Genetic variability of the stable fly, stomoxys calcitrans(l) (diptera: muscidae) over a geographical area

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The stable fly, Stomoxys calcitrans (L) (Diptera: Muscidae) is a cosmopolitan ectoparasite of livestock, wildlife and humans (Brues 1913; King and Lenert 1936; Simmons 1944; Hansens 1951; Berry and Campbell 1985; Mullens and Meyer 1987; Meyer and Shultz 1990; Thomas et al. 1990; Skoda et al. 1991; Skoda and Thomas 1993; Campbell 1995; Campbell et al. 2001; Kaufman 2002; Veer et al. 2002; Jeanbourquin and Guirin 2007; Taylor and Berkebile 2008). Both sexes are hematophagous (Brain 1912; Skidmore 1985; Campbell 1995), and feed primarily on the legs of the host animals (Skoda and Thomas 1995; Campbell et al. 2001; Mullens and Peterson 2005; Mullens et al. 2006). Stable flies react to both olfactory and visual stimuli for the location of hosts (Gatehouse 1967; Allan et al. 1987; Alzogaray and Carlson 2000; Carlson et al. 2000; Birkett et al. 2004). They will feed several times per day (Powell and Barringer 1995; Mullens et al. 2006), and at least one blood meal is required for reproduction (Skidmore 1985). Oviposition occurs on decaying organic matter such as spilled hay or grain, preferably combined with feces (Berkebile et al. 1994,1995). Stable fly parasitism has the greatest effect on the livestock industry, where animals are confined to stables or pastures, providing a pristine environment for both feeding and oviposition (Berkebile et al. 1994; Campbell 1995; Hogsette 1998; Broce et al. 2005). Their painful bite stresses confined livestock, causing them to bunch together or perform repellent behaviors, which results in significant reductions in weight gain and milk production (Campbell et al. 1977; Hall et al. 1983; Catangui et al. 1993). The stable fly is the primary pest of livestock in the United States, causing major annual economic losses estimated to be greater than $1billion to cattle in feedlots and dairies as well as poultry farms (Suszkiw and Core 2003; Taylor and Berkebile 2006; Roeder 2007).

Stable fly outbreaks also occur along beaches, causing considerable economic damage to the tourist trade (King and Lenert, 1936; Simmons and Dove 1941, 1942; Dove and Simmons, 1942; Simmons 1944; Hansens, 1951; Williams and Rogers 1976; Hogsette and Ruff 1985; Jones et al. 1991; Koehler and Kaufman 2006). In Northwest Florida they migrate on the north winds from inland livestock areas to the beaches (Fye et al. 1980), where they breed in marine grasses such as seaweed, turtle grass (Thalassia testudium), and manatee grass (Halodule wrightii) (King and Lenert 1936; Dove and Simmons 1942). They also breed in peanut litter (Simmons 1944) and waste celery (Simmons and Dove 1942). In New Jersey, stable flies were reported to breed in the marine grasses that washed onto shore, and outbreaks were concurrent with west winds (Hansens 1951).

In addition to their detrimental impact on livestock, the presence of stable flies causes legal issues between farmers and the urban population encroaching on the farmland (Meyer et al. 1990; Thomas and Skoda 1993; Campbell 1995; Suszkiw and Core 2003). Stable flies are also known to be mechanical vectors of disease (Brues 1913; Turell and Knudson 1987; Fischer et al. 2001; Veer et al. 2002; Szalanski et al. 2004; Bittencourt and De Castro 2004; Mramba et al. 2007). Due to the global distribution and adverse effects of stable fly activity, more efficient control measures are needed. Research has been carried out in areas such as chemical, biological and mechanical control mechanisms, Integrated Pest Management (IPM) practices, dispersal and overwintering, population genetics and gene flow, physiology, and DNA analysis (Campbell and Hermanussen 1971; Bailey et al. 1973; Black and Krafsur 1985; Berkebile et al. 1994; Campbell 1995; Szalanski et al. 1996; Ratcliffe et al. 2002; Skovgard and Nachman 2004; Broce et al. 2005; De Oliveira et al. 2005; Gilles et al. 2007; Taylor et al. 2007). However, no single method of stable fly control has been successful thus far. Current control methods have had no significant success in maintaining stable fly populations below the economic injury threshold (Patterson et al. 1981; Meyer et al. 1990; Clymer 1992; Hall 1992; Pickens 1992; Seymour and Campbell 1993; Andress and Campbell 1994; Cilek and Greene 1994; Campbell 1995; Weinzierl and Jones 1998; Guglielmone et al. 2004; Macedo 2004; Skovgård and Nachman 2004; Foil and Younger 2006; Taylor and Berkebile 2006; Gilles et al. 2007; Mihok and Carlson 2007), although Integrated Pest Management (IPM) practices help to reduce stable fly populations at the local scale (Campbell and Wright 1976; Lazarus et al. 1989; Campbell 1995; Skoda et al. 1996; Thomas et al. 1996).

Further research related to stable fly populations, origins of outbreaks, and dispersal patterns could lead to the development of more effective control strategies.

Life History and Biology

Stomoxys calcitrans belongs to the family Muscidae and subfamily Stomoxyinae, which includes stable flies, horn flies and buffalo flies (Zumpt 1973). There are 18 recognized species in the genus Stomoxys (Zumpt 1973), with S. calcitrans being the only species recorded in North America (Marquez et al. 2007). Stomoxyinae are characterized by their piercing proboscis and maxillary palpi. The proboscis is formed by three sclerotized parts: the labium, hypopharynx, and labrum. At rest, it is extended horizontally and can be seen beyond the head (Brain 1912). In S. calcitrans, the palpi are single-segmented and approximately ¼ the length of the proboscis (Brain 1912; Zumpt 1973). After puncturing the skin of the host, saliva is injected into the wound via the hypopharynx, then blood is drawn up into the pharynx via a tube composed of the hypopharynx and labrum combined (Brain 1912; Zumpt 1973). The mouthparts are alike in males and females, and both sexes are hematophagous (Brain 1912).

