Use of Pupal Parasitoids as Biological Control Agents of Filth Flies on Equine Facilities

Abstract

Filth flies, including house flies, Musca domestica L., and stable flies, Stomoxys calcitrans (L.) (Diptera: Muscidae), are common pests on equine farms. The use of pupal parasitoids as biological control agents for filth flies is becoming more common on equine farms; however, there is a lack of information on the execution of augmentation programs for these farms. This review of biological control of filth fly pests on equine farms provides an overview of the life history and identification of filth fly pests and common commercially available parasitoids. Additionally, recommendations for use of pupal parasitoids based on known literature are provided, and the importance of continued research in this area is highlighted. When coupled with cultural control practices and other manure management techniques, pupal parasitoids offer an environmentally sound option for mitigating on-farm fly breeding.

House flies, Musca domestica L., and stable flies, Stomoxys calcitrans (L.), belong to the group collectively called filth flies and are common pests on equine farms. These flies can be present in high numbers on farms, breeding in equine manure and decomposing organic matter such as hay and wood shavings (Pitzer et al. 2011a, Machtinger and Geden 2013). Filth flies are associated with numerous pathogens that cause disease in humans and animals including shigellosis, cholera, and poliomyelitis (Greenberg 1973, Geden 1997, Graczyk et al. 2001). Fecal and other pathogenic bacteria, such as Escherichia coli and Salmonella spp., acquired by flies can be transmitted to humans (Moriya et al. 1999, Mian et al. 2002). House flies are mechanical vectors of the protozoan pathogens, Cryptosporidium spp. and Giardia spp. (Conn et al. 2007), and the bacterium, Corynebacterium pseudotuberculosis, which causes pigeon fever in horses (Spier et al. 2004, Barba et al. 2015). Nematodes in the genus, Habronema, can cause progressive weight loss, ulcers, colic, and skin disorders and are transmitted by both house flies and stable flies (Naem 2007). Transmission of equine infectious anemia virus can result from stable fly feeding (Greenberg 1971), and arthropod hypersensitivity and pruritus (itching) in horses also has been linked to stable fly feeding (Gortel 1998). To combat these potential medical risks, an estimated US$40 million was spent in 1997 in the United States for ectoparasite control on horses (Hinkle et al. 2001). Numerous options for fly control, including biological control products, are marketed to horse owners to reduce health risks associated with filth flies.

Pupal parasitoids (Hymenoptera: Pteromalidae) are the most common biological control agents used on equine facilities for filth fly management and their use is increasing (United States Department of Agriculture [USDA] 2006, Machtinger et al. 2013). Naturally occurring populations of parasitoids typically are insufficient at suppressing the large populations of filth flies produced from the attractive and abundant development material, such as manure and soiled bedding, generated on equine farms. However, augmentation of natural populations with commercially produced parasitoids may be effective in suppressing fly populations when coupled with other management methods. Several commercial insectaries sell parasitoids to manage pest fly populations (Leppla and Johnson 2011), but instructions for use vary among vendors.

In 2005, the American Horse Council found that the equine industry contributed US$39 billion directly to the national economy each year and US$102 billion per annum when industry suppliers and employees were taken into account (American Horse Council Foundation [AHC] 2005). Despite the importance of the equine industry, pest control research and integrated pest management (IPM) extension delivery to this sector have been limited. Surprisingly little is known about pupal parasitoid associations with equine farms (Hogsette 1981, Pitzer et al. 2011b, Machtinger 2011), and no projects evaluating the efficacy of pupal parasitoids as a control method for filth flies on these facilities have been completed.

The purpose of this review is to consolidate known information, highlight areas in need of new research, and assess the use of pupal parasitoids as biological control agents of filth flies on equine facilities. The basic biology of the house fly, stable fly, and the parasitoids commonly or potentially used to manage them is reviewed, as are methods to identify appropriate situations for use of biological control agents. Suggested guidelines are detailed for planning an IPM program, obtaining the parasitoids, and monitoring parasitoid effectiveness.

IPM for Filth Fly Control

IPM focuses on a long-term approach to pest management. IPM programs are based on methodologies that incorporate the application of a diverse set of techniques to reduce insect pest populations. The goal is to implement the best combination of pest management strategies that reduce economic, health, and environmental risks while maximizing pest control.

