Stable Fly Diptera MuscidaeBiology, Management

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Journal of Integrated Pest Management, (2021) 12(1): 38; 1–23
https://doi.org/10.1093/jipm/pmab029
Profile
Published by Oxford University Press on behalf of Entomological Society of America 2021. 
This work is written by (a) US Government employee(s) and is in the public domain in the US.
Stable Fly (Diptera: Muscidae)—Biology, Management, 
and Research Needs
K. Rochon,1,7, J. A. Hogsette,2, P. E. Kaufman,3, P. U. Olafson,4 S. L. Swiger,5 and 
D. B. Taylor6,
1Department of Entomology, University of Manitoba, 12 Dafoe Rd., Winnipeg, MB R3T 2N2, Canada, 2USDA-ARS, Center for Med-
ical, Agricultural and Veterinary Entomology, 1600 S.W. 23rd Drive, Gainesville, FL 32608, USA, 3Department of Entomology, Texas 
A & M University, College Station, TX 77843, USA, 4USDA-ARS, Knipling-Bushland US Livestock Insects Research Laboratory, 2700 
Frederickburg Rd., Kerrville, TX 78028, USA, 5Department of Entomology, Texas A&M AgriLife Research and Extension Center, 1229 
N US Hwy 281, Stephenville, TX 76401, USA, 6Department of Entomology, University of Nebraska, Lincoln, NE 68583, USA, and 
7Corresponding author, e-mail: kateryn.rochon@umanitoba.ca
Subject Editor: Matthew Messenger 
Received 21 June 2021; Editorial decision 13 August 2021 
Abstract
Stable flies, Stomoxys calcitrans (L.) are global pests of livestock, companion animals, and humans. These flies 
inflict painful bites and cause significant economic losses to producers by reducing livestock production. In addition, 
they have been associated with the mechanical transmission of several pathogens causing disease in animals. 
Management of this species is difficult because: 1) their developmental habitats are often ephemeral accumulations 
of decomposing vegetation, 2) they can exploit cultural practices in many agricultural and urban environments, and 
3) the adults are highly mobile. An integrated pest management (IPM) approach is required to effectively manage 
stable flies, including integration of cultural, mechanical, physical, biological, and chemical control options. The 
challenges of stable flies in different animal commodities are discussed, and current and novel technologies for 
control are presented. Lastly, need for additional research to improve stable fly management methods are discussed.
Key words: filth fly, animal production, Integrated Pest Management, livestock pest, Stomoxys calcitrans
Stable flies (Stomoxys calcitrans (L.) [Diptera: Muscidae]) are eco-
nomically significant blood-feeding pests of livestock, companion 
animals, and humans worldwide. Originating in Africa (Zumpt 
1973, Dsouli-Aymes et  al. 2011), stable flies accompanied human 
colonization and the introduction of livestock throughout the tem-
perate and tropical regions of the world (Zumpt 1973). With their 
painful bites, stable flies reduce livestock productivity and welfare, 
the comfort of companion animals, and interrupt recreational and 
other outdoor activities of humans (Hogsette et al. 1987). Although 
not considered important vectors of human or animal pathogens 
in North America, stable flies have been implicated in the trans-
mission of several animal pathogens elsewhere in the world, some 
of which are considered invasive with the potential to establish in 
North America. Stable flies are highly adaptable and exploit changes 
in climate and agronomic practices to produce explosive outbreaks 
threatening livestock viability in several regions of the world (Kunz 
and Monty 1976, Cook et  al. 2011, Dominghetti et  al. 2015, 
Solórzano et al. 2015). Historically, similar outbreaks have occurred 
in North America (Bishopp 1913a, Simmons and Dove 1941, 1942) 
and could re-emerge with changing climate and agronomic practices.
Despite the ubiquity and economic importance of stable flies, re-
views of their life history and biology are sparse. Bouché (1834) first 
reported finding immature stable flies in manure, but it was not until 
60 yr later that Newstead (1906) described their life history and 
illustrated the immature stages. In response to severe outbreaks of 
stable flies in 1912 in the central United States, Bishopp (1913b) pro-
vided a review of their biology, life history, and economic impacts. 
The taxonomy of the Stomoxyine flies was reviewed by Zumpt 
(1973) and the role of stable flies in the transmission of pathogens 
was reviewed by Baldacchino et al. (2013a). In this review, we will 
discuss advances in the knowledge of biology, life history, and eco-
nomic impacts of stable flies as well as needs for future research.
Description and Distribution
Superficially, adult stable flies are similar in appearance to house flies. In 
fact, a common name is “biting house fly”. Adult stable flies are 4–7 mm 
in length, gray in color, and can be distinguished by the dark reddish-
brown piercing–sucking proboscis, which extends anteriorly from the 
head, two pairs of broad dark thoracic stripes, and a tessellated black 
abdominal pattern (Zumpt 1973). Wing vein m1+2 bends forward very 
slightly and meets the costa just posterior to the wing apex (Fig. 1). This 
differs from house flies (Musca domestica L. [Diptera: Muscidae]) for 
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https://doi.org/10.1093/jipm/pmab029
https://orcid.org/0000-0002-1436-3692
https://orcid.org/0000-0001-7574-3608
https://orcid.org/0000-0001-6159-8358
https://orcid.org/0000-0002-4378-4867
mailto:kateryn.rochon@umanitoba.ca?subject=
2 Journal of Integrated Pest Management, 2021, Vol. XX, No. XX
which m1+2 bends sharply forward and meets the costa anterior to the 
wing apex. Adult stable flies rest with their head held higher than the ab-
domen, and wings partially spread, revealing a portion of the abdomen.
Larvae are white and vermiform with characteristic widely 
spaced, black, rounded triangular-shaped posterior spiracular plates 
with three yellowish serpentine slits (Fig. 2; Skidmore 1985, Friesen 
et al. 2015). Stable fly larvae are not as robust as those of house fly 
and are rarely found as densely clustered in a substrate.
Stable flies, along with horn flies (Haematobia irritans (L.) [Diptera: 
Muscidae]), are members of the tribe Stomoxyini, a monophyletic sister 
group of the Muscini within the muscid subfamily Muscinae (Schühli 
et al. 2007, Dsouli et al. 2011). There are 18 species within the genus 
Stomoxys, all tropical except Stomoxys calcitrans which are found 
worldwide, having spread from their native African range with the as-
sistance of humans and the introduction of European livestock (Zumpt 
1973). Genetic analyses using mitochondrial and nuclear markers iden-
tified two stable fly populations, one in southern Asia and a second in 
Africa (Dsouli-Aymes et al. 2011). The African population is the source 
for the worldwide expansion of stable flies during the past 500 yr. Stable 
flies have been reported from Europe, Africa, Asia, Australia, and the 
Americas. Within North America, population genetic studies based on 
nuclear and mitochondrial markers demonstrated high levels of gene 
flow among populations (Krafsur et  al. 1994, Szalanski et  al. 1996, 
Kneeland et al. 2013).
General Biology
Stable flies have four stages in their life cycle: egg, larva, pupa, and adult. 
Eggs hatch into small first instar (≈ 1 mm) 12–24 h after oviposition. 
First instars increase in size to ≈1.7 mm and molt into the larger second 
instars (≈ 2.8 mm). After additional growth, second instars molt and be-
come third instars (≈ 5.2–11.0 mm). Third instars form coarctate pupae. 
Development from egg to pupariation takes ≈ 12–13 d at 27°C (Larsen 
and Thomsen 1940, Lysyk 1998). Adults emerge after approximately 
seven more days (≈ 20 d total from egg to adult). Development time and 
size are dependent upon temperature (Lysyk 1998), substrate quality 
(Florez-Cuadros et  al. 2019), and larval density(Taylor et  al. 2007, 
Friesen et al. 2016, Skovgård and Nachman 2017).
Feeding
Adults of both sexes feed on blood and are persistent biters of 
cattle and other warm-blooded animals. Although they prefer 
feeding on livestock, stable flies can be serious pests of humans 
when livestock are not available (Jones et al. 1985, Hogsette et al. 
1987, Jones et  al. 1992, Ose and Hogsette 2014, Hogsette and 
Ose 2017). Stable flies feed preferentially on the lower legs of 
cattle, horses, other livestock, and humans but favor the ears of 
dogs (Farkas and Gyurcsó 2006) and other animals (Hogsette 
et al. 1987). However, when adult fly abundance is high, they will 
also feed elsewhere on the body. When feeding on livestock, stable 
flies usually orient with their head upwards (Bishopp 1913a; Fig. 
3). This contrasts with horn flies, which normally orient with 
the head downwards. Adults use sharp, sclerotized labial teeth 
to penetrate the host’s skin (Elzinga and Broce 1986), creating 
a pool of blood from which they feed. Stable flies consume ap-
proximately 12  µl of blood per meal (Venkatesh and Morrison 
1980, Salem et al. 2012a) in 2–4 min (Harris et al. 1974, Salem et al. 
2012a). Host defensive responses to the painful bites often interrupt 
feeding prior to repletion (Schofield and Torr 2002). Stable fly saliva 
contains no anesthetics, as indicated by sialome analyses (Wang et al. 
2009, Olafson et  al. 2021), making their bites more painful than 
those of many other blood-feeding insects. Stable flies are closely 
associated with their hosts only while blood feeding. Adults can take 
their first bloodmeal 12–24 h after emergence (Parr 1962). Feeding 
frequency is temperature-dependent, with flies feeding once per day 
Fig. 1. Adult female stable fly (top) with characteristic wing venation with 
slight m1+2 bend meeting with the costa posterior to the wing apex. The 
adult house fly (bottom) showing m1+2 with sharp bend and meeting with 
the costa anterior to the wing apex.
Fig. 2. Third instar stable fly larva (A). Posterior spiracles of immature stable 
fly (B) are shaped like rounded triangles and more widely spaced than those 
of the house fly (C).
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Journal of Integrated Pest Management, 2021, Vol. XX, No. XX 3
on cooler days (Berry and Campbell 1985) and twice per day on 
warmer days (Harris et al. 1974, Müller et al. 2012).
Adult stable flies feed on nectar and other sugar sources as well 
(Tseng et  al. 1983, Jones et  al. 1985, Jarzen and Hogsette 2008, 
Taylor and Berkebile 2008). At beach sites in Florida, 23% of fe-
males and 12% of males had sugars in their guts (Jones et al. 1985) 
whereas, in Nebraska, 13% of females and 11% of males had de-
tectable sugar (Taylor and Berkebile 2008). Flies fed sugar in the 
absence of blood lived 7–8 d, five times longer than those fed water 
alone, but less than the ≥ 13-day survival time of those provided with 
blood (Moobola and Cupp 1978, Jones et al. 1992). Furthermore, 
sugar alone cannot support sexual maturation or yolk deposition in 
the ovaries (Anderson 1978, Moobola and Cupp 1978, Jones et al. 
1985). Nectar-sated flies tend to not take bloodmeals (Tseng et al. 