Stable flies take a blood meal several times per day, and are persistent feeders (Schofield and Torr 2002). Females require at least three blood meals for ovarian development, and daily blood meals thereafter (Moobola and Cupp 1978; Chia et al. 1982; Veer et al. 2002; Schofield and Torr 2002). Anderson (1978) reported that males require a blood meal to properly inseminate the females; taking a blood meal increases their virility and the aggressiveness of their mating behavior. In addition to blood meals, stable flies also feed on nectar. Lee and Davies (1979) reported that feeding on sugar increased stable fly longevity. Moobola and Cupp (1978) report that blood feeding, not sugar, increases longevity. However, they report that sugar will increase survival rate five times more than just water if no blood meals are available. Jones et al. (1992) reported that nectar feeding may supply energy for dispersing flies when no hosts are available to obtain a blood meal, but being fed sugars ad libitum may be detrimental to reproductive rate, even when given daily blood meals.

Female stable flies oviposit in moist, decaying organic matter such as pure manure (Brain 1912; Miller 1992; Hall 1992), silage, hay, grain or haylage mixed with manure (Berkebile et al. 1994; Campbell 2006), grass clippings, compost piles, dumpsters (Suszkiw and Core 2003), and seaweed (King and Lenert 1936). It has been shown that females are attracted to substrates with active microbial communities, because certain bacterial species, such as Citrobacter freundii, may aid in larval development (Romero et al. 2006). The female lays 100-400 eggs during her lifetime, at approximately 20 eggs per ovarian cycle. Two blood meals are required for each cycle (Skidmore 1985; Campbell 1997).

Stable fly development is holometabolous, consisting of the egg, 3 larval instars, pupa and adult (Zumpt 1973; Skidmore 1985). The eggs are white, about 1mm long, convex ventrally with a longitudinal groove. They hatch in 2-4 days (Brain 1912; Zumpt 1973; Skidmore 1985). Larvae grow to about 10 mm, and the larval stage lasts 2-3 weeks under normal conditions, but unfavorable weather conditions may extend it up to 80 days (Brain 1912; Skidmore 1985). Larvae migrate to drier areas of the substrate to pupariate; pupariation lasts from 2-30 days. Puparia are brown in color and approximately 6 mm long (Brain 1912; Skidmore 1985). Adults are about 7 mm in length, with 4 black longitudinal stripes on the thorax, and a checkerboard pattern of dark spots on the abdomen (Brain 1912; Zumpt 1973; Skidmore 1985). They can take a blood meal within hours of emergence (Skidmore 1985).

The development period from egg to adult is dependent on temperature. (Melvin 1931; Simmons 1944; Kunz et al. 1977; Watson et al. 1994; Campbell 1997; Lysyk 1998; Campbell and Thomas 1999; Gilles et al. 2005a,b; Barker et al. 2007). Melvin (1931) studied the development of stable flies in the laboratory at 25ºC and 30ºC. He reported the incubation period of eggs to be 32.5-35.2 hours (mean 33.4) at 25ºC, and 25.0-28.5 hours (mean 26.5) at 30ºC. Combined larval and pupal periods were observed on 2 different rearing media. On alfalfa meal and wheat bran, mean development time at 25ºC was 377 hours (15.7 days), and 311.7 hours (13 days) at 30ºC. Mean development time at 30ºC on ground oats took 320.2 hours (13.3 days) and 326.1 (13.6 days) hours in two experiments. Melvin had difficulty rearing the flies, with only 10% adult emergence.

Simmons (1944) observed stable fly development under laboratory conditions, incubating eggs at 28ºC, and larvae and pupae at 30ºC. Minimum observed time before egg hatch was 19 hr, maximum 120 hr, mean minimum 39.65 hr, mean maximum 65.1 hr., and overall mean time until hatch was 52.3 hr. Duration of larval development was recorded as overlapping instars. First instars were present from egg hatch to the 80th hr, 2nd instar from the 44th-144th hr, and third instar from the 97th hr until pupation. Minimum time until pupation was 148 hrs. For mean calculations, the larval and pupal stages were combined, with a mean developmental duration of 165.8 hrs, or 6.9 days. Separating out the pupal stage, the mean duration of this stage was 6.55 days at 28º-32ºC. The reported life cycle from egg to adult was a minimum of 13 days, with a maximum of several months under adverse climatic conditions. Simmons (1944) reported that the duration of the larval period was longer (11.2 days) during winter months, even though the temperature was sustained at 30ºC.

Kunz et al. (1977) studied development at 3 different temperatures. Mean duration of development from egg to adult emergence was 400 hrs (16.6 days) at 23.9ºC, 280 hrs (11.6 days) at 29.4ºC, and 290 hrs (12.1 days) at 35.0ºC.

Lysyk (1998) studied the relationship between temperature and life history, rearing stable flies at 15, 20, 25, 30, and 35ºC. The observed median immature development times ranged from 62 days at 15ºC-<12 days at 30ºC, with development at 20ºC being 29 days.

Gilles et al. (2005a) studied the effect of temperature on developmental time of Stomoxys calcitrans and S. niger. The mean development time observed for S. calcitrans from egg to adult was 70.66 days at 15ºC, 32.36 days at 20ºC, 16.65 days at 25ºC, 12.92 days at 30ºC and 13.17 days at 35ºC. Adult longevity (Gilles et al. 2005b) was observed to be highest at 20ºC: 23.73 days for females and 25.69 days for males.