An IPM program involving cultural control, mechanical control, and biological control has the potential to significantly reduce reliance on chemical insecticides for management of filth flies. Chemical treatments may reduce flies to acceptable numbers temporarily on some farms; however, the increasing cost, toxicity to nontarget species, environmental pollution, and increasing insecticide resistance in the target pest preclude their extended sole use (Pickens and Miller 1987, Cilek and Greene 1994, Kočišová et al. 2002, Marçon et al. 2003, Malik et al. 2007, Scott et al. 2013). House flies can rapidly develop resistance to common insecticides (Liu and Yue 2000, Kaufman et al. 2010), and stable flies have become tolerant of some chemical formulations (Malik et al. 2007, Memmi 2010, Pitzer et al. 2010). Automatic insect control systems found in many horse barns likely contribute to selection pressure leading to high levels of resistance in filth flies. In a poultry facility, resistance to pyrethrins plus piperonyl butoxide was considerably higher in filth flies exposed to insecticides at frequent intervals (Meyer et al. 1990a). While insecticide sprays for topical application to horses are compatible with pupal parasitoid use, automatic spray systems that deliver insecticide to fly breeding areas are not and may interfere with the effectiveness of biological control programs (Scott et al. 1991, Geden et al. 1992a).

Successful IPM programs include cyclical elements of pest detection and identification, monitoring of abundance, use of established threshold levels, selection and application of the best management methods, and evaluation of results. These steps are important to address before initiation of a biological control program using pupal parasitoids.

Filth Fly Identification and Life History

Proper identification of pests is necessary to avoid releasing biological control agents in a situation where they will not be effective. Pteromalid parasitoids will parasitize primarily house flies and stable flies, but not horse flies, deer flies, mosquitoes, gnats, or other common pests in equine facilities (Machtinger et al. 2013). Horn flies are a common pests in equine facilities located near pastured cattle in some areas of the United States. However, horn flies exclusively lay their eggs and develop in cattle manure and are thus beyond the reach of on-farm parasitoid use for horse owners.

Understanding the life cycle of the pest and identifying potential filth fly breeding habitats is important before purchasing and releasing pupal parasitoids. Parasitoids will not control pests already in the adult stage; therefore, identifying immature filth fly habitats will help in assessing if the use of pupal parasitoids would be effective or if other control practices are required. If adult flies are observed on the farm, it must first be determined if the flies are developing on site or moving in from surrounding areas. If flies are not actively developing on the farm and other livestock facilities are in the vicinity, it is likely adults are moving to the affected horse farm from neighboring farms. If flies are developing on the affected horse farm, those sites should be assessed for ways to improve cultural and mechanical control practices in conjunction with parasitoid releases. Sanitation remains the most economical and effective way to manage fly pests developing on site, but parasitoids can help with areas of the farm that cannot be effectively cleaned. Additionally, locating fly breeding areas will help to determine appropriate places for parasitoid releases.

House flies and stable flies are both members of the family Muscidae within the insect order Diptera (“true flies”). Both have complete metamorphosis with four distinct life stages—egg, larva, pupa, and adult (Fig. 1a). The larvae of both species are elongated and white or yellow-white with dark mouth hook. Larvae are usually found in aggregations in suitable substrates such as excrement-soiled hay, straw, pine shavings, and other bedding materials. Flies pupate once larval development is complete. Pupae are about 8 mm long, brownish-red, and resemble small pellets with a barrel or capsule shape. Under typical summer conditions, development time from egg to adult can occur in as little as 10 d for house flies (Hinkle et al. 2001) and 14 d for stable flies (Larsen and Thompson 1940). The short generation time and capability of females to lay hundreds of eggs in their lifetime can lead to rapid increases in fly numbers on equine farms in warmer months.

Fig. 1.

(a) Life cycle of the house and stable fly. The life cycle of house flies (M. domestica; pictured) and stable fly (S. calcitrans) from egg (egg mass shown) to larvae, pupae, and adult is pictured. Photo courtesy Lyle Buss and Jane Medley, University of Florida. Comparison of (b) house fly and (c) stable fly adults. Adults of both species are tan and grey with four black vertical stripes on the dorsal area of the thorax. The stable fly has a checkerboard pattern of dark spots on the dorsal surface of the abdomen and a lighter tan or grey spot between the black stripes on the thorax dorsum.

House flies and stable flies are superficially similar in appearance, but have several distinguishing characteristics (Fig. 1b and c). Adults of both species are tan and grey with four black vertical stripes on the dorsal area of the thorax. However, the stable fly has a checkerboard pattern of dark spots on the dorsal surface of the abdomen and a lighter tan or grey spot between the black stripes on the thorax dorsum. These two flies also can be distinguished by their mouthparts and where they are found feeding and resting. House flies have downward projecting, sponging mouthparts. To feed, house flies regurgitate enzyme-containing saliva onto food before ingesting it in a liquid form. Additionally, house flies often defecate where they feed, which facilitates pathogen transmission. They are attracted to mucus, sputum, and moist areas around the eyes, nostrils, and wounds. In contrast, stable flies are obligate blood feeders and have a rigid piercing proboscis that projects forward from the head. This species most often feeds on the lower portions of animals, such as the legs of horses. Once a stable fly has taken a bloodmeal, it leaves the animal, typically resting on nearby fence lines or other structures. Horse owners sometimes confuse these flies with the horn fly, Haematobia irritans (L.), which feeds on the back or belly of horses. However, horn flies are substantially smaller and have only two black thoracic stripes. Although horn flies may feed on horses, their larvae develop only in bovine manure. Unless a horse owner also keeps cattle on the property, these flies are produced off-site and management options are limited to control adults. Fly identification can be confirmed by submitting samples to local Cooperative Extension educators.