1983, Jones et al. 1985, Jones et al. 1992, Müller et al. 2012).
Reproduction
The stable fly mating process begins when males are 1–2 d old and 
females are 3–4 d old (Harris et al. 1966). Males can inseminate mul-
tiple females but females only mate once (Harris et al. 1966). Adult 
male stable flies require at least one bloodmeal prior to copulation 
and females require 2–3 (Anderson 1978, Chia et al. 1982, Morrison 
et al. 1982). Females require 3–4 bloodmeals to develop each batch 
of eggs (Harris et  al. 1974, Jones et  al. 1999). Females have an 
average of ≈100 ovarian follicles and can produce ≈100 eggs per 
ovarian cycle (Scholl 1986, Beresford and Sutcliffe 2012). Females 
frequently move while depositing eggs in small numbers throughout 
suitable substrates (Bishopp 1913a, Thomsen and Hammer 1936). 
Approximately 50% of the females survive long enough to complete 
one ovarian cycle and 50% of those survive long enough to com-
plete a second cycle (Beresford and Sutcliffe 2012). In the laboratory, 
lifetime egg production can be as high as 600–800 eggs per female 
(Lysyk 1998).
Immature Development
Stable fly larvae feed and develop in fermenting vegetative mater-
ials, often contaminated with animal wastes (Simmons and Dove 
1941, 1942, Siverly and Schoof 1955, Hafez and Gamal-Eddin 1959, 
Sutherland 1978, Campbell and McNeal 1980, Williams et al. 1980, 
Hall et al. 1982, Cook et al. 1999, Solórzano et al. 2015, Cook et al. 
2018a). Stable fly larvae do not use undisturbed, fresh bovine feces in 
the field (Bishopp 1913a), although, in the laboratory, they can develop 
in fresh feces (Hammond 2015). Horse feces aged 1–3 wk are more at-
tractive to ovipositing female stable flies and provide better sustenance 
to the larvae than fresh or >3-wk-old horse feces (Albuquerque and 
Zurek 2014). Stable flies are dependent on the microbial community 
associated with suitable substrates and cannot develop in sterilized 
oligidic substrates (Lysyk et al. 1999, Romero et al. 2006).
Larval densities within substrates can be high with average 
densities of ≈ 1,500 stable flies/m2 collected in emergence traps 
(Taylor and Berkebile 2011) and up to ≈ 3,900 stable flies/m2 
emerging from core samples (Broce et al. 2005). Silage and rotting 
vegetables can support the development of > 2,000 flies/m2 (Williams 
et al. 1980, Cook et al. 2018a). Lower densities of 24–175 larvae/
m2 have been recorded in pineapple residues, sugarcane filter cake, 
and mulch mixed with vinasse (Solórzano et al. 2013, Dominghetti 
et al. 2015).
In winter hay feeding site substrates, the probability of finding 
stable fly larvae is highest when a sample is moist (≈350% water, wet 
wt./dry wt.) with high (≈ 200 ppm) ammonium content, high (≈ 3 μS/
cm) electric conductivity, high microbial activity, and a temperature of 
≈ 23°C (Talley et al. 2009, Friesen et al. 2016). Preferences for sub-
strate pH varies from acid (Gilles et al. 2008) to neutral (McPheron 
and Broce 1996, Talley et al. 2009) to slightly alkaline (Rasmussen 
and Campbell 1981, Friesen et al. 2016). Behavioral studies indicate 
that stable fly larvae actively orient towards increasing concentrations 
of ammonium (D.B.T., unpublished data). The addition of ammonium 
to laboratory substrates increases survival (Friesen et al. 2017).
Fig. 3. Stable flies feeding with heads oriented upwards (A) and horn flies feeding with heads downward (B). The horn fly also holds its wings more angled 
away from the body.
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Traditionally, stable fly developmental substrates have been as-
sociated with decaying vegetation contaminated with animal feces 
and urine (Fig. 4). However, early reports indicated that major stable 
fly outbreaks in the central U.S. were associated with decomposing 
vegetation in the absence of animal waste. Threshing straw stacks 
were the primary sources of stable flies during a major outbreak 
in 1912 which caused the death of hundreds of horses and cattle 
(Bishopp 1913b). More recently, serious outbreaks associated with 
the application of poultry manure to agricultural land (Cook et al. 
2018b) and with crop residues have been reported (Herrero et al. 
1989, Barros et al. 2010, Corrêa et al. 2013, Solórzano et al. 2013, 
Dominghetti et al. 2015, Cook et al. 2018a, Cook et al. 2020).
Under optimal temperatures, 25–30°C, immature development 
(egg-pupariation) takes 13 d (Larsen and Thomsen 1940, Lysyk 
1998, Florez-Cuadros et al. 2019), but like other ectotherms, devel-
opment depends on the temperature. The thermal threshold for stable 
fly developmentis estimated to be 11.5°C and few flies develop suc-
cessfully at 35°C (Larsen and Thomsen 1940, Florez-Cuadros et al. 
2019). Substrate quality has little effect upon developmental rate but 
does have a large effect on imago size (Florez-Cuadros et al. 2019).
Adult Behavior and Dispersal
Adult stable flies can live up to 35 d in the laboratory, but probably 
survive less than two weeks in the field (Killough and McKinstry 
1965, Salem et al. 2012a). Adults can be active year-round in some 
locations, but numbers typically decline as daily maximum tem-
peratures increase to above 30°C (Mullens and Peterson 2005) or 
minimum temperatures decrease to below freezing (Taylor et  al. 
2007). Stable flies are more sensitive to high temperatures than house 
flies (Lysyk 1998). In temperate regions, stable fly populations tend 
to be bimodal with peaks in early and late summer (Scholl 1986, 
Broce et al. 2005, Taylor et al. 2007, Jacquiet et al. 2014). In more 
northern locations, one peak can be observed in midsummer (Lysyk 
1993, Skovgård and Nachman 2012) and in more southern, warmer, 
regions a single peak occurs in late winter or spring (Mullens and 
Meyer 1987, Pitzer et al. 2011b). Temperature and precipitation ap-
pear to be the primary drivers of stable fly population fluctuations 
(Taylor et al. 2007, Taylor et al. 2017), however, the relative import-
ance of those two parameters varies by region.
Stable flies are strong fliers. Median dispersal from larval devel-
opmental sites in a Nebraska study was 1.6 km, but fewer than 5% 
dispersed beyond 5.1 km (Taylor et al. 2010). Short-distance dispersal 
between livestock facilities was documented by Pitzer et al. (2011a) 
when stable flies containing cattle bloodmeals were collected on large 
equine farms in Florida, within 16 h of feeding at cattle premises 0.8 
and 1.5 km away. In Florida, Hogsette and Ruff (1985) demonstrated 
movement of 225 km with frontal systems by self-marked wild stable 
flies. However, whether their work demonstrated directed movement 
or drifting on air currents is not known. Actual dispersal distances ap-
pear to be determined by the distance they must fly to find hosts. They 
remain close to the ground during short-distance dispersal (Williams 
and Rogers 1976, Gersabeck and Merritt 1983, Hogsette et al. 1989), 
and once suitable hosts have been located, they tend to remain within 
the vicinity of the hosts (Bailey et al. 1973).
In temperate zones, the mechanisms for overwintering and 
recolonization in the spring are unclear. Small numbers of adult 
stable flies can be observed outdoors on warm winter days 
(Berkebile et al. 1994), and significant numbers can be observed 
in the spring prior to the accumulation of enough degree-days for 
them to have developed locally (Taylor et al. 2007, K. Friesen, un-
published data). Diapause has not been demonstrated in stable 
flies and no freeze-tolerant stages have been identified (Berry et al. 
1978). Three potential overwintering mechanisms have been pro-
posed: 1) retarded development in heat-producing substrates that 
remain above freezing throughout the winter, 2) continuous devel-
opment in protected environments such as dairy barns, and 3) re-
colonization by either short- or long-range dispersal from regions 
to the south where continuous reproduction and development 
are possible. It is possible, if not likely, that all three mechanisms 
are involved with stable fly overwintering and recolonization, al-
though the relative importance of each may depend upon local 
climate and cultural practices.
Genetics
The stable fly has five homomorphic, metacentric to sub-
metacentric chromosomes with no X or Y chromosome (Joslyn 
et  al. 1979). A  dominant male-determining locus is located on 
chromosome 1 (Willis et al. 1981), but the locus has yet to be iden-
tified. Several phenotypic markers have been assigned to chromo-
somes, including carmine eye on chromosome 2, black pupa on 
chromosome 3, and rolled wing on chromosome 4 (Willis et al. 
1981, Willis et al. 1983). Radiation-induced chromosomal trans-
locations were pursued to develop a genetic sexing system based 
on malathion resistance, i.e., resistance linked to males resulting 
in female lethality on challenge with malathion (Seawright et al. 
1986). The development of sexing systems for stable fly was not 
eventually pursued, given the challenges associated with using the 
sterile insect technique to reduce stable fly populations (see Sterile 
Insect Technique section, below). The stable fly genome was re-
cently sequenced (Olafson et  al. 2021). The sequencing effort 
identified a robust chemosensory gene family, including expanded 
lineages of chemoreceptors that are involved in bitter taste per-
ception; an immune system encoding an increased number of 
antimicrobial peptides relative to Drosophila but comparable to 
the related house fly; a large cytochrome P450 gene family that 
suggests the stable fly has an enhanced mechanism for metabolic 
detoxification. Meisel et al. (2020) combined this genomic infor-
mation with gene expression data to identify the stable fly cryptic 
sex chromosome formed when the ancestral sex chromosome 
fused with one of the five autosomes.
Fig. 4. Stable fly white larvae and reddish pupae in typical developmental 
substrate consisting of moist decaying vegetation soiled with feces and 
urine.
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Journal of Integrated Pest Management, 2021, Vol. XX, No. XX 5
Stable Fly Effects in Human and 
Animal Systems
Economic Losses
Most of the economic loss caused by stable flies can be attributed 
to direct damage and the behavioral and physiological responses 
of livestock to the adult flies’ feeding activity. During severe infest-
ations, host mortality has been observed (Bishopp 1913b, Cook 
et al. 1999, Huchzermeyer et al. 2001, Elkan et al. 2009). Stable flies 
reduce cattle productivity in the U.S. by approximately $2 billion 
per year (Taylor et al. 2012a); $2.66 billion using 2018 cattle inven-
tories and commodity prices. In other countries, economic losses are 
estimated to be $6.8 million in Mexico (Rodríguez-Vivas et al. 2017) 
and $340 million in Brazil (Grisi et al. 2014).
