flies. There are, however, new insecticides that are registered for use on other insects, but not house flies, and some of these may be effective against house flies. The Food Quality Protection Act of 1996 assesses risk based on aggregate exposures, and has led to very focused registrations for new insecticides over the last two decades as companies selected markets that would offer the greatest profitability. As a result, for the last 20 yr there have been no new insecticides coming to market that were labeled as premise sprays for house flies in the United States. Thus, if resistance to pyrethroids renders these insecticides ineffective over a wide region (beyond the single facility failures that have been observed), there are only older insecticides available to owners of livestock production facilities as premise treatments. Almost all of these older insecticides are also plagued by resistance issues (Table 4) and have unfavorable environmental and mammalian toxicological profiles (e.g., organophosphates). To complicate matters further, house flies have tremendous potential for cross-resistance to arise from overuse of a single chemical (e.g., Wen and Scott 1997, Shen and Plapp 1990, Abbas et al. 2015, Scott 1989). Baits containing neonicotinoids and spinosad were highly effective at the time of introduction but resistance has limited their effectiveness as well (Table 4).

New Tools for House Fly Management

Background

Control technologies outside of the traditional chemical insecticides and biological control agents discussed above have long been of interest to researchers. Household chemicals such as borax, bleach (chloride of lime), and table salt were studied for addition to manure to reduce fly production. Early concerns were that such larvicides might have detrimental effects on manure that might decrease its value as fertilizer (Cook et al. 1914, Hewitt 1914a,b). Further motivation for discovering alternative chemical control technologies came shortly after the DDT product GNB (Gerasol-Neocid-Base) came to market in 1942. By the late 1940s, studies of DDT failure in house fly control began to appear in the literature (e.g., Barber and Schmitt 1948, Lindquist and Wilson 1948, Wilson and Gahan 1948). Shortly after, novel methods to restore the usefulness of some important insecticides like DDT and pyrethrum were developed (Wilson 1949). These came in the form of the insecticidal synergists piperonyl cyclonene and piperonyl butoxide (PBO), the latter of which is still widely used in house fly control products. More attention was also turned to traps and attractants (Yates 1951, Muto and Sugawara 1965).

Current Outlook

Today, with widespread resistance to nearly every insecticide marketed for house fly control (Freeman et al. 2019), discovery of novel chemistries has never been more important. Chemistries that have insecticidal activities or synergistic potential with currently used insecticides are of particular interest. One broad group of compounds, the essential oils, have received considerable attention recently (Pavela and Benelli 2016, Khater and Geden 2019). However, of the hundreds of studies published on their insecticidal potential per year for numerous insect pest species, few provide the chemical information or positive control comparisons (i.e., comparisons to current insecticides) necessary for proper evaluation (Isman and Grieneisen 2014). It is therefore imperative that future work include these important parameters. An active component of eucalyptol, 1,8-cineole, appears to be among the most potent (Palacios et al. 2009, Rossi and Palacios 2015). When applied as a fumigant, toxicity of 1,8-cineole (LC₅₀: 3.3 [1.1–10.4]) mg/dm³, was similar to deltamethrin (LC₅₀: 9.2 [2.8–29.5] mg/dm³). 1,8-cineole also appears to synergize well with the pyrethroid deltamethrin (deltamethrin + 1,8-cineole: 1.0 [0.08–12.1]) mg/dm³, deltamethrin + PBO: 1.5]0.2–11.4)] mg/dm³). Other essential oils such as geraniol show some repellent potential and are sometimes classified as minimum risk compounds (USEPA 2015). Geraniol-based fly repellent sprays are popular on the market (e.g., Outsmart, Pyranha Zero Bite All Natural).

Few new traditional insecticides have come to market for house fly control over the past 5 yr. Perhaps the most prominent is cyantraniliprole, belonging to the second-generation of anthranilic diamide insecticides (Yu 2015). Insecticides in this class bind to the insect ryanodine receptor, which is a new mechanism of action that minimizes the chance for cross-resistance from previous insecticide use. Cyantraniliprole was registered in a house fly granular bait in 2015 (Zyrox, renamed Cyanarox). In 2018, BASF initiated an EPA registration of the meta-diamide broflanilide (federal register citation: 83 FR 34128). Included in the registration is use of broflanilide in two fly baits (Vedira) targeting multiple filth flies, including house flies. No new insecticides have been registered as premise treatments for >20 yr.