Adult female filth flies seek appropriate media for oviposition, which may include manure piles, bedding in stalls, soiled hay, and fresh manure (Fig. 2). House flies develop consistently in pure manure (Meyer and Shultz 1990, Broce and Haas 1999, Larraín and Salas 2008) and, in choice tests with various equine-associated substrates, they preferred fresh horse manure and manure mixed with pine shavings (Machtinger et al. 2014). Stable flies prefer decomposing plant matter mixed with animal wastes, including improperly composted material, and material and debris around large rolled hay bales (Hall et al. 1982, Meyer and Peterson 1983, Meyer and Shultz 1990, Skoda and Thomas 1993, Broce et al. 2005, Talley et al. 2009). Stable flies are more attracted to equine manure for oviposition when compared against bovine manure (Jeanbourquin and Guerin 2007). In choice studies, stable flies were primarily attracted to equine manure mixed with pine shavings. This species was flexible in oviposition and development on common substrates found on equine farms (Machtinger et al. 2014), although stable flies preferred aged horse manure, 1–3 wk old, over fresh manure (Albuquerque and Zurek 2014).

Fig. 2.

Typical locations of fly development on equine farms. (a) Areas of accumulated manure, such as manure piles. Typically, fly development proceeds along the perimeter in fresher material. (b) Around the perimeter or between stall mats in horse stalls. Shavings and straw are bedding materials that promote fly development. (c) Old hay surrounding round hay bales. Horses defecate and urinate on this waste hay creating optimal environments for stable fly development. (d) Isolated manure either in wheelbarrows, manure buckets, or run-in sheds is ideal for house fly development (Photos courtesy Erika Machtinger).

Both house flies and stable flies are abundant on equine farms and their population size is affected by temperature and moisture. In general, fly populations begin to increase in the spring when temperatures rise and then decline in the autumn as weather becomes too cool for fly development. In the southeastern United States, house flies are most common in the summer months (LaBrecque et al. 1972, Machtinger 2011, Pitzer et al. 2011b), whereas stable flies are usually active in the warmer periods of winter and spring (Simmons 1944, Machtinger 2011, Pitzer et al. 2011b).

Establishing a Pest Monitoring Program

A monitoring program to evaluate fluctuations in pest fly populations should be established to guide decisions on when to conduct additional pest management strategies, as well as to evaluate the effectiveness of the pest management program. Monitoring records can be saved and used to anticipate fly population increases in subsequent years. Methods for monitoring larvae and pupae are labor intensive and require a greater degree of knowledge to interpret appropriately, but adult fly monitoring can provide the necessary relative numbers over time to draw conclusions on the effectiveness of the management program.

House flies and stable flies have different behaviors as adults; therefore, monitoring strategies differ for each species. A number of monitoring methods are available for horse owners to evaluate relative adult house fly and stable fly abundance over time (Kaufman et al. 2000). Lysyk and Axtell (1985) found that house fly catches in baited jug traps and spot cards change proportionally with changes in house fly density. Spot cards are impractical for outdoor situations, but can easily be used in a barn (Rutz et al. 1992, Geden 2005). The sticky alsynite-based Williams trap, and subsequent modifications, captures large numbers of stable flies provided that they are properly serviced to avoid accumulation of debris on the sticky material (Williams 1973, Ruff 1979, Patterson 1981, Gersabeck et al. 1982, Rugg 1982, Hogsette 1983, Broce 1988).

For both flies, numbers of flies or spots should be recorded weekly. Definitive data on the numbers of flies that cause damage do not exist for horses. Horse owners are encouraged to record both fly numbers and animal behaviors to develop farm-specific fly thresholds. By examining these data over several fly seasons, horse owners can determine fly levels that are problematic and take corrective measures before these fly thresholds are reached.

