Introduction

The house fly, Musca domestica L. (Diptera: Muscidae), is a serious pest of livestock and humans (Scott et al 2000, Kaufman et al 2001, Deacutis et al 2006), causing health and environmental problems by vectoring diseases from animals to man and animal to animal. House flies reproduce on all eatable materials and wastes and are a known vector of more than 100 disease pathogens, including bacteria, protozoa, helminths, and viruses (Förster et al 2007, Acevedo et al 2009, Khan et al 2012, Abbas et al 2014a). Outbreaks of diarrheal diseases are associated with seasonal abundance of house flies in urban and rural settings of many countries, including Pakistan (Graczyk et al 2001, Khan & Akram 2014). The house fly is also a vector of the bird flu virus, an avian influenza that is a serious problem in poultry throughout the world (Barin et al 2010, Wanaratana et al 2011). Millions of domestic poultry flocks are affected by the avian influenza virus which has caused more than 150 deaths of humans globally (Otte et al 2007). In areas of poultry farming, like Punjab Pakistan, uncovered poultry manure provides an ideal condition for growth and reproduction of house flies. High-density populations of flies irritate workers, stress hens, and reduce the aesthetic value of livestock, leading to economic losses (Acevedo et al 2009, Abbas et al 2014a).

Synthetic insecticides, including organophosphate, carbamate, pyrethroid, and novel chemistry classes, have been used to control the house fly worldwide, including Pakistan. Judicious and appropriate insecticide applications are helpful in conditions where high densities of the house fly are associated with avian influenza in poultry (Nielsen et al 2011, Wanaratana et al 2011) and diarrhea in humans (Chavasse et al 1999, Khan & Akram 2014), but season-specific use of insecticides is required. However, inappropriate chemical use and poultry manure stored in open places could result in the development of resistance to insecticides by house flies. These factors may be responsible for avian influenza and diarrhea cases in poultry and humans, respectively, particularly in developing countries. This resistance development is also promoted by a number of biological traits such as adaptability to different environments, high fecundity, a short developmental time, and cross-resistance (Kaufman et al 2010a, Khan et al 2013b). Resistance to all chemical classes is common in house flies worldwide (Scott et al 2000, Acevedo et al 2009, Kaufman et al 2010b, Khan et al 2013a,b, 2014a, Abbas et al 2014a,b).

The determination of the stability of insecticide resistance has practical implications in developing effective resistance management strategies (Khan et al 2014b, Afzal et al 2015). The removal of insecticides from spray schedules may bring the resistance level down if resistance is unstable, and consequently, the efficacy of an insecticide can be extended. A decrease in resistance has been reported for insecticide-selected insect populations in the field and laboratory (Carriére & Tabashnik 2001). Effective resistance management programs depend upon different mechanisms of resistance and factors that enhance susceptibility recovery. A reduction in resistance levels may be due to the negative effects of resistance genes on fitness components without insecticide selection pressure. When insecticide exposure is not a constant factor, the susceptibility of resistant individuals may change in the absence of selection pressure under laboratory conditions (Roush 1993, Kristensen et al 2000, Abbas et al 2014b).

We assessed the fluctuations in insecticide resistance of house fly populations from Khanewal, Punjab, Pakistan. House fly peak densities start in March and continue until August. Therefore, house fly populations were collected in different seasons (late summer, early winter, and early spring) to assess the variation in susceptibility to pyrethroids (cypermethrin, lambda-cyhalothrin), organophosphate (chlorpyrifos), neonicotinoids (imidacloprid, acetamiprid, nitenpyram), spinosyn (spinosad), oxadiazine (indoxacarb), and avermectins (abamectin and emamectin benzoate). The results of this study can have a significant role in developing strategies for house fly resistance management through the selection of the most appropriate insecticide to deal with the unstable nature of resistance in specific months of the year for the management of house flies.

Material and Methods

Insect rearing

About 200 adult house flies were collected from a poultry farm located at Khanewal (32°4′50N, 73°38′10E) in three different seasons denoted by March, July, and November with a sweep net. Adult flies were kept in meshed plastic jars (34 × 17 cm) and fed on a mixture of sugar and powdered milk (1:1 w/w). Cotton wicks soaked with water were provided in a Petri dish in order to supplement moisture requirements. Larvae were reared on an artificial diet comprising powdered milk, sugar, yeast, grass meal, and wheat bran (1.5:1.5:5:5:20 w/w) and 65 mL of water (Abbas et al 2014a). The colony was maintained under laboratory conditions according to Abbas et al (2014a). The susceptible strain was collected from an insecticide-free area (Multan) and maintained in the laboratory without exposure to insecticides.