Stable flies impact livestock production by reducing weight gain 
and feed efficiency in beef feeder calves (Campbell et al. 1977, Berry 
et al. 1983, Catangui et al. 1993). Stable flies have a greater effect 
on cattle fed a “growing” ration, 0.9  kg/d, than those fed a “fin-
ishing” ration, 0.2  kg/d (Campbell et  al. 1987). For both rations, 
feed efficiency was reduced by 11–12%. Three-fourths of the loss is 
due to the negative effects of bunching and heat stress and the re-
maining quarter is the result of blood loss and metabolic energy ex-
pended in defensive behaviors (Wieman et al. 1992). In dairy cattle, 
increases in milk production following the implementation of biting 
fly control have been reported (Bruce and Decker 1958, Morgan and 
Bailie 1980, Block and Lewis 1986). The effects of stable flies on 
milk production, and probably other livestock production systems 
as well, are confounded by interactions with host nutrition (Cheng 
and Kesler 1961, Miller et al. 1973). This is especially relevant given 
that most of the studies examining the impacts of stable flies on live-
stock production were done 30–50 yr ago; significant advances have 
been made in dairy system productivity and nutrition since that time.
Stable fly outbreaks have been associated with decomposing 
crop residues in several regions of the world during the last 20–30 
yr. Devastating outbreaks in Australia, Central and South America 
(Monty 1972, Kunz and Monty 1976, Herrero et al. 1989, Herrero 
et al. 1991, Cook et al. 1999, Koller et al. 2009, Dominghetti et al. 
2015, Solórzano et al. 2015) produced infestation levels greater than 
1,000 stable flies per animal (Fig. 5; Solórzano et al. 2015), greatly 
exceedingthe economic threshold (≈15 flies per animal; Berry et al. 
1983, Campbell and Berry 1989). Such infestations can last for sev-
eral months, reduce animal productivity to zero and cause mortality 
in livestock and companion animals. Flies arising from these out-
breaks can be especially difficult to control because, unlike those 
generally associated with livestock, developmental sites, extend over 
great areas (i.e., tens of thousands to millions of hectares). The agro-
nomic industries creating the fly developmental substrates are not 
usually negatively affected by the flies and thus are not incentivized 
to expend resources for stable fly control. Although such outbreaks 
have not been observed in the U.S.  in recent years, they have oc-
curred previously (Bishopp 1913a) and, with changing climatic con-
ditions and agronomic practices, have the potential to reoccur.
Animal Health Concerns
Stable fly bites are more painful than those of other biting insects 
(Elzinga and Broce 1986, Hogsette and Ose 2017). They cause 
discomfort that disrupts feeding, reproductive, and resting behav-
iors of livestock. Additionally, stable fly bites can induce strong 
defensive behaviors and physiological stress. Stable fly infestation 
levels can exceed 1,000 flies per animal in instantaneous counts 
(Solórzano et  al. 2015). LaBrecque et  al. (1975) estimated that 
for every fly observed on a host, 50–60 are present in the vicinity. 
Hence daily biting rates are 50–60 times higher than instantaneous 
counts. Since stable flies imbibe ≈12  µl of blood per bloodmeal 
(Salem et al. 2012a), infestations at the economic threshold level, 
≈15 flies per animal (instantaneous count) (Berry et  al. 1983, 
Campbell and Berry 1989), can result in the loss of ≈15  ml of 
blood, while instantaneous counts of 1,000 flies can result in the 
loss of >1  L of blood per day. In response to stable flies, cattle 
bunch together to protect their legs from the biting flies (Wieman 
et al. 1992, El Ashmawy et al. 2019) and exhibit defensive behav-
iors such as tail flicking, skin twitching, and leg stamping (Mullens 
et al. 2006). In addition to their impact on livestock, stable flies can 
disrupt human recreational activities (Hansens 1951, Fosbrooke 
1963, Newson 1977, Merritt and Newson 1978, Hogsette et  al. 
1987, Urban and Broce 1998, Elkan et al. 2009). However, stable 
flies are not known to be primary vectors of any medically im-
portant human pathogens.
Compared with other biting flies such as mosquitoes (Diptera: 
Culicidae) and tsetse flies (Diptera: Glossinidae), stable flies are poor 
vectors of pathogens. The mode of transmission for nearly all patho-
gens associated with stable flies is mechanical, requiring interrupted 
feeding on an infected host followed by subsequent feeding on an 
uninfected host within a short timeframe. Due to their painful biting, 
interrupted feeding is common for stable flies under natural con-
ditions, with nearly three out of every four feeding attempts being 
interrupted (Schofield and Torr 2002). Because most stable fly hosts 
are herd animals, many hosts are usually within close proximity, al-
lowing opportunities for completing the mechanical transmission 
cycle. In heavy infestations (i.e., instantaneous counts > 100 flies 
per animal) biting rates can be > 6,000 per day. Significant levels of 
disease transmission can occur under these conditions, even when 
transmission efficiency is low.
Stable flies have been associated with the transmission of several 
pathogens (Baldacchino et  al. 2013a, Sharif et  al. 2019), however, 
for most, one or more of the four criteria for implicating a species 
as a primary vector (Barnett 1960) have not been empirically dem-
onstrated. Several pathogens are expanding their ranges and could 
potentially invade North America. One of the most important inva-
sive livestock diseases in Europe, African Swine Fever, can be trans-
mitted by stable flies either through their bite (Mellor et  al. 1987) 
or via ingestion of infected flies (Olesen et al. 2018). Several of the 
important or potentially concerning infectious agents, the diseases or 
Fig. 5. Top view of a calf with a large infestation of adult stable flies. Flies 
feeding on the back, shoulder and ears is unusual under normal infestation 
levels, where they typically feed on the lower legs.
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conditions they produce, and their association with stable flies are 
described below.
Habronematidosis
Habronema microstoma Creplin (Spirurida: Habronematidae), one of 
several species of Habronema responsible for stomach worms of horses, 
is the only pathogen known to be transmitted cyclodevelopmentally by 
stable flies (Traversa et al. 2008), meaning the worms undergo develop-
mental changes in the insect before they are transmitted. Adult worms 
in the horse stomach release eggs which pass through the digestive 
system and into the environment with the feces. Eggs hatch soon after 
being excreted and first instar larvae are eaten by stable fly larvae in 
manure. The worms molt twice in the fly larvae and develop into in-
fective (L3) larvae by the time the adult flies emerge. The L3 migrates 
to the fly’s head and escapes while it is feeding, usually around the 
nose or lips of the definitive host. L3 larvae are ingested by the host, 
pass to the stomach where they mature and complete the cycle (Pugh 
et al. 2014). Gastric habronematidosis can cause anorexia, diarrhea, 
and weight loss in horses.
If the L3 nematode larvae are deposited near a wound or the eyes, 
they are unable to develop into adults and continue to migrate in the 
skin causing inflammation and a condition referred to as “summer 
sore”. Summer sore lesions are proliferative and granulomatous, often 
bloody, itchy, ulcerated, and contain calcified granules (Barlaam et al. 
2020). Lesions may heal or be recurrent. Recurrent lesions will ap-
pear to heal during colder weather and reappear as weather warms 
in spring. Lesions may attract more flies leading to superinfection 
(Barlaam et  al. 2020). Deposition of L3 larvae near mucus mem-
branes, such as the eyes or genitals, can result in mucocutaneous 
habronematidosis. When the eyes are involved, symptoms include 
conjunctivitis, blepharitis, photophobia, and lacrimation. Urogenital 
infections can produce dysuria and frequent urination.
Equine Infectious Anemia (EIA)
Stable flies have been implicated in the mechanical transmission of 
several pathogens. EIA is possibly the best documented in North 
America. EIA is an infectious, viral disease of horses and other equids 
with worldwide distribution. The virus is an RNA virus in the family 
Retroviridae, genus Lentivirus. Also known as Swamp Fever, EIA 
is most active during seasons and at locations where biting flies are 
abundant (Scott 1922). Deer flies (Chrysops spp.; Foil et al. 1983), 
horse flies (Tabanus spp (Stein et al. 1942) and Hybomitra spp.) and 
stable flies (Stein et al. 1942, Foil et al. 1983) can transmit the virus 
mechanically. The relative importance of these flies as vectors de-
pends upon season and geographical region.
Exotic Diseases With Invasive Potential
Lumpy Skin Disease (LSD)
Lumpy skin disease is caused by a virus in the family Poxviridae, 
genus Capripoxvirus, and infects cattle and water buffalo in Africa, 
the Middle East, expanding into southeastern Europe (EFSA 2019, 
2020). The virus causes considerable economic losses due to emaci-
ation, hide damage, infertility, mastitis, loss of production, and mor-
tality (Davies 1991, Al-Salihi 2014). Stable flies can transmit LSD 
virus between sheep (Kitching and Mellor 1986) and cattle (Sohier 
et al. 2019, Issimov et al. 2020) and outbreaks are associated with 
high stable fly infestation levels (Kahana-Sutin et al. 2016, Gubbins 
et al. 2018).
Besnoitiosis
Besnoitiosis is a chronicand debilitating, emergent disease of cattle 
and other ruminants. The etiological agent, Besnoitia besnoiti, is 
an apicomplexan protozoan related to Toxoplasma gondii with 
a similar life cycle. Most cases of besnoitiosis are in Africa; how-
ever, the disease is spreading to the Middle East, southern Asia, 
southern Europe, and South America (Álvarez-García et al. 2013). 
Besnoitia besnoiti bradyzoites can be transmitted mechanically by 
the bites of blood-feeding flies (Álvarez-García et al. 2013, Duvallet 
and Boireau 2015). In France, the temporal pattern of Besnoitia 
sp. transmission coincides with stable fly abundance (Liénard 
et  al. 2011) and in some locations, stable flies appear to play a 
primary role in besnoitiosis epidemiology. Another Besnoitia spe-
cies, Besnoitia bennetti, infects equines and has been reported from 
North America (Ness et al. 2005, Elsheikha et al. 2020). The mode 
of transmission of B. bennetti is unknown, but S. calcitrans must be 
considered as a participant.
Trypanosomiasis (Trypanosomosis)
Trypanosomes are parasitic flagellate protozoans in the order 
Trypanosomatida, family Trypanosomatidae, genus Trypanosoma. 
Several species of Trypanosoma cause serious and debilitating dis-
eases in humans, livestock, and companion animals. Human tryp-
anosomiasis, also known as African Sleeping Sickness, is caused by 
Trypanosoma brucei gambiense Dutton  and Tryanosoma brucei 
rhodesiense Stephen and Fantham  which undergo cyclical de-
velopment in their primary vector, Glossina spp. Several other 
Trypanosoma species can be transmitted directly from host-to-host 
via mechanical transmission; stable flies have been implicated as vec-
tors, as described below.