Cultural and Mechanical Control

Manure management to reduce areas suitable for fly development is an integral component to ensure success in any fly IPM program, especially when using pupal parasitoids. Cultural control was found to be the most important method for on-site reductions of stable flies (Greene 1983), and Kaufman et al. (2005) found lower house fly captures in farms with the best sanitation practices. Currently, there is a need to educate equine farm owners and operators on how best to manage facilities to minimize fly abundance and maximize the use of biological control agents (Machtinger et al. 2013).

Mechanical control practices can reduce fly abundance by breaking up existing fly breeding areas or modifying the area such that adult flies will not oviposit in the habitat. A manure storage area that is dry and well ventilated is critical to fly management, as manure moisture promotes filth fly development (Watson et al. 1998). Covering manure piles with burlap or tarps to increase the temperature was shown to prevent exposure to pests and made the substrate unsuitable for filth fly development (Fay 1939). Correctly composting manure increases the internal temperature of the waste and lowers moisture content, rendering the substrate unsuitable for filth fly development (Abu-Rayyan et al. 2010). An alternative to stacking manure is spreading it over a pasture. Spreading a thin layer of manure on agricultural fields encouraged drying and reduced fly development (Axtell 1986). For pastured horses, breaking manure piles with a drag also facilitates quick drying.

Other organic matter, such as soiled hay and shavings, can facilitate developing flies, but can be modified easily. Waste hay produced from rolled round hay bales can be a major site of stable fly development (Broce et al. 2005, Talley et al. 2009). Burning or stacking aging hay bales and hay waste, placing hay on a mobile wagon, and frequently relocating round bale feeding sites may reduce house fly and stable fly breeding areas (Broce et al. 2005). Sodium bisulfate has been found to be a safe additive for horse stalls that acidifies the substrate thus reducing larval house fly development (Sweeney et al. 2000a,b; Calvo et al. 2010). Both low (2.3 kg/9.3 m²) and high (4.5 kg/9.3 m²) concentrations of sodium bisulfate added to stalls weekly reduced fly populations.

Additionally, using alternative bedding in horse stalls can reduce fly development. Schmidtmann (1991) found that, of the bedding types tested in calf hutches, sawdust and gravel bedding were the most effective at reducing fly numbers. Machtinger et al. (2014) found that both house flies and stable flies laid fewer eggs and had less successful development in horse manure mixed with sand or soil, while the most attractive substrate was horse manure mixed with pine shavings.

Trapping adult flies is complementary with biological control and is recommended to improve pest management effectiveness. Biological control of filth flies specifically targets the pupal stage of flies developing on the farm; however, farms often are located near other livestock and thus may experience adult fly pressure from populations immigrating from neighboring facilities. Adult flies from neighboring farms will not be controlled by the localized use of pupal parasitoids, but traps can help to lower adult fly numbers to more acceptable levels. At least 14 types of baited house fly traps were described in the literature between 1900 and 1995 (Pickens 1995), and there are many products on the market today. Baited traps, such as the Fly Terminator or Captivator traps (Farnam Products, Inc. Phoeniz, AZ), are common and effective for adult house flies (Geden et al. 2009). Traps targeting adult stable flies are more limited in number than those developed for house flies. However, the alsynite trap first developed by Williams (1973) proved effective at capturing stable flies. This trap was modified into a cylinder-shaped device (Broce 1988). A subsequent commercial version of the sticky trap is available from Olson Products (Medina, OH) with a smaller commercial trap, the Knight Stick, available through BugJammer, Inc. (Pennington, NJ). Ultra-violet light traps are available and used commercially for adult house fly and other nuisance pest control.

Biological Control

Augmentative biological control of filth flies has become a commonly used technique for fly control on equine facilities (USDA 2006, Machtinger et al. 2013). Given suitable habitats and hosts, commercially available pupal parasitoids have the potential to suppress populations of filth flies. The effectiveness of augmentation programs has not been evaluated on equine farms. However, in other livestock facilities, studies with various species have yielded mixed results, with suppression of fly populations with augmentative releases of parasitoids in some situations (Morgan and Patterson 1990; Geden et al. 1992b; Petersen and Cawthra 1995; Crespo et al. 1998, 2002; Skovgård and Nachman 2004; Geden and Hogsette 2006) and failures in others (Morgan 1980, Meyer et al. 1990b, Andress and Campbell 1994, Weinzierl and Jones 1998, McKay and Galloway 1999, Kaufman et al. 2001, 2002). These contradictory results may be partially attributed to environmental factors that affected parasitoid abundance and distribution (Skovgård 2004). Site-specific environmental factors include sensitivity to insecticides, use of low-quality commercial colonies, microhabitat preferences, availability of hosts, and a lack of optimal timing and methods of release (Petersen and Meyer 1985, Patterson and Rutz 1986). Additionally, it is possible that existing active parasitoid populations may inhibit the establishment or function of a newly released species in augmentation programs (Legner et al. 1990). Quarles (2006) suggested that the success of a biological control program using pupal parasitoids relies on matching the released species with the climate and habitat of the release location, most appropriately by deploying endemic species.