The results of these studies suggest that the developmental time of S. calcitrans from oviposition to adult emergence ranges from 11 days to several months, depending on the ambient temperature.

Distribution and Climatic Variables

Stomoxys calcitrans is native to Palaearctic regions of the Old World, and is now distributed worldwide, where it is most abundant in temperate regions (Brues 1913; Zumpt 1973; Skidmore 1985; Szalanski et al. 1996). It likely arrived in North America with the immigrants from Europe, and is reported to have been abundant in Philadelphia as early as 1776 (Brues 1913). Distribution patterns vary with climate, with precipitation and temperature having significant effects on population dynamics (Cruz-Vazquez et al. 2004; Mullens and Peterson 2005; Rodriguez-Batista et al. 2005; Taylor et al. 2007). In the Midwestern United States, stable fly populations follow a bimodal pattern of seasonal activity. They begin to appear in late March or early April and increase in numbers until they peak at the end of June. During the warmest part of summer the numbers recede, then peak again in mid-September (Mullens and Peterson 2005; Taylor et al. 2007). In California, the population peaks only once, in the late spring, but an active population remains throughout the year (Mullens and Peterson 2005). A study in Brazil showed stable fly activity during the spring and summer, (the time of year with the most precipitation) with a peak during November and December, and a smaller peak at the beginning of fall; there was no activity during the winter months. The results of this study suggested that stable fly population increases were related to rainfall (Rodriguez-Batista et al. 2005). A study in an arid region of Mexico, however, found no correlation between rainfall and stable fly populations. Instead, their results showed that the increase in population was correlated with relative humidity, and temperature was the primary factor in decreasing populations (Cruz-Vazquez et al. 2004).

The results of these studies suggest that climatic factors such as rainfall, relative humidity, and temperature all have an effect on stable fly populations. In the warmer climates populations declined during midsummer, showing sensitivity to high temperatures (Mullens and Peterson 2005). The correlation between population increases and rainfall or relative humidity indicates that stable flies require sufficient moisture to survive. Therefore climatic variables need to be considered in any population studies.

Stable Fly Parasitism of Livestock

Stable flies are the most important arthropod pest of livestock in the United States (Campbell 1995; Roeder 2007). They feed mainly on the legs of host animals, and the bite is extremely painful. Brain (1912) described the bite as having “a decided stab after the first puncture had been completed”. The pain and annoyance to cattle results in economic losses due to reductions in weight gain, feed efficiency and milk production, as well as the expense of control measures (Hall et al. 1983; Campbell 2006; Mullens et al. 2006; Roeder 2007).

Many studies have been performed on the effect of stable flies on the weight gain of cattle (Campbell et al. 1977; Catangui et al. 1993, 1995, 1997; Campbell et al. 2001; Broce et al. 2005), and the economic injury level on cattle in feed lots is estimated to be an average of five flies per front leg (Campbell et al. 2001). Cattle have developed repellent behaviors to dislodge the flies, such as leg stamps, tail and ear flicking, skin twitches and head throws (Campbell 1997; Schofield and Torr 2002; Mullens et al. 2006). They will also bunch together or stand in water in an attempt to escape the fly annoyance (Campbell 1995; Campbell and Thomas 1999). The bunching behavior may cause heat stress, which adds to the overall discomfort of the animal (Campbell 1995). Being stressed by the flies and engaging in repellent and avoidance behaviors, the cattle do not feed, which results in decreased weight gain from .1 pound to .48 pound per head per day, and a decrease in milk production of up to 40 percent (Campbell 1995).

Stable flies were originally considered pests of cattle confined to feed lots, but they are now recognized as a pest of pastured cattle as well (Campbell et al. 2001; Mullens et al. 2006). There are numerous breeding sites on feed lots, such as drainage areas, the edge of holding ponds, in areas where manure and soil or spilled feed can accumulate, such as along fences, in corners of pens, and at the edge of feed handling and storage areas (Campbell 1997). In pasture environments immatures have been found under large round bales or where the round bales are distributed and a portion of the hay is wasted. The wasted hay mixes with manure and urine, and if the bales are placed in the same area consistently, the substrate becomes attractive as a breeding site for the stable flies (Hall et al. 1982; Berkebile et al. 1994; Broce et al. 2005. Detritus from large round bales also provides a competent site for overwintering (Berkebile et al. 1994).

Control Methods

Many types of control methods for the stable fly have been researched, including insecticides, baits, biological control, and sterile release methods (Campbell and Hermanussen 1971; Campbell and Wright 1976; Campbell and Doane 1977; LaBrecque et al. 1981; Patterson et al. 1981; Williams et al. 1981; Gersabeck et al. 1982; Black and Krasfsur 1985; Andress and Campbell 1994; Hammack and Hesler 1996; Bartlett and Staten 1996; Floate et al. 2001; Ratcliffe et al. 2002; Guglielmone et al. 2004; Kaufman et al. 2005; Geden et al. 2006; Taylor and Berkebile 2006; Mihok and Carlson 2007; Mihok et al. 2007). No single control method tested thus far is effective in decreasing stable fly populations below the economic injury threshold (Meyer et al.1990; Seymour and Campbell 1993; Cilek and Greene 1994; Thomas et al. 1996; Macedo 2004; Taylor and Berkebile 2006). The current procedure recommended as the most efficient means of stable fly control is an Integrated Pest Management approach, which stresses the importance of sanitation, and utilizes a combination of the methods listed above (Watson et al. 1994; Powell and Barringer 1995; Campbell and Thomas 1999; Campbell 2006).