Trypanosoma evansi (Steel)  (= Trypanosoma brucei evansi) is 
widely distributed in Saharan Africa, southern Asia, and South 
America (Radwanska et  al. 2018) and is the etiological agent of 
surra, a debilitating and economically important disease of equines, 
bovines, and camelids (Aregawi et al. 2019). T. evansi cannot de-
velop in fly hosts and, therefore, can only be transmitted mechan-
ically. Stable flies and horse flies (Tabanidae) have been implicated 
as the primary vectors (Luckins 1988, Lun et al. 1993, Desquesnes 
et al. 2008). Stable flies have been associated with the transmission 
of T. evansi in time and space (Desquesnes et al. 2008, Rodríguez 
et  al. 2014), but transmission by S.  calcitrans has not been ad-
equately demonstrated. Bouet and Roubaud (1912) concluded that 
Stomoxys spp. play a major role in surra epidemiology. However, 
their experiments lacked controls and the rigor expected in more 
recent studies. Sumba et al. (1998) demonstrated transmission by 
Stomoxys species other than S.  calcitrans. Failure of laboratory 
transmission studies should not be taken as strong evidence that 
S.  calcitrans is not involved in the transmission of T.  evansi be-
cause most such studies involve relatively small numbers of flies 
or fly bites (Ngeranwa and Kilalo 1994, Desquesnes et al. 2013). 
In the field, animals can sustain tens of thousands of bites per day 
during severe outbreaks. Such numbers can overcome low odds for 
transmission. Additional research is needed to elaborate the role 
of Stomoxys spp. and specifically S. calcitrans, in the epidemiology 
of surra.
Stable Fly Challenges for Animal Systems
Intensive Animal Systems
Dairy Cattle
Stable flies are one of the most damaging insect pests for the dairy 
industry. The severe and intense biting behavior of the adult stable 
flies negatively impacts dairy cows by disrupting feeding and causing 
bunching of cattle, both decreasing milk production. As few as one fly 
per animal can affect the behavior of dairy cattle leading to protective 
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bunching (El Ashmawy et  al. 2019). Bishopp (1913a) indicated 
that a stable fly outbreak in the late summer of 1912 reduced milk 
production in cows by 40–60% and, in some instances, cows went 
completely dry.
Management systems for dairy cattle production include ani-
mals on grass pasture, in dry lots, and in free stall barns. Pastured 
and dry lot dairy cows have many of the same issues with stable 
flies as do pastured and confined beef cattle respectively. Although 
stable flies do not tend to enter buildings, they will venture a limited 
distance into open-sided barns (20–30 m). As a result, the animals 
will bunch towards the middle of the barns, avoiding areas near the 
open exterior walls.
Calf hutches, both individual and group, can be primary develop-
mental sites for stable flies in dairies (Schmidtmann 1988, Meyer and 
Shultz 1990). Calf hutches should be sited to promote good air flow 
and drainage to avoid or eliminate moisture in the bedding. Fans can 
minimize adult fly movement around calves and help keep bedding 
dry (J.A.H, unpublished data).
Some bedding materials favor developing stable flies more than 
others. In addition to calf hutches, stable flies develop in spilled or 
waste feed. Proper sanitation should eliminate those developmental 
substrates (see Management section, below).
Confined Cattle/Feedlot
Although stable flies can develop in aged bovine feces mixed with 
soil, larval densities are far lower than in substrates where manure 
is mixed with vegetation. Primary developmental sites for stable flies 
in confined animal operations are adjacent to the feed bunks where 
waste feed combines with animal wastes and under fence lines where 
manure accumulates undisturbed (Skoda et  al. 1991). Spilled feed 
in front of feed bunks, in the feed mill, and elsewhere in the facil-
ities and poorly maintained bunker silos and silage bags can also be 
sources.
Control options for stable flies in confined cattle facilities 
are limited. As for all systems, sanitation is the primary concern. 
Although regular removal of manure, every two weeks, has the po-
tential to reduce stable fly populations by 30–50% (Thomas et al. 
1996), few feedlots have the resources to maintain such a schedule. 
Maintaining good drainage, avoiding leaking watering systems, and 
avoiding accumulations of waste feed will reduce the potential for 
stable fly development. Traps and targets can assist in the control of 
adult flies, although their use in the feedlot environment has not been 
thoroughly explored. Insecticides can be used in extreme situations; 
however, their effectiveness is very short term and ear tags are inef-
fective (see Chemical Control section, below)
Zoological Gardens
Zoological gardens (“zoos”) are collections of animals, native and/
or exotic, usually in or near cities, and these animals can serve as 
hosts for stable flies. It has not been determined whether adult stable 
fly populations at zoos are from nearby areas and have been con-
centrated by the attraction of the zoo, or if they have dispersed 
from more distant locations. During a stable fly project in Garden 
City, Kansas in 1996, large numbers of stable flies arrived suddenly 
with the passing of a weather system. During this event, stable flies 
were observed at a local zoo feeding on lions, tigers, leopards, bears, 
emus, and especially canids (Hogsette and Farkas 2000, Farkas and 
Gyurcsó 2006). Summer stable fly populations have been docu-
mented at urban zoos throughout the U.S. (Kanzia 2015, Hogsette 
and Ose 2017).
Extensive Animal Systems
Range and Pasture Cattle
As their name implies, stable flies are usually considered to be pri-
marily pests of livestock in the barnyard environment where soiled 
animal bedding provides suitable substrates for larval development. 
However, since the early 1980s, they are being recognized increas-
ingly as serious pests of range and pastured livestock (Hall et  al. 
1982, Campbell et al. 2001). Broce et al. (2005) implicated the tran-
sition from feeding cattle small bales of hay, often spread throughoutthe pasture, to the use of large round bales in stationary feeders for 
the increased numbers of stable flies found in pastures; a conclu-
sion supported by subsequent studies (Talley et al. 2009, Taylor and 
Berkebile 2011). However, Taylor et al. (2020) suggest that other, un-
identified, developmental sites may also be contributing large num-
bers of flies to pasture populations.
The impacts of stable flies on range cattle are understudied. The 
most frequently observed sign of stable fly activity in pastured cattle 
is bunching of animals and leg stomping. Infested cattle will often 
stand in water to protect their legs from the biting flies. Bunching 
results in damage to forage and, on fragile soils, may create blowouts 
(i.e., hollows eroded by the wind). Bunching of cows can result in 
injury to calves if they get stepped on. Foot stomping and retreating 
to water holes can result in foot rot and water pollution. Because of 
this relatively recent increased activity in pastured cattle as a result 
of these husbandry changes, studies of weight gain and economic im-
pacts have been limited. Although weight gain differences have been 
noted in pastured cattle, differences shown in studies by Cutkomp 
and Harvey (1958) and Cheng (1958) cannot be attributed to only 
stable flies due to the presence of both horn fly and stable fly popu-
lations on control animals. Campbell et al. (2001) reported a 21% 
improvement in average daily gain with partial control of a relatively 
low population of stable flies on pastured yearling cattle (stockers).
Mixed Animal Systems
Equine
Stable flies cause irritation and weakness in horses and account for 
considerable blood loss in severe cases. Wounds caused by feeding 
flies can serve as sites for secondary infection. As with many other 
hosts, stable fly feeding occurs mainly on the front legs. Heavy 
feeding can result in blood-matted hair and an unthrifty appearance. 
In a study on large equine farms in Florida, Pitzer et  al. (2011a), 
demonstrated that most of the bloodmeals in field-collected stable 
flies were from cattle (64.6%), equines (24.3%), humans (9.5%), 
and dogs (1.6%). With no cattle within 0.8 km, these findings sug-
gests that many flies found on equine facilities possibly originate 
elsewhere and are not targeting horses as preferred hosts. Defensive 
behaviors (Mullens et al. 2006) expressed by horses can make them 
difficult to show or ride (Hogsette 1981), and in heavily infested 
areas, animals cannot be pastured during the day.
Stable Fly Management: An Integrated 
Approach
The primary elements of an Integrated Pest Management (IPM) pro-
gram are economic thresholds, monitoring, and management. Economic 
thresholds for stable flies are difficult to calculate. Because their bites 
are more painful than those of most blood-feeding insects, even very 
small numbers can evoke considerable responses and losses. As few as 
one fly per cow foreleg in an instantaneous count can induce bunching 
of dairy cattle (El Ashmawy et al. 2019). Hence, on cattle, the economic 
threshold is exceptionally low: five flies per leg or about 15 flies per 
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animal (Berry et al. 1983, Campbell and Berry 1989). Economic thresh-
olds have not been determined for equines.
One of the primary principles of IPM of stable flies, and most 
other filth flies, is to attack the pests when they are most concen-
trated, least mobile, and most accessible (Novak et al. 2010). This 
indicates source reduction of immature developmental sites should 
be the target of control efforts. However, most filth flies develop in 
ephemeral substrates and immatures are spatially clumped. In add-
ition, developmental sites are often difficult to identify. Dispersal of 
adults from immature developmental sites appears to be dependent 
upon the distribution of available hosts and is therefore not well 
defined. These factors can make monitoring and control difficult. 
Once adult populations have reached the economic threshold, it is 
too late to initiate sanitation and other management options dir-
ected towards the immature stages. Sanitation and immature stage 
management efforts must be implemented prophylactically to avoid 
losing these most critical elements of the arsenal for managing stable 
flies and other filth flies.
Monitoring
Several methods are available for monitoring stable fly populations. 
Adult monitoring usually consists of either on-animal counts or trap 
catches. On-animal and trap counts tend to be relative to the overall 
stable fly population, however, both are also influenced by static and 
dynamic environmental variables (Berry et al. 1983, Thomas et al. 
1989, Guo et al. 1998). On-animal counts have the advantage that 
they are causally related to the level of animal irritation (Berry et al. 
1983, Mullens et al. 2006), but the probability of error is also higher 
because of the difficulty involved with counting moving flies and 
visibility of the flies at different positions on the animals and on 
animals of different colors (Berry et al. 1983). On-animal counts are 
also labor-intensive and therefore usually encompass a shorter time 
period than trap counts (Taylor et al. 2020). Traps can be employed 
for extended periods of time without attention thus their counts can 
average dynamic environmental factors such as weather. However, 
they are still influenced by static environmental factors such as prox-
imity to hosts, developmental sites, and other physical structures 
such as buildings and vegetation (Berry et al. 1983, Guo et al. 1998, 
Taylor et al. 2013).
On-Animal
Stable flies prefer feeding on the front legs of cattle below the chest 
floor (Berry et al. 1983). As such, when recording on-animal num-
bers it is appropriate to use leg counts, which is the number of stable 
flies visible from a single point of view on the outside of one front 
leg and on the inside of the other (Campbell and Hermanussen 
1971, Berry et  al. 1983). Leg counts are an instantaneous metric, 
representing the number of flies present on the host at the time of 
the count. Given that stable flies feed for 2–4 min and then leave 
the host to digest their meal, each fly observed in an instantaneous 
count may represent 50–60 flies present in the vicinity that will feed 
on the host during the course of the day (LaBrecque et al. 1975). 
On-animal counts are normally done during the late morning and/
or early afternoon, but ideal times will vary according to climate. 
Biting activity is reduced at temperatures below 15 and above 30°C 
(Hafez and Gamal-Eddin 1959), so times of day when these condi-
tions are common should be avoided. Berry and Campbell (1985) 
found leg counts in Nebraska to be more closely correlated with time 
of day than temperature and humidity. This may be because the tem-
peratures during their study were relatively consistent and remained 
within the window of stable fly activity.