Within the past few years, the non-nutritive polyols erythritol and xylitol have been assessed for their potential as house fly adulticides and larvicides, with mixed results (Burgess and King 2017, Fisher et al. 2017, Burgess and Geden 2019). Under no-choice conditions, over 50% of flies die within two days when exposed to 2 M solutions of the polyols. However, given acute exposures of 24 h, followed by access to normal food, fly mortality barely reached 25% after 20 d, and was not significantly different from the normal food control. Similarly, larvicidal LC₅₀ values against late second instar flies were in the parts-per-thousand range for house flies, making them orders of magnitude worse than typical insect growth regulators. Lower concentrations were very effective at killing eggs/early instar larvae, with 0% of eggs developing to adults in the erythritol treatment and only 23.3% of eggs developing to adults in the xylitol treatment. This is compared to 46.0 % of control eggs developing to adults. The polyols erythritol and xylitol also appear to be attractive substitutes for sucrose in baits, especially those centered around the entomopathogenic fungus, Beauveria bassiana (Burgess et al. 2018). Current work suggests that they may also play a role in the house fly gut physiology and microbiome, potentially leading to new methods for controlling house flies by exploiting their physiology in ways independent of traditional insecticides.

House flies are not generally regarded as suitable candidates for conventional (irradiation-based) sterile insect technique (SIT) methods. Population densities are too large and fluid to overwhelm with irradiated males, and male flies are as pestiferous and capable of transmitting pathogens as females. Emerging technologies may create new opportunities to examine SIT and other genetic methods (Scott et al. 2018). Promising results with mosquitoes have been seen using Wolbachia/cytoplasmic incompatibility (Bourtzis et al. 2014, Mains et al. 2019) and the release of genetically modified males to deliver dominant lethal alleles (RIDL) (Harris et al. 2012). Moreover, the availability of the genome of house flies (Scott et al. 2014) and related species may lead to novel management approaches that exploit vulnerabilities in the life cycle. Little work has been done using RNAi in adult house flies, but constructs have been identified that, when injected, shut down ovarian development (Sanscraint et al. 2018).

Research Priorities

The house fly remains a formidable adversary with many unanswered questions despite over 100 yr of scientific research. Topics that warrant highest priority for future research needs fall under four broad categories.

Management

Monitoring flies remains a highly inexact science. More research is needed to identify which monitoring tools are most appropriate in different animal commodity systems. Nuisance/action/economic thresholds are in urgent need of revisiting, as current threshold values are based on little more than traditional use that has been repeated for nearly 50 yr. The use of monitoring house fly density in the decision making for insecticide applications is sorely needed.

Biological control has won acceptance as a management tool in some settings, but there are still gaps in availability of products that are effective and economically feasible. Many mycopesticides have been developed and labelled for crop pests, and screening these for house fly control would facilitate label expansion to include this pest. Dozens of laboratory studies on efficacy of fungal pathogens for house flies have been conducted, but only a handful of field tests have been published. Field testing of promising pathogens is critically needed to encourage commercial producers to invest in developing pathogens as viable and competitive products. One of the main liabilities of fungal pathogens is their slow kill rate compared to chemical insecticides. Further research is needed to identify fast-killing strains and improve the virulence of known strains by selection, genetic modification, and better formulations either as stand-alone agents or in combination with other components and adjuvants. New formulations could also include baits and attract-and-infect stations that could be used as autodissemination devices. Other biological control research needs include the development of predator mites for augmentative control and improvements in timing, species selection, and release rate recommendations for pteromalid parasitoids in different animal systems.

Traps are frequently used by producers (USDA 2014) and will always play an important role in house fly management, yet there have been no substantial improvements in house fly attractants for traps in over 40 yr. Renewed research is needed to identify superior attractants, especially ones that are not malodorous to humans and that would preferentially attract female flies.

Novel chemistries are needed to develop new insecticide products to provide rapid reduction of house fly populations resistant to currently available insecticides. Screening of currently-registered insecticides not already in use for house fly control is also needed, with special emphasis on space sprays and modes of action distinct from those of pyrethroids, neonicotinoids, and organophosphates. Integrated resistance management (IRM) plans that take an area-wide approach would be helpful in extending the effective product life of those insecticides that remain effective. Screening of essential oils and other plant-derived compounds for insecticidal and repellent potential should continue, with rigorous testing standards that include testing against positive controls and inclusion of information on chemical composition and purity. New insecticide synergists are needed as well, especially molecules that will fit an OMRI or EPA minimal risk profile.

Rising global temperatures will require fly management to be conducted under increasingly hot conditions in many locations. High temperatures can impact the effectiveness of biological (Geden et al. 2019) and chemical control (Scott and Georghiou 1984). Research on fly management under hot conditions, especially field studies, is needed to prepare for future challenges.