Sanitation is an important control factor in feedlots and dairies. The removal of organic waste such as spilled feed and manure, regular cleaning, and good drainage decreases larval development sites. Manure can be spread out to dry, or piled in mounds with sufficient drainage. During wet weather the edges of the mounds should be scraped away in order to dry (Campbell 1995, 1997, 2006; Watson et al. 1994; Campbell and Thomas 1999).

Chemical controls can be effective for short periods, but require that the treatment be repeated regularly. Animal sprays give some relief, but are washed off when the cattle walk through damp grass or stand in water for avoidance (Campbell and Hermanussen 1971; Watson et al. 1994; Campbell 1997). Residual sprays applied to fly resting surfaces such as fences, buildings and vegetation can be effective for up to 14 days, provided that extensive alternate resting places are not accessible nearby. They may also be washed off in the rain or decomposed by direct sunlight (Watson et al. 1994; Campbell 1997; Campbell and Thomas 1999). Area sprays are effective in areas where flies congregate, but are not a long term solution as they only kill the flies they contact (Watson et al. 1994; Campbell 1997; Campbell and Thomas 1999).

Feed-through fly controls, which contain larvicides or insect growth regulators, pass through the digestive system of the host animal and remain in the feces. These controls are not effective for the control of stable flies, as they do not oviposit in fresh cow manure (Campbell 1997).

The application of larvicides on fly breeding areas is not an effective control method. The acidity of the substrate decomposes the larvicides rapidly, and frequent application promotes insect resistance to the chemicals (Campbell 1997; Campbell and Thomas 1999).

Releasing parasitic wasps is not an effective means of stable fly control. The numbers of flies are not significantly reduced, and the cost of the parasitoids is more than the economic loss from the flies (Andress and Campbell 1994; Campbell 1997, 2006).

Baits are not an effective method against stable flies, since they feed on blood and are not attracted to the baits (Campbell 2006). Traps, however, have been effective at capturing stable flies. They are attracted to certain olfactory stimuli such as CO2, ammonia, and phenylpropanoid compounds (Gatehouse 1967; Hammack and Hesler 1996), and visual stimuli such as Alsynite fiberglass which reflects UV light (Gersabeck et al. 1982; Black and Krafsur 1985; Allan et al. 1987). Stable flies respond to wavelengths of light in the UV range (360 nm) and the blue range (450-550 nm) (Allan et al. 1987).

A sterile insect release program combined with IPM practices was conducted in St. Croix, US Virgin Islands with some success (LaBrecque et al. 1981; Patterson et al. 1981; Williams et al. 1981; Willis et al. 1981, 1983; Bartlett and Staten 1996). However, the success of the sterile insect technique would be unlikely with stable flies on a large scale. Although 4 out of 5 feasibility factors for the method (Knipling 1955) apply to stable flies, the populations may be too large, and immigration of wild flies into an area would be a problem due to their long distance dispersal capability (Bailey et al. 1973; Hogsette and Ruff 1985).

Stable Flies and Disease

In addition to being a pest of livestock and other animals, stable flies are known to be mechanical vectors of many diseases. In India, they are abundant pests of animals such as sambar, deer, mithan, blackbuck, and various carnivores, and are mechanical vectors of surra disease and equine infectious anemia (EIA) virus (Veer et al. 2002). In a study in the Czech Republic and Slovakia, two species of Mycobacterium were isolated from adult stable flies at a farm raising both cattle and pigs (Fischer et al. 2001). A laboratory study by the US Army Medical Research Institute of Infectious Diseases showed that stable flies can mechanically transmit Bacillus anthracis, the agent of anthrax, and Rift Valley fever virus (Turell and Knudson 1987). Bartonella henselae type M was isolated from stable fly DNA during a study in California (Chung et al. 2004).

Enteric bacteria are transmitted by stable flies, as could be expected from their association with animal feces. Campylobacter spp. were detected in stable flies collected from turkey production facilities in Arkansas (Szalanski et al. 2004). In a laboratory experiment in which stable flies were orally inoculated with Enterobacter sakazakii, over 50% of the flies still carried the pathogen 20 days after inoculation. E. sakazakii also had significant positive effects on stable fly development (Mramba et al. 2007). Escherichia coli have a positive effect on stable fly larval development when in a mixed bacterial community. The larvae ingest the E. coli but do not digest it readily, so cattle feeding on silage containing the infected larvae may ingest the bacteria (Rochon et al. 2004). Puparia of infected larvae have also been found to contain large amounts of E. coli (Rochon et al. 2005).

Some bacteria are pathogenic to stable flies, such as Aeromonas sp., Pseudomonas aeruginosa and Serratia marcescens (Lysyk et al. 2002). Further research on the efficacy of such pathogens in causing mortality in stable flies could be another potential control method.

Stable Fly Genetics

The majority of the genetic research on stable flies to date has been focused on genetic variation among or between populations, in an attempt to determine their origin and dispersal patterns, and genetic control strategies such as sterile male release programs.

Genetic control methods

Following the success of the screwworm (Cochliomyia hominivorax) sterile male release program on the island of Curacao, Knipling (1955) suggested several applications for this method, including the control of small numbers of naturally occurring pests, newly established populations of pests, or in conjunction with other integrated pest management practices (Knipling 1955). In 1974, a sterile male release program was initiated on St. Croix, US Virgin Islands, as a component of an integrated pest management program to control stable fly populations. At that time, populations averaged 9.5 x 105 during the wet season and 2.5 x 105 during the dry season (La Brecque et al. 1981). The program included mass rearing of 250,000-300,000 flies per day, with 70,000 required for colony maintenance. Males from 24-48 h old were sterilized by exposure to 2 kR of cobalt-60 gamma radiation (Williams et al. 1981). For 18 months during 1976-1977, sterile males were released at a rate of 1 x 105 per day, 5 days per week. By the end of the project, the stable fly population was reduced to ~350 flies, although not entirely eliminated. However, after cessation of the project, during just 3 generations, populations rose to 210,000 (Patterson et al. 1981).