Insecticides

Commercial formulated insecticides were used for bioassays including conventional: chlorpyrifos (Lorsban® 40EC, Arysta Life Sciences), cypermethrin (Arrivo® 10EC, FMC), and lambda-cyhalothrin (Karate® 2.5EC, Syngenta); and newer chemistries: spinosad (Tracer® 240SC, Arysta Life Sciences), indoxacarb (Steward® 150SC, Arysta Life Sciences), abamectin (Alarm® 1.8EC, DJC), emamectin benzoate (Proclaim® 019EC, Syngenta), acetamiprid (Mospilan® 70WP, Arysta Life Sciences), imidacloprid (Confidor® 20SL, Bayer Crop Science), and nitenpyram (Paranol® 10EC, Kenzo Agro Chemicals).

Toxicological bioassays

The toxicities of the tested insecticides were assessed by using a feeding bioassay (Kaufman et al 2006). Ten 2–3-day-old male and female flies were provided with a cotton wick (3 cm) soaked in 20% sugar solution plus insecticide and kept in plastic jars (14 × 9 cm) (Abbas et al 2014b). Five concentrations were prepared as serial dilutions for each insecticide, and each concentration was replicated three times (10 flies/replicate). The range of concentrations was 8–128 μg·mL−1 for organophosphate, pyrethroid, and neonicotinoid; 0.125–2 μg·mL−1 for avermectins; and 1–16 μg·mL−1 for spinosyn and oxadiazine in all tested strains. Cotton wicks soaked in 20% sugar solution without insecticide were provided to control flies (3 replicates of 10 flies). Cotton wicks were hydrated (without sugar solution) after 24 and 48 h to avoid wick drying (Kaufman et al 2006). All flies were kept under laboratory conditions as earlier described. Mortality data was assessed 48 h after exposure to conventional and 72 h after exposure to newer chemistry insecticides. Ataxic flies were considered as dead (Abbas et al 2014a).

Data analysis

Concentration response data were analyzed by probit analysis (Finney 1971) with POLO software (Software 2005) to determine LC50 values, their 95% confidence intervals (CI), slopes, and their standard errors. Control mortality was corrected by the formula of Abbott (1925), whenever required. Resistance ratio (RR) and its 95% CI were calculated by dividing the LC50 value and its 95% CI of test strains by the LC50 of the susceptible strain (Ban et al 2012). LC50 values were not considered significantly different when their 95% CI overlapped. The resistance ratio was considered significant if the 95% CI did not include the value of 1 (Robertson & Preisler 1992).

Variation in resistance per month (R) was estimated with the following formula:

$$ R= \log \left({\mathrm{final}\ \mathrm{L}\mathrm{C}}_{50}\right)- \log\;\left({\mathrm{initial}\ \mathrm{L}\mathrm{C}}_{50}\right)/n $$

In the above equation, n indicates the months between two collections. Positive and negative values for R indicate the increase and decrease in resistance, respectively.

Coefficient of correlation of mortality responses to insecticides were used to calculate the coefficient of determination by taking square of the coefficient of correlation of responses between any of two tested insecticides (Sayyed et al 2005). LC50 values were not considered significantly different when their 95% CI overlapped (Robertson & Preisler 1992).

Results

Toxicity of insecticides to a house fly-susceptible strain

The susceptible strain of the house fly was more susceptible to the newer chemistry insecticides. Abamectin, indoxacarb, emamectin benzoate, and spinosad were more toxic compared with other tested insecticides. Chlorpyrifos, lambda-cyhalothrin, cypermethrin, and nitenpyram were equally toxic. Imidacloprid and acetamiprid were less toxic (Table 1). The LC50 values of spinosad, indoxacarb, abamectin, and emamectin benzoate were significantly lower compared with other insecticides tested except chlorpyrifos (95% CI did not overlap). There was no significant difference in LC50 values of cypermethrin, lambda-cyhalothrin, acetamiprid, and nitenpyram (95% CI overlapped). The LC50 of imidacloprid was higher compared with all other tested insecticides (95% CI did not overlap) (Table 1).