Successful use of biological control agents requires an integrated approach. Even if a biological control program is successful initially, immigrating flies from neighboring livestock areas can increase fly populations rapidly (Quarles 2006). An understanding of house fly and stable fly biology and species-appropriate monitoring and management methods are compulsory for equine facility owners to effectively develop and implement a successful IPM program for filth flies. Prior to the pest season, consideration should be given to select a parasitoid species for release that is compatible with the region and facility type and to locate an insectary that can supply that species. Finally, decisions should be made on handling parasitoids after shipment and frequency and times of release.

Adult pteromalid parasitoids used in biological control are tiny wasps that are black, metallic green, or bronze. These wasps are harmless to humans and horses and will not bite or sting. Adult parasitoids drill through the fly puparium and deposit one or more eggs on the surface of the fly pupa located within, depending on the species of wasp. The developing immature wasps kill the host fly as they develop, preventing the fly from emerging as an adult (Fig. 3). Development time of the parasitoid at constant temperature depends on the wasp species, ranging from less than 2 to over 4 wk before emerging as an adult.

Fig. 3.

Life cycle of a pteromalid pupal parasitoid of filth flies. Adult flies (top left) lay eggs that develop into larvae and then pupae. Female parasitoid wasps lay one or more eggs in the fly puparia (center). The immature wasps feed on the developing fly pupa, killing the fly and effectively breaking the fly life cycle. Adult parasitoids emerge from the fly pupa (right) to reproduce and continue the cycle (Drawing courtesy Erika Machtinger).

Several naturally occurring pupal parasitoids are found throughout the United States, though limited research has been conducted on parasitoid species dominance and population dynamics on equine farms. In Florida, Spalangia spp. are the most common parasitoids, comprising over 90% of species recovered from equine farms (Hogsette 1981, Machtinger 2011, Pitzer et al. 2011b). Spalangia cameroni Perkins was found to dominate parasitoid recoveries on equine farms, making up over 70 and 86% of species recovered on large and small equine farms, respectively (Machtinger 2011, Pitzer et al. 2011b). Similarly, in North Carolina, S. cameroni made up the majority of parasitoids recovered from equine farms (D. W. Watson, unpublished data). Muscidifurax raptor Girault and Sanders also has been found in limited numbers in equine farms, but is recovered more frequently in other livestock facilities.

Six species of filth fly pupal parasitoids from three genera are commonly sold commercially, each with slightly different life history characteristics. Limited research has been conducted on behavior, dispersal, and monitoring strategies for these common parasitoids. However, life history characteristics and competition have been investigated more thoroughly in the six pteromalid parasitoids presented below than for other parasitoids in the family that parasitize filth breeding flies.

A considerable amount of research has been conducted on the life history and effectiveness of M. raptor, a very common pupal parasitoid of filth flies. This is a solitary species (i.e., one wasp offspring produced per fly pupa) and, although highly dependent on temperature, lifetime fecundity has been reported to range from 69 to 211 progeny (Morgan et al. 1989, Geden et al. 1992c, Lysyk 2001a). In the field, this species has been found to be more effective at parasitizing hosts than Muscidifurax zaraptor Kogan and Legner (Floate 2002). For all species of parasitoids, development is temperature dependent, but at 25°C, M. raptor males emerge as adults in ∼17 d and females emerge in 19 d (Geden 1997). Dispersal of M. raptor following release has been found to be <30 m from the point of release on dairy farms (Smith et al. 1989). This species searches for host fly pupae in the top 3 cm or less of the host development substrate (Rueda and Axtell 1985, Geden 2002). Microhabitat preferences in equine-associated substrates are not known; however, in dairies this species was found more often around drainage areas and edges than in shavings in calf hutches (Olbrich and King 2003). In the laboratory, this species was found to prefer bovine manure over equine manure in olfactometer experiments (Machtinger and Geden 2015). M. raptor was found primarily outdoors in spilled feed, straw, and manure piles (Smith and Rutz 1991, Skovgård and Jespersen 2000) and rarely in poorly lit areas (Skovgård and Jespersen 2000), although a complete assessment of diel activity is not available.