Traps
Traps can be used for monitoring, sampling, and managing stable 
fly populations. All traps have biases, whether relative to chrono-
logical or physiological age, sex, or mating status. Although quantity 
of catch is important for management programs, it is not necessarily 
the most important criteria for monitoring or sampling programs. 
Trap placement in the environment is also important and can im-
pact fly capture. More stable flies are captured when traps are placed 
as close as possible to the host animals (Hogsette and Ose 2017). 
Trap height above the ground or vegetation (Beresford and Sutcliffe 
2008), sunlight (Rugg 1982), and number of traps in the environ-
ment (Taylor et al. 2013) can impact trapping success. The biases of 
monitoring and sampling methods must be understood and compat-
ible with the goals of the program (Gilles et al. 2007, Taylor et al. 
2020). Contemporary stable fly trapsare based upon visual attrac-
tion. Olfactory stimulants can increase trap catches (Cilek 1999, 
Mihok et al. 2007, Phasuk et al. 2016, Zhu et al. 2016), but have 
not been used widely in management programs. Stable fly traps com-
monly used today are either adhesive traps or cloth traps with a 
funnel collection mechanism.
Sticky traps made from Alsynite fiberglass (Williams 1973, Broce 
1988) are particularly effective and have been frequently used for 
monitoring and management. The traps act as visual targets, but 
the exact mechanisms of attraction are unknown. It is generally ac-
cepted that flies are attracted to the wavelengths of sunlight reflected 
by the Alsynite panels (Thimijan et  al. 1973, Agee and Patterson 
1983, Zachs and Loew 1989). The Olson trap (Arbico Organics, 
Oro Valley, AZ) is a commercially available Alsynite-based trap with 
preglued sleeves used extensively for stable fly monitoring and re-
search (Ose and Hogsette 2014, Machtinger et al. 2016a, Hogsette 
and Foil 2018) as was the Broce trap (Taylor et al. 2007, Dominghetti 
et  al. 2015, Taylor et  al. 2017). Most studies using Alsynite traps 
have observed an excess of males in the collections, often approxi-
mately two males for every female (Scholl 1986, Guo et al. 1998, 
Taylor and Berkebile 2006), but Hogsette and Ruff (1990) did not 
observe this on the Olson trap. In addition to Alsynite, several other 
materials have been found to be effective. Corrugated plastic ma-
terials Aluma Panel (Cumming, GA; Cilek 2003) and Coroplast 
(Vanceburg, KY; Beresford and Sutcliffe 2006) are convenient ma-
terials for making stable fly traps. White Coroplast captures more 
stable flies per unit area than Alsynite (Beresford and Sutcliffe 2006) 
and is also more readily available, less expensive, and easier to cut 
and handle than Alsynite. Blue polyethylene screens with a peak of 
reflectance around 460 nm and covered with transparent sticky film 
have also been evaluated with success (Sharif et al. 2020). Care must 
be taken to service adhesive traps before they become saturated with 
flies. Stable flies tend to avoid adhesive traps with many trapped flies 
already present (Beresford and Sutcliffe 2017).
Sticky traps designed to attract stable flies tend to collect few 
other insects, but the non-target species collected will vary by loca-
tion and season. Small flying insects will stick randomly to the trap 
if they are carried by the wind, as will any very abundant insect in 
the vicinity of the trap. Polyethylene blue screens caught very few 
pollinators (Sharif et al. 2020). Insectivorous birds or reptiles may 
be attracted to the trap and come in contact with the glue when at-
tempting to eat the dead flies; in most cases, those animals are strong 
enough to free themselves from the trap, especially when the traps 
are full of flies.
Several variations of blue-black cloth traps originally developed 
for tsetse fly control in Africa have been evaluated for collecting 
Stomoxys spp. including S.  calcitrans (Mihok et  al. 1995, Mihok 
2002, Mihok et al. 2006, Onju et al. 2020). Of these, the Vavoua 
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and Nzi traps are best characterized (Fig. 6). Blue-black cloth traps 
tend to collect slightly older stable flies than do Alsynite and other 
adhesive traps (Taylor and Berkebile 2006). Because they do not rely 
on an adhesive to collect flies, these cloth traps require less frequent 
servicing than sticky traps (Gilles et al. 2007) but are seldom used in 
the U.S. apart from research endeavors.
Substrate Sampling
Sampling substrates for immature flies (larvae) can be used as a 
monitoring method. Core samples are the most direct evaluation 
method (Skoda et al. 1991, Berkebile et al. 1994, Talley et al. 2009) 
and have the advantage that all occupants of the sample can be iso-
lated and characterized as well as the physical and microbial proper-
ties of the substrate (Talley et al. 2009, Wienhold and Taylor 2012, 
Friesen et al. 2016). Most stable fly immatures will be found within 
5 cm of the surface (Taylor and Berkebile 2011), so core samples do 
not need to be deep. However, core samples can be difficult to collect 
due to the fibrous nature of most developmental substrates and the 
high moisture content. Related to core samples is non-quantitative 
trowel sampling. This method can be used for determining presence/
absence of immature stable flies within a substrate and their ap-
proximate age. Other fly species including house flies can be found 
in substrates similar to those occupied by stable flies. Accurate iden-
tification of immature flies in substrate samples is essential (Fig. 2).
Pupal traps work on the premise that stable fly larvae seek a 
slightly drier location within the substrate to pupariate. Traps are 
normally constructed with 6.3 mm (1/4-inch) hardware cloth filled 
with wood chips or other coarse substrate (Hogsette and Butler 
1981, Skoda et al. 1996). A disadvantage of pupal traps is that they 
are susceptible to damage when animals are present (Skoda et  al. 
1996) unless more durable traps are constructed (Taylor et al. 2020).
Adult fly emergence traps are essentially cages placed over devel-
opmental substrates that capture flying insects as they emerge from 
the substrate. Most emergence traps have wood, metal, or plastic 
frames with screen cones or pyramids leading to the collecting 
chamber and a funnel mechanism at the top to collect emerging flies 
(Taylor and Berkebile 2011, Taylor et  al. 2020). Emergence traps 
are easy to install and service and have the advantage that insects 
remove themselves from the substrate. Emergence traps also sample 
a relatively large area, so fewer traps are needed compared to core 
samples. The primary disadvantage of emergence traps is that they 
do not sample non-flying denizens of the substrate, nor do they pro-
vide a picture of the age structure of the immatures at a given point 
in time. Emergence traps exclude ovipositing females, therefore for 
long term studies, they should be relocated at least every two weeks 
(the approximate time required for egg to adult stable fly develop-
ment). The final disadvantage of emergence traps is that animals may 
interfere with the traps, therefore, livestock must be excluded from 
test areas to protect the traps.
Management
Stable fly management poses unique challenges. Stable fly adults are 
on the hosts for a relatively short time and tend to congregate on the 
lower parts of the legs, reducing the efficiency of on-host insecticide 
treatments. Immature developmental sites can produce vast numbers 
of flies from relatively small areas. Because developmental substrates 
are ephemeral, they are often no longer productive by the time adults 
are biting and a search is done for their source. To overcome these 
challenges, multifaceted or integrated approaches are needed for the 
management of stable fly populations. Such programs usually begin 
with cultural control and sanitation, the removal of potential de-
velopmental sites, and modification of animal production methods 
to reduce the development and impacts of stable flies. Mechanical, 
biological, and chemical control efforts are doomed to failure if not 
done in conjunction with cultural changes and sanitation. Because 
stable flies are strong fliers, localized management efforts may have 
a limited effect on reducing adult fly populations due to the immi-
gration of adult flies from off-site developmental substrates. Keep 
in mind that at a given location, the number of flies from a devel-
opmental site will be related to the reciprocal of the square of the 
distance to that site (Taylor et al. 2010). Therefore, even though lo-
calized control efforts will not eliminate stable flies, the lack of such 
efforts will result in far greater local populations. Optimally, stable 
flymanagement programs should be coordinated area-wide efforts 
(Taylor 2021).
Cultural Control
Sanitation to minimize on-site development of immature populations 
is critical (Thomas et al. 1996) and serves as the foundation of a suc-
cessful control program, regardless of operation type, particularly 
on facilities where management options are limited (i.e., accepted 
organic pesticides). Waste management practices such as making 
waste and bedding easy to remove from a facility, storing manure 
Fig. 6. Blue–black targets developed for tsetse fly control in Africa have 
proven useful for stable flies. The Vavoua (top) and Nzi (bottom) traps have 
been evaluated under many conditions and locations. When utilized as 
traps or when fabric is treated with an approved insecticide, these traps can 
capture or kill large numbers of flies.
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and waste to allow for drying, and removing waste frequently can 
reduce stable fly development. In pastures, fly control should start 
with fly source reduction through debris management. This involves 
removal and proper disposal of residues, starting with winter hay 
feeding sites, and continuing through summer with other accumula-
tions of rotting feed and bedding contaminated with animal wastes. 
While timely removal of stall litter and other potential fly habitats 
can be helpful, improper management of these materials after they 
have been removed can move the problem from one place to another 
(Cook et  al. 1999). It is essential to consider the potential for fly 
development in the material once removed from the facility struc-
tures or pastures. Soiled bedding should be removed and properly 
disposed of weekly. Removed materials can be spread thinly (i.e., less 
than 5 cm deep), harrowed into soil, or densely piled vertically and 
composted (Hogsette et al. 1987, Pickens et al. 1994). If removal is 
not possible, bedding can be treated with an insect growth regulator 
(IGR) as described in the Chemical Control section below.
Keeping vegetation and animal waste separate will reduce the 
number of stable flies. Because hay bale waste can be a significant 
source of stable fly development, periodically moving round bale hay 
placement sites can reduce fly development. Similarly, keeping silage 
dry and off the ground, avoiding accumulations of undisturbed 
silage at the edges of feed bunks, and covering silage can significantly 
reduce fly problems. Incorporating wet crop residues into the soil at 
least 15 cm deep (Cook et al. 2020) as quickly as possible following 
harvest and stopping irrigation once harvest is complete can pre-
vent fly development in crop residues. Compaction of some soil types 
after incorporation can reduce fly emergence (Cook et al. 2020).
Management of feed and water (e.g., with efficient and leak-
free water systems and good drainage) can reduce moisture in the 
environment. Cleaning or avoiding feed spillage can also reduce 
fly numbers. Bedding choice can minimize stable fly development: 
sand, gravel, and sawdust are least conducive to stable fly devel-
opment while straw, corn, and soybean stalks are more conducive 
(Schmidtmann et al. 1989, Schmidtmann 1991).
Mechanical and Physical Control
Trapping can be effective for adult stable fly management. The 
same types of traps discussed in the Monitoring section can be 
used for management. Proper placement of traps, numbers of traps 
per unit area, and routine servicing are essential for success (Ose 
and Hogsette 2014). The Starbar Bite Free (Central Life Sciences, 
Schaumburg, IL) and the Knight Stick (BugJammer, Inc., Stockton, 
NJ) are two commercially available preglued stable fly traps. 