During the 1980's, genetic mutations were investigated as potential control mechanisms. The stable fly has 5 pairs of chromosomes, with 4 recessive mutations being reported at the time. (Hunter et al. 1992) Chromosome 1 contains the sex locus, chromosomes 2, 3, and 4 contain the carmine eyes (ca) mutant, the black pupa (bp) mutant, and the rolled down wing (rd) mutant, respectively (Willis et al. 1981; Willis et al. 1983; Hunter et al. 1992). The fourth recessive mutant, subcostal incomplete (sci) was reported in 1992 (Hunter et al. 1992). Possibilities for stable fly control using mutations consisted of DNA recombination techniques such as reciprocal translocations (Willis et al. 1981), and genetic sexing techniques to eliminate females (Willis et al. 1983; Seawright et al. 1986; Bartlett and Staten 1996). The latter was accomplished using chemical susceptibility genes or the black pupa mutant gene (Willis et al. 1983; Seawright et al. 1986; Bartlett and Staten 1996). These methods could be effective when combined with an integrated pest management program (Bartlett and Staten 1996).

Population genetics

Due to the ubiquity and pestiferous nature of stable flies, it would be an advantage to determine the origin of seasonal populations and their dispersal patterns. It has been shown that immature stable flies are able to overwinter in livestock areas, in build-ups of substrate that retain some heat, such as piles of wasted hay, silage, grass clippings and compost piles (Berkebile et al. 1994; Broce et al. 2005). However, the source of the populations remains unknown (Broce et al. 2005). Stable flies are able to fly long distances and to disperse with the wind (Bailey et al. 1973; Gersabeck and Merritt 1985; Hogsette and Ruff 1985; Beresford and Sutcliffe 2009). In a flight-mill test, a stable fly was reported to fly 29.11 km in 24 hours (Bailey et al. 1973). A study in Northwest Florida, in which stable flies were marked with fluorescent dust, released, then recaptured in Williams traps, reported a flight range of 225 km.In this area, stable fly populations become so dense on the beaches that tourism comes to a standstill. It is believed that the flies migrate to the beach areas, since there are few breeding sites available (Hogsette and Ruff 1985).

Several studies of stable fly population genetics have attempted to determine dispersal patterns and sources of the populations. Jones et al. (1991) tested allele frequencies of 10 different enzymes using protein electrophoretic analysis. They collected 100 stable flies from each of 37 sample sites from 1982-1985, including 8 beach sites in Florida, 7 dairies in Florida, dairies in Indiana, Maryland, New York and Texas, and a feedlot in Nebraska. Their data showed very little variation among populations, suggesting a high level of gene flow across the United States. They suggested that the movement of stable flies is due to drifting on weather fronts rather than migratory behavior, and that flies on the Florida beaches could have originated as far away as Nebraska (Jones et al. 1987; Jones et al. 1991).

Szalanski et al. (1996) performed a study using the polymerase chain reaction-restriction fragment length polymorphism (PCR-RFLP) technique. They screened portions of the cytochrome oxidase (CO) I, II, and III mitochondrial DNA genes, NADH 4 and 5 genes, and nuclear ribosomal DNA genes. Their samples were primarily from Nebraska, with samples from Texas and Manitoba included. The results were similar to Jones et al. (1991), and they also reported very low levels of genetic differentiation among populations (Szalanski et al. 1996).

Gilles et al. 2007 also reported low levels of genetic differentiation in the stable flies on dairy farms on La Reunion Island, a small (2507 km2) island in the Indian Ocean east of Madagascar. They noted more differentiation at two sites which used dissimilar farming methods. The methods used in this study were the analysis of 7 microsatellite loci which had been sequenced previously (Gilles et al. 2004). The 7 loci were amplified by PCR and then sequenced (Gilles et al. 2007).

Contradictory data were reported by Marquez et al. (2007). They examined r16S and COI mitochondrial DNA loci from 11 different countries including the United States,by amplifying the DNA and sequencing the PCR products. This group reported considerable variation in the mtDNA of stable flies (Marquez et al. 2007).

Other genetic research

The hematophagous nature of stable flies generates interest in the innate immune responses developed by this insect to resist the pathogenic microbes it is exposed to during feeding. Of particular interest are the antimicrobial peptides (AMPs), most of which are produced by the fat body. Three AMPs specific to the stable fly have been sequenced, which are unique in that they are specific to the anterior midgut. Two are defensins: stomoxys midgut defensins (smd) 1 and 2 described by Lehane et al. (1997), which exhibit anti-Gram negative activity. The third, stomoxyn, was identified by Boulanger et al. (2002). It is a cecropin-like peptide which exhibits a wide spectrum of anti-microbial activity against bacteria, fungi, and trypanosomes (Boulanger et al. 2002).

Genetic Analysis Techniques

A variety of techniques are utilized in genetic studies, depending on the nature of the research, including the analysis of proteins, nuclear DNA and mitochondrial DNA (mtDNA). DNA technology advances rapidly, and methods are continually modified for optimum performance (Jones et al. 1987; Jones et al. 1991; Gilles et al. 2004). Interest in the mitochondrial genome dominates current research, with the sequencing of the cytochrome oxidase subunits I and II of the stable fly and other insects (Szalanski and Owens 2003; De Oliveira et al. 2005; Marquez et al. 2007).