M. zaraptor is similar in biology to M. raptor and is perhaps the most common commercially produced pteromalid parasitoid. Coats (1976) found that this species had a lifetime fecundity of ∼265 offspring and was highly competitive against other parasitoids. Adult M. zaraptor emerge in ∼18 d, with females emerging slightly later than males (Lysyk 2001b). This species was found to disperse no more than 8 m from a release point by Pawson and Petersen (1988). As with most Muscidifurax spp., searching in host development substrates is limited to just under the surface (Legner 1977). Some strains may search deeper. Microhabitat associations of M. zaraptor have not been explored in equine or other livestock facilities. However, like M. raptor, this species was found to prefer bovine manure over equine manure in the laboratory (Machtinger and Geden 2015). Mullens et al. (1986) reported that this species is exclusively active during daylight hours. M. zaraptor is frequently recovered in western U.S. states, whereas M. raptor is more common in eastern states.

Muscidifurax raptorellus Kogan and Legner (Fig. 4a) is gregarious (i.e., multiple offspring produced per pupa) and prolific. Total lifetime fecundity ranges from 85 to 166 progeny at 25°C and adults of both sexes emerge at ∼16 d (Petersen and Currey 1996, Lysyk 2001c). Dispersal after release seems to be limited in some situations. Parasitism by M. raptorellus was not observed greater than 6 m away from a release point in a high rise poultry building, regardless of release rate (Tobin and Pitts 1999). Further distances of 22.5 m and 48 m were observed in outdoor cattle systems by Lysyk (1995) and Petersen and Cawthra (1995), respectively. The competitive searching ability of this species has been investigated minimally, but it was found to be ineffective at locating and parasitizing pupae in soiled shavings from an equine farm under laboratory conditions (Pitzer et al. 2011a). The microhabitat and circadian activity preferences of M. raptorellus on equine or other livestock facilities have not been investigated.

Fig. 4.

Female pteromalid parasitoids (a) Muscidifurax raptorellus Girault and Sanders parasitizing a house fly pupa. Photo courtesy Lyle Buss, University of Florida. (b) Spalangia cameroni Perkins on a house fly pupa. Photo courtesy Lyle Buss, University of Florida. (c) Nasonia vitripennis (Walker) on a Sarcophagidae spp. Photo courtesy Lyle Buss, University of Florida.

Spalangia cameroni (Fig. 4b) is a solitary parasitoid with an average total lifetime fecundity ranging from 38 to 52 progeny per female (Morgan et al. 1989, Machtinger 2015). Adult emergence occurred at ∼27 d, with males emerging slightly earlier than females (Geden 1997). Dispersal was found to be <10 m from a release point in indoor swine facilities (Skovgård 2006), and similarly in outdoor situations, linear dispersal was generally <10 m (Machtinger et al. 2015a). Under interspecific competition, female S. cameroni will search deeper for hosts in host development substrates than Muscidifurax spp., with depths of up to 10 cm reported (Rueda and Axtell 1985, Geden 2002). S. cameroni was found to be effective at locating and parasitizing house fly hosts that had pupated in soiled shavings from an equine farm (Pitzer et al. 2011a) as well as other common equine-generated substrates such as horse manure, hay soiled with horse manure, and sand mixed with manure (Machtinger and Geden 2013). In addition, S. cameroni was more attracted to equine manure containing house fly larvae over pupae (Machtinger et al. 2015b) and to odors from horse manure versus bovine manure in the laboratory (Machtinger and Geden 2015). These laboratory data support the limited field work on equine farms showing Spalangia as the dominant genus on equine farms in Florida (J.A. Hogsette, unpublished data, Hogsette 1981) and North Carolina and, specifically, S. cameroni as the most frequently collected species (Machtinger 2011, Pitzer et al. 2011b, D. W. Watson, unpublished data). Preferred microhabitats on equine farms have not been evaluated, but on dairies S. cameroni was recovered primarily indoors and preferred loose substrate, such as straw and calf bedding (Smith and Rutz 1991). Skovgård and Jespersen (2000) corroborated these findings and reported S. cameroni most frequently in indoor pig farms and cattle farms. S. cameroni was found to prefer calf hutches with shavings on dairies in Illinois (Olbrich and King 2003). Preferred moisture levels of substrates were 45–65% in poultry manure, but when hosts were limited this species was somewhat flexible, parasitizing hosts in substrates with up to 75% moisture (Geden 1999). Mullens et al. (1986) reported that S. cameroni was active during daylight hours and to some degree at night.