Compared with the Olson trap, the Bite Free prototype trap col-
lected 50% more stable flies (Taylor and Berkebile 2006) while the 
Knight Stick trap (Fig. 7) captured three times more stable flies and 
the glue on the sticky sleeves caught and held flies over a wider tem-
perature range (Hogsette and Kline 2017). Unfortunately, traps are 
expensive and labor-intensive. Effective trap usage can be achieved 
on dairies (Miller et al. 1993) and feedlots with proper budgeting 
and in-depth knowledge of seasonal stable fly patterns. Traps may be 
used more often to protect high-value animals such as equines and 
zoo animals. The strategy for managing stable fly adults in zoos is to 
minimize populations faster than they arrive, preferably without the 
use of pesticides. Evaluation of sanitation systems, e.g., elimination 
of animal wastes, is prudent, but in many cases, stable flies are not 
developing on-site (Ose and Hogsette 2014). Sticky Alsynite traps 
(Rugg 1982) and Knight Stick sticky traps (Kanzia 2015, Hogsette 
and Ose 2017) have been used for stable fly management in zoos. In 
Costa Rica, where thousands of adhesive traps are employed around 
pineapple farms to control stable flies, traps are made from white 
plastic trash bags painted with an adhesive (Solórzano et al. 2015; 
Fig. 8).
Biological Control
Predators
Predators, including macrochelid mites and staphylinid and histerid 
beetles, have the most significant effect of all biological control 
agents on immature stable fly populations (Legner and Olton 1970, 
Smith et al. 1985, Azevedo et al. 2018). However, predators are diffi-
cult to mass rear, and the substrates in which stable flies develop are 
often ephemeral, limiting the ability of predators to colonize them 
(Seymour and Campbell 1993). Currently, only Carcinops pumilio 
(Erichson) (Coleoptera: Histeridae) is sold commercially.
Parasitoids
Microhymenopteran parasitoids from 6 families parasitize stable 
flies (Table 1). Natural parasitism varies from 0 to 10%, usually early 
in the season, to as high as 70–80% at the end of the season (Greene 
et al. 1989). The wasp species parasitizing stable flies vary depending 
upon region and substrate. Spalangia species are the predominant 
parasites of stable flies in most environments, with Spalangia 
cameroni Perkins, Spalangia endius Walker, Spalangia nigroaenea 
Curtis, and Spalangia nigra Latreille  (Hymenoptera: Pteromalidae) 
being more prevalent (Pitzer et al. 2011b). Natural parasitism has 
an impact on stable fly populations. However, attempts to increase 
parasitism rates by augmentative releases of parasitoids have had 
mixed results. Releases of Muscidifurax raptor Girault and Saunders 
(Hymenoptera: Pteromalidae) and S. nigroaenea on dairies and feed-
lots showed minimal to no impact on stable fly populations (Miller 
et al. 1993, Andress and Campbell 1994, Skovgård and Nachman 
Fig. 7. Knight Stick sticky trap covered with stable flies.
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2004), while a later study showed the release of S. cameroni on four 
organic dairy farms in Denmark reduced the densities of both house 
flies and stable flies (Skovgård 2004).
Entomopathogenic Bacteria
Generally, known insect pathogens fail to produce infections in stable 
flies, yet studies to identify bacteria pathogenic to stable flies con-
tinue. Such properties were demonstrated for Serratia marcescens, 
Aeromonas sp., and Pseudomonas aeruginosa (Lysyk et  al. 2002, 
Lysyk et  al. 2010, Lysyk et  al. 2012, Lysyk and Selinger 2012). 
Additionally, five Bacillus thuringiensis (Bt) isolates with high tox-
icity to stable fly immatures were identified from a screen of 85 iso-
lates (Lysyk et al. 2010). Bt isolates controlled 87–99% of larvae at 
cooler temperatures (15°C) but lost some efficacy at warmer temper-
atures (30°C); the isolates were also less effective against 2nd and 3rd 
instars (Lysyk and Selinger 2012). Of interest, isolate B. thuringiensis 
thompsoni 401 consistently caused adult stable fly mortality when 
fed inblood and when applied topically (Lysyk et al. 2012).
Entomopathogenic Fungi
Metarhizium anisopliae has been commercialized as a biopesticide 
for fly control. M. anisopliae is virulent to stable fly eggs but not 
to larvae or pupae (Moraes et  al. 2008). An aqueous formula-
tion (Ma134) applied to naturally infested Holstein cows reduced 
stable fly populations and cattle defensive behaviors (Cruz-Vasquez 
et al. 2015). Another Metarhizium formulation using Metarhizium 
brunneum (Met52 EC) deterred stable fly oviposition (Machtinger 
et al. 2016b), and Weeks et al. (2017) observed effective adult stable 
fly mortality when exposed to Met52 EC. Strains of Beauveria 
bassiana have also been commercialized as a biopesticide (e.g., 
Mycotrol O, BotaniGard ES, balEnce); however, these were less 
effective against stable flies than Met52 EC (Machtinger et  al. 
2016b, Weeks et  al. 2017). Moraes et  al. (2014, 2015) identified 
Stenotrophomonas maltophilia as a bacterium in stable fly larval 
mucus with antifungal activity against B. bassiana; this suggests that 
larval microbiota can compromise the use of B. bassiana fungus to 
control stable fly larvae.
Entomopathogenic Nematodes
Nematodes in the families Heterorhabitidae and Steinernematidae 
can infect and reproduce in stable flies in laboratory studies (Clark 
2001, Mahmoud et al. 2007, Pierce 2012, Leal et al. 2017). Infective 
juvenile (L3) nematodes survive for up to 10 wk in manure substrates 
in the laboratory (Clark 2001). However, in the field, infective nema-
todes could not be isolated with Galleria mellonella (L.) (Lepidoptera: 
Pyralidae)  larvae 24  h after treatment (D.B.T., unpublished data). 
Steinernema feltiae (Filipjev) (Rhabditida: Steinernematidae)  was 
the most virulent species in comparative studies (Clark 2001, Pierce 
2012), but Leal et al. (2017) found Heterorhabditis bacteriophora 
Poinar  and Heterorhabditis baujardi Phan, Subbotin, Nguyen 
and Moens (Rhabditida: Heterorhabditidae)  to be highly viru-
lent as well. Pierce (2012) observed significantly fewer stable flies 
emerging from treated substrates relative to untreated substrates in 
year one of a two-year study, suggesting the potential for the use of 
entomopathogenic nematodes of the genus Heterorhabditis for the 
control of stable flies.
Chemical Control
Chemical agents registered to control of adult and immature stable 
flies are general use products applied against other biting flies, lice, 
or ticks. Products registered for stable flies vary based on the target 
animal, treatment site (animal/manure/structure) and the specific in-
secticides available vary by state (https://www.veterinaryentomology.
org/vetpestx). The products contain various active ingredients, but 
most are pyrethroids, often permethrin. These products may be ap-
plied as diluted concentrates delivered as sprays to the animal, as 
a ready-to-use pour-on products applied along the backline of an 
animal, or as diluted concentrates applied as indoor and outdoor 
premise sprays targeting the resting sites of stable flies. For control 
of biting adults on range and pasture cattle, area treatments with 
ultralow volume (ULV) insecticides can provide short-term relief, 
usually less than 24 h (Foil and Hogsette 1994), depending on the 
distance from fly breeding sites. Forced self-treatment devices such as 
the 3-D Quik Hand (3-D Cattle Equipment, LLC, Pine Ridge, AR), a 
motion-activated travel sprayer, are an option for high-valued stock.
Efficacy of on-animal applications for stable fly control is often 
poor and/or short-lived because the flies’ preferred feeding site is 
quite distant from the insecticide application site unless the lower 
legs are specifically and thoroughly sprayed. Ear tags are not ef-
fective against stable flies because of the fly’s preference for feeding 
on the animals’ lower legs. Furthermore, the lower legs are often wet 
or dirty, causing insecticide residues to be washed away or covered 
up, respectively. The window for insecticide contact is quite limited 
because the stable fly only visits host animals when feeding.
Fig. 8. White plastic bags covered with adhesive and positioned in fields near agricultural farms in Costa Rica are used to help reduce a massive stable fly 
infestations originating in pineapple crop residues.
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Table 1. Microhymenopterous parasitoids of stable flies
Family Species Location References
Pteromalidae Muscidifurax spp. South Africa, Florida, Missouri, 
Illinois,Nebraska,
(Legner and Olton 1968, Meyer and Petersen 1982, 
Petersen and Meyer 1983, Smith et al. 1987, Greene 
et al. 1989, Olbrich and King 2003, Romero et al. 
2010)
 M. raptor Lab, Denmark, Australia, Florida, 
Canada, California, Illinois, Indiana, 
Nebraska, 
Hungary, Australia
(Legner 1966, Legner et al. 1967, Legner and Olton 
1968, King 1990, Meyer et al. 1990, Meyer et al. 
1991, Hogsette et al. 1994, Jones and Weinzierl 
1997, Skovgård and Jespersen 1999, Hogsette et al. 
2001, Gibson and Floate 2004, Lysyk 2004, Geden 
et al. 2006, Pitzer et al. 2011b)Hogsette unpubl.
 Musidifurax raptorellus Lab, Costa Rica (Geden and Moon 2009) Taylor unpubl.
 Musidifurax zaraptor Lab, California, Illinois, Nebraska (Meyer et al. 1990, Meyer et al. 1991, Jones and 
Weinzierl 1997, Lysyk 2004) Taylor unpubl.
 Spalangia cameroni Lab, Japan, Germany, Denmark, Is-
rael, Kenya, Uganda, South Africa, 
Mauritius, Australia, New Zealand, 
Philippines, Hawaii, Florida, Can-
ada, Illinois, California, Nebraska, 
Indiana, Hungary
(Legner 1966, Legner et al. 1967, Legner and Olton 
1968, Meyer and Petersen 1982, Petersen and Meyer 
1983, Greene et al. 1989, King 1990, Meyer et al. 
1990, Meyer et al. 1991, Hogsette et al. 1994, Jones 
and Weinzierl 1997, Skovgård and Jespersen 1999, 
Hogsette et al. 2001, Skovgård and Steenberg 2002, 
Olbrich and King 2003, Gibson and Floate 2004, 
Geden et al. 2006, Romero et al. 2010, Pitzer et al. 
2011b, Machtinger et al. 2016a, Matsuo 2020)
 S. endius Lab, Japan, Florida, Brazil, Israel, 
Kenya, Uganda, South Africa, 
Mauritius, Australia, New Zealand, 
Philippines, Missouri, Illinois, Cali-
fornia, Nebraska, Indiana, Hungary
(Legner 1966, Legner et al. 1967, Legner and Olton 
1968, Meyer and Petersen 1982, Petersen and 
Meyer 1983, Smith et al. 1987, Greene et al. 1989, 
King 1990, Meyer et al. 1990, Meyer et al. 1991, 
Hogsette et al. 1994, Jones and Weinzierl 1997, 
Hogsette et al. 2001, Olbrich and King 2003, Geden 
et al. 2006, Romero et al. 2010, Brandão et al. 2011, 
Pitzer et al. 2011b, Matsuo 2020)
 Spalangia haematobiae Missouri, Florida (Smith et al. 1987, Romero et al. 2010)
 Spalangia gemina Lab, Costa Rica, Australia (Geden et al. 2006)Hogsette unpubl.,Taylor unpubl.