Protein Electrophoresis

Water-soluble proteins are extracted from the sample and absorbed onto a paper wick. The wick is placed in the well of a starch or acrylamide gel, and the gel is placed into a buffer. Electricity is applied to the gel buffer, and electrophoresis continues for several hours. The gel is then sliced horizontally and the thin slice is incubated with a stain that contains a substrate that is specific to the target enzyme. The gel is visualized in a light box and the bands are compared to known samples (Jones et al. 1987; Avise 2004). This method is simple and not too time consuming. Many different codominant alleles can be identified at numerous loci, and the data can be easily replicated, although there is only moderate resolution of genetic differences. This method is useful for studying population genetics and relationships between species (Avise 2004).


DNA is extracted from the sample, and restriction enzymes are added to the DNA. Restriction enzymes cleave the DNA at specific sites which are usually 4, 5, or 6 nucleotides in length. The recognition for EcoRI, for example, is the base sequence 5'-GAATTC-3' (Avise 2004). The cleaved DNA is then electrophoresed on agarose or acrylamide gel to separate the different sized fragments. The gel is transferred to a nylon membrane, where radioactive probes are added. The probes bind to the DNA fragments, the gel is dried and an X-ray film is placed over it. When developed, the results are an autoradiograph on which the DNA fragments can be visualized. This method is called Southern hybridization, or Southern blot (Hoy 2003; Avise 2004).

The development of the polymerase chain reaction (PCR) in the 1990's facilitated the RFLP technique, allowing for extraction of DNA from much smaller samples. PCR is an automated method of exponentially amplifying DNA, which involves 20-30 cycles of 3 different temperatures: 94ºC for 20 seconds for denaturation of the template DNA, 55ºC for 20 seconds for annealing primers to the template, 72ºC for 30 seconds for extension of the DNA. The PCR cycles must be optimized for each organism. Reactions require a PCR mix containing dNTPs, Taq DNA polymerase, PCR buffer, MgCl2, specific forward and reverse primers and template DNA. During the denaturation step, the double strands of the DNA separate. Specific primers (added to the PCR mix) anneal to the ends of the target sequence on the DNA, and the DNA between the primers is replicated. Each cycle doubles the amount of DNA, allowing for millions of copies to be replicated in a very short time (Hoy 2003; Avise 2004; Varsha 2006). The Southern blot has been replaced by the addition of dyes such as ethidium bromide to the gel, which can then be visualized under UV light (Clark 2000).

PCR-RFLP is used for both nuclear and mitochondrial DNA analysis (Szalanski et al. 1996).

It requires more time than other techniques, but the data can be replicated without difficulty. The bands indicate codominant alleles using nDNA and maternal alleles using mtDNA. Usually few loci are assayed, but many alleles per loci can be identified (Avise 2004).


Randomly amplified polymorphic DNA (RAPD) is a technique in which universal primers are used to amplify unknown DNA sequences. Short primers are used which have the ability to generate multiple fragments. Polymorphisms are detected when the PCR product is separated by gel electrophoresis. Using the RAPD technique, one is able to detect small differences in populations, species, and individuals because it generates numerous DNA fragments and many loci can be analyzed in one reaction. It requires only a small amount of DNA and is relatively inexpensive (Hoy 2003; Christen 2008). This method is a popular method in population biology, but is not very reproducible and only reveals dominant markers.


Amplified fragment-length polymorphism is a technique that combines the RFLP and RAPD methods (Bensch and Akesson 2005). Genomic DNA is digested by two restriction enzymes, usually EcoRI and MseI. Adapters (short DNA fragments complimentary to the loci cut by the enzymes) are ligated to the ends of each fragment. Then a preamplification is run with primers that are complimentary to the adapter and the enzyme, plus an additional nucleotide. Following preamplification, a small amount of the product is added to a PCR mix containing specific primers, amplified and separated on polyacrylamide gel (Avise 2004; Bensch and Akesson 2005).

AFLP is a relatively inexpensive method of screening a large number of loci, and has become a useful tool in the field of population genetics (Campbell et al. 2003; Bensch and Akesson 2005). Although it reveals only dominant markers, and is based on the presence or absence of a band (Bonin et al. 2007), it has been shown to be as efficient as techniques that separate codominant markers, such as microsatellites (Bensch and Akesson 2005).

DNA sequencing

Historically, two methods were developed for DNA sequencing, one by Maxam and Gilbert, the other by Sanger. Each involved isolating and denaturing the DNA, labeling the ends with radioactive primers, and separating the fragments by gel electrophoresis. Each method required four reactions, one for each deoxynucleotide (Avise 2004). Currently, sequencing is predominantly automated, using PCR thermocyclers connected to sequencing machines (Avise 2004).

Other techniques

Other methods of DNA analysis could be useful in analyzing stable fly population genetics. Short interspersed elements (SINEs), single-strand conformational polymorphism (SSCPs), single nucleotide polymorphism (SNPs), techniques described in Avise (2004), are methods that could be employed in population genetics. Essentially, experimental designs should consist of available techniques that provide the maximum quantity and quality of data with the least expenditure of time and money.


Effective control strategies for stable fly populations are of primary importance in the livestock industry, as well as other affected areas such as tourism on Florida beaches and the convergence of urban and agricultural habitats. Because stable flies are distributed world-wide, and have the ability to travel long distances, single control methods have unsatisfactory results. The application of Integrated Pest Management strategies appears to be the most successful approach so far. Improved sanitation, such as removal of manure and wasted feed, washing of stalls and milking areas, and good drainage systems are important practices to eliminate breeding areas and larval development sites. Insecticides, traps and biological controls aid in reducing fly numbers. However, these practices do not reduce stable fly numbers to an acceptable level.