Spalangia endius Walker is a solitary parasitoid. Lifetime fecundity has been found to be between 10 and 40 progeny per female (Ables and Shepard 1974, Morgan et al. 1976), with an average adult life span of <4 d (Morgan et al. 1976). Adult emergence occurs in ∼19 d at 26°C (de Araujo et al. 2012). Dispersal information on this species is unknown. S. endius will search uniformly for hosts throughout the substrate, but does parasitize at depths of 5–10 cm (Rueda and Axtell 1985, Geden 2002), much deeper than Muscidifurax spp. Additionally, S. endius was found to be as successful as S. cameroni at locating and parasitizing hosts in pine shavings soiled with equine waste (Pitzer et al. 2011a). In the laboratory, S. endius preferred dry poultry manure at about 45% moisture, although it would search at broader moisture levels when hosts were limited (Geden 1999). This species, like S. cameroni, had a preference for equine manure over bovine manure in the laboratory (Machtinger and Geden 2015). Other microhabitat preferences and circadian activity have not been evaluated on equine or other livestock facilities for this species.

Nasonia vitripennis (Walker) (Fig. 4c) is gregarious, widely distributed in North America, and easy to rear. Although sold commercially, this species is a generalist that is most commonly found in carrion and calliphorid systems (Peters 2010). Despite its inexpensive cost to consumers, and aggressive marketing to equine owners and managers, purchasing N. vitripennis is not recommended for use in filth fly biological control programs on equine farms due to disappointing results in field trials (McKay and Galloway 1999, Kaufman et al. 2001). The complete genome of this species has been sequenced, and this will provide invaluable tools for future work with the more economically important Muscidifurax and Spalangia relatives (Werren et al. 2010).

Suggestions for Implementing a Biological Control Program With Pupal Parasitoids on Equine Farms

Biological control using pupal parasitoids has been used in livestock facilities for many years. Husbandry practices in equine farms differ from cattle, swine, and poultry facilities where the majority of research on pupal parasitoid use has been conducted. The filth fly and parasitoid system is less known in equine farms, and there are few guidelines on the best execution of parasitoid release in these facilities. While some recommendations are universal, others are based on information from use in other livestock facilities and what is known about parasitoid life history and preferences.

Deciding When, How Often, and How Many Parasitoids Should be Released

Because pupal parasitoids work better to prevent unacceptable population levels as opposed to a curative control method, it is important to anticipate fly problems using the biology of the fly species and monitoring records that indicate when fly populations might increase. Releases of parasitoids should begin before pests become a problem. This information can be collected from previous monitoring data, or from generalizations of population increases regionally. Fly flight, mating, and oviposition begin at ∼10°C and increase in activity as temperature increases (Keiding 1986). Filth flies are found on livestock facilities from spring to autumn in most areas of the country and year-round in the very southern United States. Depending on the geographic location of the facility, releases may need to start in the winter or early spring in the South and mid-spring in cooler regions as temperatures begin to increase, although releases can occur at any time. Numbers of released parasitoids can be adjusted based on fly monitoring results and will be driven largely by the availability of fly breeding habitats.

The number of parasitoids to release typically is expressed as a number of parasitoids per unit, either by animal number or by acre. There is limited empirical evidence to support specific release rates for the many types of livestock facilities, as they are managed in so many different ways and sanitation can vary dramatically. Recommended releases at cattle farms range from 200 parasitoids per cow per week to upwards of 500 to 2,000 per calf, where calves are the driving force of fly breeding sites (Rutz et al. 1992, Kaufman et al. 2012). Unfortunately, there are no data available to provide recommendations for parasitoid releases on equine facilities. However, suppliers have suggested a release rate of 2,000 parasitized pupae per stalled horse on a biweekly basis as a starting point. The use of farm acreage to determine parasitoid purchase volume is not recommended, as parasitoid need is much more related to the number of waste-producing animals than farm size (which may have few or many horses per acre).

Most commercial insectaries prefer to automatically ship their products every 4 wk for equine clientele. Monthly releases may be convenient, but weekly or biweekly releases are considerably more likely to be effective in light of the life history differences between the parasitoid and the fly targets. Horse owners are encouraged to survey several insectaries or distributors to find the best place to purchase parasitoids that match their price and delivery expectations.

Purchasing Pupal Parasitoids

Pupal parasitoids can be purchased from several commercial insectaries or distributors. Buyers should be cautious and informed when selecting a supplier and ask relevant questions before ordering. These companies are competitive and differ in the services they provide as well as the products they sell. Buyers should ask if the supplier offers professional consulting services to follow up with the buyer on the success or failure of the program. The buyer should know if the supplier produces their own product or if they are a distributor of a product. Additionally, the buyer should be aware of which species of parasitoid is being supplied and whether a qualified entomologist or taxonomist made that species determination. The Latin binomial species name should be given for clarity (i.e., Spalangia cameroni is different from Spalangia endius, but both could be referred to as Spalangia spp.). The supplier should be asked how often the parasitoids are checked for quality and, if the product contains a mix of parasitoid species, what is the species ratio. The number or percentage of expected females in each order should be provided as well. The equine owner should compare answers from the various suppliers and choose the supplier that best meets his/her needs.