 Spalangia longepetiolata Kenya, Uganda (Legner and Olton 1968, Legner and Greathead 1969)
 Spalangia nigripes Denmark (Skovgård and Jespersen 1999, Skovgård and 
Steenberg 2002)
 Spalangia nigroaenea Lab, Japan, Germany, Uganda, South 
Africa, Mauritius, New Zealand, 
Philippines, Florida, Canada, Mis-
souri, Illinois, California, Nebraska, 
Texas, Kansas, Louisiana, Hungary, 
Australia
(Pinkus 1913, Legner 1966, Legner et al. 1967, 
Legner and Olton 1968, Hogsette 1981, Meyer and 
Petersen 1982, Petersen and Meyer 1983, Smith 
et al. 1987, Greene et al. 1989, Meyer et al. 1990, 
Meyer et al. 1991, Hogsette et al. 1994, Jones and 
Weinzierl 1997, Hogsette et al. 2001, Olbrich and 
King 2003, Gibson and Floate 2004, Geden et al. 
2006, Romero et al. 2010, Pitzer et al. 2011b, 
Matsuo 2020) Hogsette unpubl.
 S. nigra Denmark, Florida, Canada, Missouri, 
Illinois, Nebraska
(Meyer and Petersen 1982, Petersen and Meyer 
1983, Smith et al. 1987, Jones and Weinzierl 1997, 
Skovgård and Steenberg 2002,Olbrich and King 
2003, Gibson and Floate 2004, Romero et al. 2010, 
Pitzer et al. 2011b)
 Pachycrepoideus 
vindemiae
Denmark (Skovgård and Jespersen 1999)
 Sphegigastersp. Uganda, South Africa (Legner and Olton 1968, Legner and Greathead 1969)
 Trochomalopsis sp. 
Trichomalopsis dubius
Hungary, Australia 
Illinois
(Hogsette et al. 2001)Hogsette unpubl. 
(Olbrich and King 2003)
 Trichomalopsis 
sarcophagae
Lab (Lysyk 2004)
 Trichomalopsis 
viridescens
Florida (Romero et al. 2010)
 Urolepis rufipes Canada, Illinois, California (Meyer et al. 1991, Olbrich and King 2003, Gibson 
and Floate 2004)
Ichneumonidae Phygadeuonsp. Ireland (Legner and Olton 1968)
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Insecticide-treated blue-black fabric targets attract and kill 
stable flies (Foil and Younger 2006). When impregnated with 
0.1% lambda-cyhalothrin or 0.1% zeta-cypermethrin, targets in 
northern Florida remained effective in the field for four months 
(Hogsette et al. 2008). Hogsette and Foil (2018) determined that 
small, cylindrical (30  cm high × 63  cm circumference) targets 
made of blue, black, and blue and black fabric worked equally 
well. The average stable fly resting time on the targets is 30  s, 
which is long enough to acquire a lethal dose of the insecticide. 
Similarly, insecticide-impregnated screens (Nagagi et  al. 2017, 
ZeroFly) can be located on the perimeter of indoor or outdoor 
animal units where stable flies are likely to rest while entering 
to feed or leaving after feeding. An advantage of targets and 
screens over traps is that they do not saturate or fill up with cap-
tured insects; hence they do not need regular servicing. However, 
screens must be cleaned regularly to remove accumulated dust 
occluding the surface, limiting the contact and effectiveness of the 
insecticide-treated surface (D.B.T. and J.A.H., unpublished data). 
The primary disadvantage of targets is perception, as users are 
accustomed to seeing captured flies in or on traps, but targets do 
not have collection devices. Flies falling off the targets may blow 
in the wind, be carried away by scavengers, or die shortly after 
leaving the target, leaving no obvious evidence of their demise.
Larvicides
Chemicals targeting stable fly larvae and development substrates 
of stable flies have been investigated. Liquid and granular larvicide 
applications of pyriproxyfen and buprofezin, insect growth regu-
lators, to substrates inhibited immature stable fly development 
(Liu et  al. 2012), and cyromazine reduced adult stable fly emer-
gence (Taylor et  al. 2012b, Donahue Jr et  al. 2017). Novaluron 
(benzoylphenyl urea insecticide) applied to manure (Lohmeyer and 
Pound 2012) or hay feeding sites (Taylor et al. 2014, D.B.T., un-
published data) resulted in a significant reduction in adult emer-
gence, as did novaluron delivered as a feed-through (Lohmeyer 
et al. 2014). Ideally, IGRs should be applied 3–5 d after clean-out 
of barns and calf pens or 2-3 wks before expected adult emergence 
from hay feeding sites and other pasture sites. IGRs should remain 
active for 10–12 wk (Taylor et al. 2012b). Because cyromazine and 
novaluron have different modes of action (Insecticide Resistance 
Action Committee (IRAC) groups 17 and 15 respectively), proper 
rotation should inhibit the development of resistance. Field trials 
in Costa Rica found triflumuron, diflubenzuron, novaluron, and 
cyromazine to all be effective larvicides in postharvest pineapple 
stubble (Solórzano et al. 2013).
Managing Resistance
Pyrethroid resistance in stable flies has been documented in sev-
eral countries. In the United States, 3.5–12-fold resistance to per-
methrin was described in stable flies collected from Florida dairies 
and equine farms (Pitzer et  al. 2010), while flies occupying sugar 
cane production sites in Brazil exhibited 6.8–38-fold resistance to 
cypermethrin (Barros et al. 2019). Analysis of the gene encoding the 
voltage-sensitive sodium channel, a target site for pyrethroids, iden-
tified single nucleotide polymorphisms that are associated with the 
knockdown resistance (kdr) phenotype. Stable flies from the U.S. and 
Brazil have a mutation resulting in a leucine to histidine substitution 
at this position (L1014H; kdr-his; Olafson et al. 2011, Egito 2020). 
Resistance to cypermethrin, deltamethrin, lambda-cyhalothrin, and 
permethrin was observed in stable flies from livestock farms in 
France (Salem et al. 2012b, Tainchum et al. 2018), and flies from 
one of these regions (Toulouse) harbored a leucine to phenylalanine 
(L1014F) kdr mutation (Olafson et al. 2019). Reissert-Oppermann 
et al. (2019) described deltamethrin-resistant populations of stable 
flies in northeast Germany. While metabolic detoxification mechan-
isms contribute to pyrethroid resistance in both house flies and horn 
flies, its role has not been studied in pyrethroid-resistant stable fly 
populations.
Natural Products/Bio-Pesticides
Insecticidal and behavior-modifying properties of plant by-products, 
i.e., plant extracts, essential oils, and fatty acids, are promising alter-
natives to synthetic insecticides, especially for control of biting flies 
(Showler 2017). The essential oil of catnip or catmint, Nepeta cataria, 
and its dehydrogenation by-products are toxic to adult stable flies 
(Zhu et  al. 2011), and are effective at deterring blood-feeding and 
oviposition (Feaster et al. 2009, Zhu et al. 2009, Zhu et al. 2010, Zhu 
et al. 2012). Larval development is also prevented probably because of 
antibacterial activity (Zhu et al. 2014). Essential oils and constituents 
of Zanthoxyllum sp., specifically Zanthoxyllum piperitum (Japanese 
pepper) and Zanthoxyllum armatum (winged prickly ash), are also 
toxic to stable fly adults, albeit several orders of magnitude less ef-
fective than synthetic organophosphates (Hieu et al. 2012). Further, 
Zanthoxyllum essential oils reduce stable fly electroantennogram 
responses to host volatile compounds and repel stable fly adults in 
human hand bioassays (Hieu et al. 2010b, Hieu et al. 2014). Tamanu 
seed oil, isolated from Calophyllum inophyllum, synergizes the effect 
of these Zanthoxyllum essential oils and increases protection time 
to that comparable to DEET (Hieu et al. 2010b, Hieu et al. 2010a). 
Tamanu oil also synergizes repellency of essential oils from clove bud, 
clove leaf, thyme white, patchouli, or savory to stable flies at levels 
Family Species Location References
 Phygadeuon fumator Denmark, Florida, Canada (Skovgård and Jespersen 1999, Skovgård and 
Steenberg 2002, Gibson and Floate 2004, Romero 
et al. 2010)
 Diplazon laetatorius Missouri (Smith et al. 1987)
Braconidae Alysia manducator Uruguay (Legner et al. 1967)
Encyrtidae Tachinaephagus 
zealandicus
Lab, Australia, New Zealand (Legner and Olton 1968, Geden and Moon 2009)
Diapriidae Trichopriaspp. Missouri, Uganda, New Zealand, Phil-
ippines, Hungary, China
(Legner and Olton 1968, Smith et al. 1987, Hogsette 
et al. 1994, Guo et al. 1997)
 Trichopria nigra Lab (Geden and Moon 2009)
Chalcididae Dirhinus himalayanus Lab (Geden et al. 2006)
Table 1. Continued
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similar to DEET (Hieu et al. 2010a, Hieu et al. 2015). Lemongrass 
oil has been identified as an effective feeding deterrent against adult 
stable flies (Baldacchino et  al. 2013b). A  formulation comprised of 
straight chain saturated octanoic (C8), nonanoic (C9), and decanoic 
(C10) fatty acids, commercially available as C8910 (Stratacor Inc.), 
is repellent against stable flies (Mullens et al. 2009) and was regis-
tered by the Environmental Protection Agency in 2009 for use as a 
fly repellent on livestock. Further, these three straight chain saturated 
fatty acids(C8, C10, and C12) and their corresponding methyl esters 
isolated from hydrolyzed coconut oil deterred adult stable fly feeding 
in laboratory assays (Zhu et al. 2018), with the C8 methyl ester ex-
hibiting a spatial repellent effect and flies observed to be paralyzed 
(knocked down) during feeding tests using the C8, C10, and C12 me-
thyl esters (Roh et al. 2020). While significant progress has been made 
in evaluating insecticidal and behavior-modifying properties of plant 
compounds towards stable flies, the residual activity is short (24 h or 
less) and would require repeated application. As a result, commercial-
ization of these products for stable fly control is limited. Lower cost, 
longer active on-animal repellents are needed for more general use on 
pastured cattle (Zhu et al. 2018), but range and pasture management 
systems will require re-application intervals to be 1–3 wk if on-animal 
repellents are to become operational.
Novel Technologies for Stable Fly Management
Targeting Olfaction
Stable fly adult trapping systems focus on adult stable fly visual at-
traction to various materials (reflectance, color, etc.). Improvements 
to these systems may occur if supplemented with chemical attract-
ants (Zhu et al. 2016). Identifying compounds effective at “pulling” 
flies away from their host has bolstered this effort (Cook et al. 2007). 