An enormous amount of possibilities exist in the study of stable flies and their control, and the logical direction is to implement control strategies at their source. On a local scale, more efficient methods could be developed to eliminate larval development and overwintering sites. New biological control methods could be investigated, such as recombinant DNA technologies that block the production of essential hormones or antimicrobial peptides. Further study on dispersal patterns using release and recapture techniques could aid in locating local sources of stable fly outbreaks.

Further research is needed in stable fly population dynamics, to investigate dispersal patterns and possible sources of stable fly populations. To date, studies have indicated low differentiation and high gene flow among populations, even on a small island scale (Szalanski et al. 1996; Gilles et al. 2007), although Marquez et al. (2007) reported a high level of variation in stable fly mitochondrial genomes on a global scale. Each group used different DNA analysis techniques and different sample areas. Szalanski et al. (1996) examined populations in Nebraska by PCR-RFLP; Gilles et al. (2007) concentrated on La Reunion Island using microsatellites which were amplified and sequenced; and Marquez et al. (2007) chose a wider experimental area including 11 different countries, with direct sequencing of mtDNA. Different techniques could generate more informative data on stable fly population dynamics. Larger sample sizes could prove beneficial, and many more loci could be evaluated using AFLP, including DNA sequences that are as yet undescribed. Examining populations on a global scale, such as the research by Marquez et al. (2007), would be more likely to show differentiation between populations, so it would be logical to begin at a large scale and work toward a smaller scale.

Technology in the fields of genetics and molecular biology advances rapidly, and concomitantly, the capability for further understanding of stable fly biology and habits increases. Since the purpose of stable fly research is population control, advancement is a step toward success. The more knowledge we acquire about stable fly population genetics, the further we are toward understanding the methods to control their numbers. While investigating control methods, we may also discover beneficial ecological niches for the stable fly that compel us to more readily accept their presence.

Research Objectives

More research is needed in the population genetics of stable flies to increase our knowledge of their dispersal patterns and the sources of outbreaks. Advancements in DNA technology over the past decade offer more efficient methods of screening a larger number of genetic loci with an equivalent input of time and expenses. Describing variations in population distribution, as well as finding genetic markers that define the variations, will expand our understanding of the population dynamics of the stable fly. The goal of this research is to investigate genetic variation in stable fly populations across the United States and on a global scale.

I propose that more genetic variation will be revealed using the Amplified Fragment Length Polymorphism (AFLP) method of genetic analysis than in previous studies using other methods of DNA analysis. AFLP may reveal genetic markers for stable flies that have not been previously described and entered into a database such as GenBank. There will be genetic variation in populations separated by geographical barriers, both globally and within the United States.

Objective 1:

Analyze genetic variations in stable fly populations from several geographic areas, including numerous locations in the United States, and other locations where samples may be obtained, such as Mexico, Central and South America, Africa, Europe, and Australia.

Null hypothesis: No genetic variation will be found, which will support the results of Jones et al. (1991), Szalanski et al. (1996), and Gilles et al. (2007).

Hypothesis: Genetic variation will be evident using the AFLP method because a greater number of loci will be screened than in previous experiments. Genetic variation will be evident across geographical barriers. This hypothesis would support the results of Marquez et al. (2007).

Objective 2:

Describe unreported genetic markers specific to Stomoxys calcitrans.

Null hypothesis: No specific genetic markers will be located.

Hypothesis: Using the AFLP method, I will find genetic markers that are specific to Stomoxys calcitrans.

Objective 3:

Analyze genetic variations of stable fly populations across the United States, in north-south, east-west, northwest-southeast, and southwest-northeast transects.

Null hypothesis: There will be no genetic variation between populations of stable flies in the United States.

Hypothesis: Genetic variation will be found in populations across the United States when divided by geographical barriers such as large mountain ranges.

Experimental Design

A sample of 50 stable flies will be collected from each experimental area to assure that I will have a sample size of at least 30 flies for genetic analysis. The samples will be obtained by the cooperative participation of colleagues in each area except for the local areas in Nebraska which I will collect. Adult flies will be soaked in 95% ethanol. The ethanol will be poured off for shipping, and will be reapplied when the samples are received. The samples in 95% ethanol will be stored at 4°C until processed. For each sample set, DNA will be extracted from the thorax using the CTAB protocol, modified from Saghai-Maroof et al. (1984), and analyzed following the AFLP protocol in our lab. Climatic and topographical data for each collection area will be gathered from the appropriate sources and considered as factors in the experiments. Statistical methods for analyzing genetic variation will include the Analysis of Molecular Variance (AMOVA). Outlier species will be used for controls, such as Musca domestica (Diptera: Muscidae), Calliphora vicina, or Phormia regina (Diptera: Calliphoridae). Samples of the Calliphorids are currently available in our lab. As a non-dipteran outlier, I will use Diabrotica vergifera vergifera or Diabrotica barberi (Coleoptera: Chrysomelidae), which are available in our lab.

Materials and Methods

For each of sample set:

30 stable fly adults will be analyzed from each site.

DNA extraction will follow the CTAB (Cetyltrimethyl ammonium bromide) protocol.

The method of DNA analysis will be AFLP, following our lab protocol.

AFLP will be performed on the LI-COR machine in our lab.

AFLP polyacrylamide gels will be scored using the SAGA software (version 3.3, LI-COR), and will be converted into a numerical matrix using 1 for presence or 0 for absence of bands (Maliphan 2006). The matrix will then be analyzed using the Analysis of Molecular Variance (AMOVA).

Outlying groups will be used for controls.