Buyers should consider the conditions for parasitoid shipment. The product should be sent with minimal travel time, preferably in an insulated container to protect the parasitoids from temperature extremes. Instructions for handling and release should be provided by the supplier, including information on how soon the parasitoids are expected to emerge. Buyers should inquire what options are available for quality determination. Complications from shipping, colony quality, or other unforeseen circumstances can occur, even with reliable and knowledgeable suppliers. The consumer should implement a quality assurance protocol to determine the condition of the parasitoids received. A simple protocol to verify quality involves collecting and holding a small quantity, ∼100, of the pupae from each shipment in a small container, such as an old medicine bottle. Record the date of the shipment and leave the container in a climate-controlled location out of direct sunlight for 3–4 wk. Pupae can be assessed for parasitoid emergence after this holding time by counting emergence holes (little round holes) in the pupae that indicate each fly puparium produced a live parasitoid (Fig. 5). The holes are easily seen with a magnifying glass and good lighting. Vendors should adjust the number of pupae shipped to account for actual percent parasitism, but an end-user quality check will document if there are problems with mortality during shipping. If pupae are being placed in stations, not scattered, pupae from the station can be examined for emergence holes after the same time frame and compared with the samples held in climate-controlled conditions. This can be a useful check to make sure that the pupae are not being placed where environmental conditions are harmful for parasitoid emergence.

Fig. 5.

Adult pteromalid pupal parasitoid emergence holes in house fly pupae.

Handling Parasitoids

Pupal parasitoids are small, living insects that require careful handling to avoid mortality. Parasitoids usually arrive in small bags with the purchased number of house fly pupae mixed with shavings. Prior to shipment, pupae were parasitized and will produce only adult wasps. Once received, specific instructions for handling the parasitoids provided by the company should be followed carefully. Parasitoids should be kept in the packaging they were received in under regulated temperatures and out of direct sunlight until they are released.

Timing and Locations of Release

A critical component of a biological control program is the timing and location of the release of the control agents. It is suggested that releases be made during the morning or late afternoon to avoid high temperatures that can increase mortality. Other weather conditions, like rain and wind, can have negative impacts on survival, and releases during inclement weather should be avoided unless provisions are made for providing shelter for the pupae.

Unfortunately, there are limited data on the most effective way to release parasitoids of filth flies. Research suggests that parasitoids are fairly limited in their dispersal from a release point (see information on commercially available species in above section). Some suppliers suggest sprinkling the parasitized pupae on the ground in areas where fly development is likely or previously confirmed, such as manure piles, horse stalls, feeding and watering stations, or near aging hay (i.e., round bales). It is important to keep in mind that locations where immature flies develop often are quite different from where the most adult fly activity is observed. Releases should be made on the perimeter of these areas and preferably in locations protected from horse and human traffic. Some suppliers recommend releasing parasitoids in hanging tubes or mesh bags in the barn or in a shaded, outdoor location. Both of these methods require careful consideration to avoid loss from predation. If ants, mice, chickens, and wild birds are a concern, buyers should work with the supplier to develop a release plan that is suitable for their situation.

Conclusions

Although pupal parasitoids of filth flies are sold commercially and aggressively marketed in the equine industry, little is known about their basic biology in this system. There is a strong stakeholder interest in biological control, with over 80% of respondents in a 2012 survey requesting that more research be conducted on biological control of filth flies (Machtinger et al. 2013). Despite this interest, the effectiveness of pupal paSrasitoids for fly management has not been evaluated on equine farms and there is a pressing need to fill the knowledge gap on use of parasitoids on these facilities. With increased support for research directed at the equine industry, further improvements to recommendations for parasitoid use can be made.

Biological control for filth flies using pupal parasitoids has the potential to be effective in reducing fly populations on equine farms, as has been shown in other livestock systems. It is important to remember that the use of these biological control agents is not a quick fix to a fly outbreak, but careful consideration must be taken to ensure the releases are being made in the appropriate situations and where cultural control methods for manure management are practiced. With careful planning, horse owners can use pupal parasitoids as an environmentally-sound method to reduce pest house fly and stable fly numbers and reduce the risks to horses and their owners. Owners are encouraged to share their experiences with other owners, parasitoid suppliers, Extension personnel, and their veterinarians so that the community can benefit and learn together how to best incorporate biological control into practical management solutions.

Acknowledgments

The authors would like to thank Drs. Erica Lacher and Nancy Hinkle for their extensive reviews of this manuscript and helpful comments. This project was supported by a United States Department of Agriculture Southern Sustainable Agriculture and Research Education (SARE) grant (GS11-101).