Stable flies were exposed to host-derived volatiles of Holstein-
Friesian cattle, and compounds that elicited an electroantennogram 
response included naphthalene, p-cresol, octenol, 2-heptanone, 
2-decanol, sulcatone, linalool, and citronellol (Birkett et al. 2004). 
Jeanbourquin and Guerin (2007a) further isolated volatiles from 
the rumen contents of grass-fed steers and identified dimethyl 
trisulphide, butanoic acid, and p-cresol as attractive to stable flies 
using wind tunnel experiments. Horse and cow dung were assessed 
for attractiveness to adult stable flies. While flies preferred to ovi-
posit on horse dung, no difference in electroantennogram response 
was observed when flies were exposed to dung from either host. 
Stable flies responded to twenty-five compounds found in both 
the cow and horse dung, with the highest response to dimethyl 
trisulphide (Jeanbourquin and Guerin 2007b). Cattle manure 
slurry has also been evaluated for volatile compounds attractive 
to stable flies, and blends of phenol, p-cresol, and m-cresol were 
most attractive (Tangtrakulwanich et  al. 2011). Understanding 
stable fly responses to semiochemicals informs trap enhancement. 
Preliminary descriptions of gene families involved in chemosensory 
recognition and signal transmission were identified from stable fly 
expressed sequence tag databases (Olafson et  al. 2010). Further, 
the highly conserved insect odorant co-receptor, Orco, was isolated 
and the spatial and temporal transcript and protein expression pat-
tern described (Olafson 2013), providing the basis for studies to 
identify chemosensory targets for behavior-modifying strategies.
Targeting Neuroactive Molecules
Insect neuroactive molecules are integral to pathways regulating vital 
functions, including feeding, excretion, and reproduction. In the stable 
fly, periviscerokinin/cardioacceleratory (PVK/CAP2b) neuropeptides 
were identified that stimulate Malpighian tubule fluid secretion 
(Nachman et  al. 2002, Nachman et  al. 2006), as well as a factor 
similar to the natriuretic peptide that regulates sodium ion excretion 
in the primary urine (Chen et al. 1997). Liu et al. (2013) observed 
a reduction in sperm transfer by reproductive stable fly males and 
reduction in oviposition by gravid females who had been separately 
treated with reserpine, a chemical that depletes biogenic amines. In 
males, this was likely due to a transient reduction of serotonin in the 
24h after treatment as serotonin immunoreactivity was depleted in 
neurons innervating the testes during this period (Liu et  al. 2013). 
Serotonin also plays a critical role in stable fly larval and adult feeding, 
as the cibarial pump during both these life stages is innervated by 
serotonin immunoreactive neurons (Liu et  al. 2011). The design of 
mimetic analogs to target these functions by disrupting their signaling 
pathways is an attractive avenue for control technology development.
Mining Stable Fly-Omics Resources
Advancements in nucleic acid and protein sequencing technolo-
gies provide opportunities to broaden our understanding of stable 
fly biology by examining its genome, transcriptome, and proteome. 
MicroRNAs are a class of molecules that affect gene expression by 
repressing messenger RNAs post-transcriptionally, regulating bio-
logical processes from metamorphosis to feeding and reproduction. 
Stable fly microRNAs expressed during immature and fed adult 
stages were identified by Tuckow et  al. (2013), but their function 
and role remain unknown. Future studies to characterize stage- and 
tissue-specific expression of microRNAs and to identify those that 
are of interest for disruption would inform the development of 
microRNA mimetics that can be developed as a control technology, 
as in Zhang et  al. (2015). The recent sequencing of the stable fly 
genome sequence provides an additional resource to further define 
those pathways that regulate critical behaviors (Olafson et al. 2021). 
Future studies will require functional characterization of these gene 
family members to elucidate their importance to stable fly biology. 
For this, gene editing technologies such as CRISPR have shown 
promising application in muscid flies, and stable flies are amenable to 
genetic transformation, as demonstrated by O’Brochta et al. (2000).
Sterile Insect Technique
The sterile insect technique (SIT) is an area-wide pest control method 
based upon the mass release of sterilized insects that mate with na-
tive insects, rendering them sterile. The SIT method has been used 
to control and/or eradicate several species of insects including New 
World Screwworm (Cochliomyia hominivorax (Coquerel) [Diptera: 
Calliphoridae]; Wyss 2006) and Mediterranean Fruit Fly (Ceratitis 
capitata (Wiedemann) [Diptera: Tephritidae]; Enkerlin et al. 2017). 
In preliminary studies during the early 1970s in Florida, LaBrecque 
et al. (1972) reduced the population of wild stable flies significantly 
with the release of chemo-sterilized stable flies but were unable 
to eradicate locally because of immigration from the surrounding 
area. A program was initiated on the island of St. Croix, U.S. Virgin 
Islands in 1974 to demonstrate the feasibility of using SIT to eradi-
cate an isolated population of stable flies. Models were developed 
using data on the population dynamics of stable flies on St. Croix to 
develop an integrated strategy including SIT within the limitations 
of infrastructure and rearing capacity for the island (LaBrecque et al. 
1972). The stable fly population on St. Croix averaged 1.9 flies per 
animal (=950,000 flies on the island) during the wet seasons and 
0.5 flies per animal (=250,000 total flies on the island) during the 
dry season. Methods were developed to rear and sterilize ≈250,000 
stable flies per day (Williams et al. 1981). In addition to SIT, major 
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larval developmental areas were treated with 1% methoxychlor and 
the pupal parasitoid Spalangia endius was released at some of the de-
velopmental sites. The sterility of native flies increased to over 90% 
and numbers dropped by more than 99% after one year of sterile 
insect releases (Patterson et al. 1981).
Although the St. Croix pilot program was successful, SIT use for 
stable fly control is not presently a viable option (Taylor 2021). SIT 
programs usually target release rates of five to ten sterile insects for 
every native insect. The stable fly population on St. Croix at the time of 
the pilot programwas quite low, 0.4–6.0 flies per host animal, which 
is well below the economic threshold of 5 flies per front leg (≈15 flies/
animal) (Campbell and Berry 1989). During severe outbreaks, stable 
fly infestations can exceed 1,000 per host animal (Bishopp 1913a, 
Cook et al. 1999, Solórzano et al. 2015) and 15–50 per animal are not 
uncommon (Taylor et al. 2012a). It is unlikely that livestock produ-
cers or the public would tolerate the release of the required numbers 
of sterile stable flies given that both males and females bite and feed 
on blood up to two to three times daily (Harris et al. 1974), their bites 
are extremely painful, and their predilection to bite humans when live-
stock are not available. An effective method requiring the release of 
much smaller numbers of stable flies is desirable.
Current Needs to Improve Stable Fly 
Management
Management
The economic thresholds in use to make management decisions 
today were developed in the early 1980s. Since then, factors that 
affect the impact of stable flies on animals, like animal nutrition and 
genetics, have evolved. Monitoring methods adapted to today’s hus-
bandry practices are essential to allow producers to make quick and 
reliable decisions for the welfare of their animals.
• Develop a standard for monitoring adult stable fly populations 
and reaching an economic threshold.
• Re-evaluate economic thresholds using modern production 
methods and livestock.
• Develop monitoring methods that do not involve a host animal.
In contast, trapping technologies have changed little over the last 
thirty years. Traps generally remain less attractive than host animals 
and require a fair bit of maintenance. We still do not have a solid 
grasp of what attracts adult stable flies to the hosts, resting sites, or 
even trapping devices.
• Design traps that are more competitive with animal hosts.
• Develop attract-and-kill or capture devices that are efficacious but 
not labor-intensive, e.g., virtual counters connected to phone apps.
• Develop traps that simulate stable fly adult resting sites, e.g., 
shrubbery.
• Determine how flies perceive traps to exploit these tendencies.
The crux of successful stable fly management resides in destroying 
larval substrate. However, we cannot quickly and reliably locate pro-
ductive immature fly development sites or estimate larval numbers 
within these sites. More research is required to improve knowledge 
of immature development substrates.
• Characterize microbial communities associated with develop-
mental substrates.
• Develop sensors to identify sub-surface larval activity.
• Develop pesticide-free methods for preventing stable fly develop-
ment in larval substrates.
• Develop better understanding of larval chemical ecology and 
orientation within their habitat.
Biology and Life History
Dispersal by adult stable flies leads to management problems be-
cause the arrival of adults from off-site locations can create the ap-
pearance that on-site management techniques are inadequate. When 
few immatures are found on-site, adult abundance is difficult to 
explain. Therefore, dispersal and the factors affecting it need to be 
better defined and integrated into management decisions.
• Identify factors necessary for predicting pest-level population 
dispersal.
• Develop methods to associate adult stable flies with their devel-
opmental substrate to gain insight into the sources of pest popu-
lations.
• Explain adult stable fly abundance on facilities and its relation-
ship to larval abundance.
• Determine dispersal patterns of stable flies, including both mean 
flight range and mechanisms of long-range flight.
• Determine the role of weather patterns in stable fly dispersal and 
the influence of climate change on stable fly populations.
• Assess whether genome-wide sampling techniques effectively dis-
tinguish stable fly populations, providing a tool for interpreting 
dispersal events.
Efforts to define permethrin resistance in stable fly populations have 
gained some traction within the last decade. However, there are still 
gaps in our understanding of the extent of resistance to this class of 
insecticides and whether resistance to other classes of insecticides is 
circulating.
• Evaluate stable fly resistance to other classes of insecticides.
• Define nationwide levels of permethrin resistance.
• Investigate whether mechanisms other than target-site insensi-
tivity may contribute to the permethrin-resistant phenotype.
Methods are currently being developed and evaluated that would use 
genetic drive mechanisms to suppress insect populations, particularly 
those of importance to animal and human health. These methods 
would require the release of smaller numbers of genetically modified 
insects to “drive” a deleterious trait into the population, ultimately 
reducing the size of the native population. Importantly, such methods 
involve persistent genetic modifications to the population (Carter and 
Friedman 2016) and are pending societal acceptance.
• Explore the use of gene-drive constructs that would specifically 
target stable fly population replacement.
Acknowledgments
We thank the members of the S-1076: Multistate research project Fly Man-
agement in Animal Agriculture Systems and Impacts on Animal Health and 
Food Safety for their support and review of this manuscript, especially Erika 
Machtinger, Doug Ross, Roger Moon, Alec Gerry, Jeff Scott, Kyle Harrison 
and Luisa Domingues for helpful comments and suggestions. We also thank 
Matt Bertone for providing the beautiful photos of the adult stable fly and 
house fly, stable fly larva, and close-up of spiracles. Thanks to Michael Behrens 
for technical assistance. This multi-state extension work is/was supported by 
the United States Department of Agriculture National Institute of Food and 
Agriculture Extension Smith Lever funding under Project number PEN04540 
and Accession number 1000356 and by the various Multistate Hatch projects 
from collaborators.
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