CTAB Protocol

Samples stored in 95% ethanol at 4°C will be washed twice in nanopure water. Then samples are transferred to a 1.5mL centrifuge tube containing 250µl CTAB extraction buffer. The samples are homogenized by hand using a pestle, and another 250µl CTAB buffer is added to the tube. Proteinase K (10µl per tube) is added, tubes are vortexed slowly to mix, then placed on a heat block at 65°C for 1 hour, with mixing every 20 minutes by inverting the tubes. After one hour, RNase is added to the tubes (15µl per tube) and they are placed on a heat block at 37°C for 2 hours, with mixing every 20 minutes. After adding RNase, the tubes are not vortexed.

Tubes are centrifuged for 5 minutes at 14,000rpm and 25°C to separate the solids from the liquids. Supernatant is removed and transferred to a new 1.5mL tube and the solids are discarded. 500µl chloroform: isoamyl alcohol (24:1) is added to each tube, and they are mixed by inverting several times. Tubes are centrifuged for 20 minutes at 14,000 rpm and 25°C. There will be two layers in the tube. The top aqueous phase is removed and put into a new tube. The chloroform layer is discarded in the chloroform waste container. The chloroform: isoamyl step is repeated.

The top aqueous phase is removed without disturbing the interface between layers, and transferred to a new tube. The chloroform layer is discarded. Chilled isopropanol (-20°C) is added (400µl) to each tube. Tubes are gently mixed, then placed in the 4°C refrigerator overnight.

Samples are centrifuged for 30 minutes at 14,000rpm and 4°C. A pellet should be seen at the bottom of the tubes. Liquid is poured off, and 500µl chilled (-20°C) absolute ethanol is added to each tube. Tubes are tapped to free the pellet from the bottom and centrifuged for 5 minutes at 14,000 rpm and 4°C. Supernatant is removed and 500µl chilled 70% ethanol is added to each tube. Samples are centrifuged again for 5 minutes at 14,000rpm and 4°C and ethanol is removed using a pipette, so that as much as possible is removed from the tube. Tubes are left open in the hood to air dry. After all the ethanol has evaporated, 50µl TE buffer is added to each tube, and samples are put in the 4°C refrigerator overnight.

DNA Quantification

Samples are analyzed on the Nanodrop® spectrophotometer. They are also electrophoresed on 1% agarose gel to determine quantity and quality of DNA in comparison to known standards.

AFLP Protocol

A restriction digestion mixture is made containing MseI and EcoRI restriction enzymes. 5.5µl of the mixture is added to the tubes, and 7.0µl of template DNA is added to the restriction digestion mixture. The samples are digested and enzymes denatured using the following program on a therm cycler: 60 minutes at 37°C, 90 minutes at 37°C, 15 minutes at 70°C.

Adapters, corresponding to the restriction enzymes and DNA ligase are added to the tubes, making a volume of 15µl. Samples are diluted by adding 135µl TE buffer to the digestion/ligase mixture in a .5mL PCR tube, and left overnight at 4°C.

A preamplification mixture is made containing pre amp primer mix, PCR buffer, and DNA polymerase. In sterile PCR tubes, 11.5µl of the mix is added to 1.25µl of the diluted template DNA. Tubes are run on the thermal cycler on the following program: Twenty cycles of 30 seconds at 94°C, 1 minute at 56°C, 1 minute at 72°C. Samples are left in thermal cycler at 4°C overnight.

PCR product is diluted 1:20 by adding 190 µl nanopure water to 10µl of the pre amp PCR product. PCR mixes are prepared for each selective primer pair to be used. 8.5µl selective primer mix is put into tubes and 2.0µl of the DNA is added. Samples are run on the “selective amplification” thermal cycler program which consists of: 1 cycle of 30 seconds at 94°C, 30 seconds at 65°C and 1 minute at 72°C; 12 cycles of 30 seconds at 94°C, 30 seconds at 65°-56°C, 1 minute at 72°C; 23 cycles of 30 seconds at 94°C, 30 seconds at 56°C, 1 minute at 72°C. Samples can be left on the thermal cycler overnight at 4°C.

Polyacrylamide Gel Preparation and Electrophoresis

Plates are washed with cleaning solution, rinsed thoroughly, rubbed with pledge furniture polish, then covered with alcohol and left to air dry. After plates are dry, they are clamped together and the binding solution and gel are prepared. Binding solution is applied to the plates in the area where the comb will be placed. Gel matrix is mixed with ammonium persulfite solution and the gel is applied between the plates using a pipette. A comb is placed at the top of the gel. The gel is allowed to set for 2 hours. After gel is set, it is loaded into the LI-COR machine and buffer is added.

Stop solution is added to each sample (2.5µl), and they are denatured on the thermal cycler for 3 minutes at 94°C and cooled on ice for 5 minutes. Samples are loaded in the gel. The gel is electrophoresed for 2 hours and recorded on the computer.


Collections will be made throughout the year as seasons differ in collection locations. Collections in the Midwest will be made during the summer of 2009 and 2010. All collections should arrive by the spring of the next year. Analysis will be performed after each collection, and will be ongoing through the winter of 2009 and 2010. Since this is a laboratory experiment, there should be no time restraints to obtaining results, except for receipt of the samples. Geographical data will be obtained before the experiment begins, and climatic and weather data will be obtained after each collection. Data will be analyzed with the Analysis of Molecular Variance (AMOVA), using Arlequin software (Schneider et al. 2000).


If the results support the null hypothesis, that no genetic variation exists between stable fly populations, it will support the aforementioned studies that assume a high level of gene flow between populations. However, if my hypotheses are supported, and there is a level of genetic variation between populations, as well as genetic markers that are specific to Stomoxys calcitrans, possibilities for further research into genetic variations will be revealed. Patterns in the phylogenetic relationships between populations may occur, which may lead to locating the sources of stable fly populations and subsequently the development of more efficient methods of control.