Biology and Life History

There is no satisfactory answer to the important question of how flies typically live in the field. Field longevity experiments are notoriously difficult but are needed to identify factors affecting fly survivorship under natural conditions. For example, how is longevity affected by animal production systems, food resources within those systems, and locally available microbiomes? Research on fly functional genomics, transcriptomics, and proteomics, leveraged by knowledge gained in other dipteran pests, may reveal as-yet undiscovered fly vulnerabilities that can be exploited. We are just beginning to explore the roles that microbiomes play in larval and adult fly fitness. Further research on this topic is critically needed and could point to ways to interfere with the acquisition and retention of needed microbes. Several important aspects of fly behavior are still poorly understood, especially regarding flight activity and dispersal of flies. A better understanding of the role of fly sex, local availability of food resources, and fly response to environmental conditions could help identify triggers that lead to dispersal and increased synanthropy.

Role as Reservoirs and Transmitters of Pathogens

Flies that originate from concentrated animal feeding and production operations can disseminate microbes within the operations (creating an animal health concern) or away from operations (creating a human health concern). The studies listed in Table 3, as well as many other surveys (reviewed in Nayduch and Burrus 2017), demonstrate that wild-caught flies harbor a diverse and abundant community of human and animal pathogens in a variety of settings. Furthermore, the studies we review above demonstrate that flies can acquire, harbor, and transmit a number of pathogenic bacteria, and may facilitate lateral transfer of resistance and virulence genes among microbes in the gut. Several critical research gaps remain in defining the role house flies play as both reservoirs and transmitters of microbes. What are the most important microbial and fly-specific variables that impact acquisition, persistence and transmission of pathogens? What factors underlie the temporal dynamics of pathogen persistence in infected flies? How long do flies harbor pathogens? How long do they shed pathogens? Are pathogens being shed in doses sufficient to infect animals or humans? How significant a role does house fly play in contaminating human foods/crops with pathogens? A better understanding of these factors, along with variables from prior studies outlined in this review, should be used to develop predictive models as improved risk assessment tools for transmission of animal and human pathogens. Further demonstrations also are needed to document that effective fly management mitigates pathogen prevalence in the field.

Systems Needs

Intensive animal agriculture production systems provide materials that attract adult house flies and allow for the development of immature house fly populations on site. These include animal manures, collected and stored, and feed ingredients, many of which are suitable for house fly development. These materials are continually replenished. Fly management programs in all intensive system focus on fly habitat management/reduction, and the available management tools are applicable across different animal production systems. However, additional research is needed to develop more effective manure and feed storage systems that limit fly access and are easy to use.

Equine systems have received the least research attention with regard to house fly management. Fly populations tend to be comparatively low, and many of the flies present may have been produced off-site on cattle or other facilities. Research is needed to determine high-risk larval development habitats and to assess the proportion of flies on horse farms that have developed on-site. Many equine facilities use fly parasitoids, yet almost nothing is known about the efficacy of parasitoid releases on horse farms. There are currently no scientifically based recommendations for the numbers or even the species of parasitoids needed to keep flies at acceptable levels in this difficult market.

The poultry industry is changing rapidly and is now composed of organic and conventional production systems using a wide array of housing types, flock sizes, and management practices. Much of the research on house fly management has been conducted on farms with high-density caged layer houses that house >100,000 hens each. Europe and California have banned this type of housing, and future fly control work will be conducted on smaller open-floor facilities of different designs. Research is needed to assess how fly populations are affected by housing type and husbandry practices in this rapidly evolving industry. The swine industry faces similar challenges, with a range in production types and over wide geographic areas. Best management practices are critically needed in the poultry and swine industries for fly monitoring and biological control options under different housing, management, and climate conditions.

Acknowledgments

We thank members of the S1076 multistate project ‘Fly Management in Animal Agriculture Systems and Impacts on Animal Health and Food Safety for manuscript discussion and advice. We also appreciate the financial support from Penn State Extension grant to hold a workshop in Orlando, Florida, for the discussion of this manuscript and others in the series. We thank Dana Johnson and Brianna Davis for help in organizing the references and the following individuals for reading and editing the manuscript before it was submitted: Fallon Fowler, Bethia King, David Taylor, Jerome Hogsette, Roger Moon, and Kateryn Rochon. Mention of trade names or commercial products in this publication is solely for the purpose of providing specific information and does not imply recommendation or endorsement by the U.S. Department of Agriculture. USDA is an equal opportunity provider and employer.