Table 1 Toxicity of insecticides to the susceptible strain of Musca domestica.
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Stability of resistance to conventional insecticides

Resistance to conventional insecticides in poultry facility-collected strains varied significantly with the season (Table 2). In populations collected at 3-month intervals, the resistance level for chlorpyrifos decreased significantly in March compared with July and November but increased slightly in November compared with the strain collected in July. The resistance to cypermethrin and lambda-cyhalothrin decreased significantly in March compared with July and November. The declined rate of resistance to cypermethrin (−0.2) and lambda-cyhalothrin (−0.3) was similar for the strain collected in March.

Table 2 Seasonal variation in Musca domestica resistance to different insecticides.
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Stability of resistance to newer chemistry insecticides

Resistance to newer chemistry insecticides varied significantly with the season in the poultry facility-collected house fly strains (Table 2). The resistance to spinosad remained stable throughout the season in which strains were collected. The resistance to indoxacarb, abamectin, and emamectin benzoate decreased significantly in November compared with the strains collected in July and March. However, resistance remained stable for the strains collected in July and March. The declined rate of resistance against indoxacarb (−0.2), abamectin (−0.2), and emamectin benzoate (−0.3) was similar for the strain collected in November. The resistance to imidacloprid remained stable throughout the season in which strains were collected. However, the resistance to acetamiprid and nitenpyram increased significantly in November compared with the strain collected in July and March. The resistance to acetamiprid and nitenpyram did not decrease significantly in July compared with March.

Correlation in insecticides

All conventional and newer chemistry insecticides depicted significant correlations with each other in all tested seasons except nitenpyram/lambda-cyhalothrin and nitenpyram/imidacloprid, which showed a non-significant correlation in the population collected in July. Similarly, acetamiprid/cypermethrin, acetamiprid/lambda-cyhalothrin, and nitenpyram/acetamiprid showed non-significant correlation in the population collected in November (Table 3).

Table 3 Correlations in mortality to insecticides in different seasons for different populations of Musca domestica.
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Coefficient of determination

The coefficient of determination for cypermethrin/chlorpyrifos decreased between July and November while it increased between November and March. The coefficient of determination for lambda-cyhalothrin/chlorpyrifos and lambda-cyhalothrin/cypermethrin increased between July and November while it decreased between November and March (Fig 1a). The coefficient of determination of acetamiprid/chlorpyrifos or cypermethrin or lambda-cyhalothrin decreased between July and November but increased between November and March (Fig 1b).

Fig 1
figure 1

Variations in coefficients of determination of different combinations of a cypermethrin, lambda-cyhalothrin, and chlorpyrifos; b of acetamiprid, chlorpyrifos, cypermethrin, and lambda-cyhalothrin; c of imidacloprid, chlorpyrifos, cypermethrin, and lambda-cyhalothrin; and d of nitenpyram, chlorpyrifos, cypermethrin, lambda-cyhalothrin, acetamiprid, and imidacloprid from July to March for different populations of Musca domestica.

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The coefficient of determination for imidacloprid/acetamiprid decreased between July and November and increased between November and March. The coefficient of determination increased for imidacloprid/cypermethrin while it decreased for imidacloprid/chlorpyrifos from July to March. The coefficient of determination for imidacloprid/lambda-cyhalothrin remained stable between July and November but decreased between November and March (Fig 1c).

The coefficient of determination for nitenpyram/chlorpyrifos increased from July to March. The coefficient of determination for nitenpyram/cypermethrin decreased between July and November while it increased between November and March. In contrast, the coefficient of determination for nitenpyram/lambda-cyhalothrin increased between July and November and decreased between November and March. The coefficient of determination for nitenpyram/acetamiprid decreased between July and November but increased between November and March. The coefficient of determination for nitenpyram/imidacloprid increased between July and November and remained stable between November and March (Fig 1d).

Discussion

Organic insecticides are the major alternative available for house fly control at livestock farms. However, the development of house fly resistance to insecticides slows down the effectiveness of new preparations, making quite difficult the selection of the most effective insecticides (Khan et al 2013a). The use of insecticides with long residual action in close vicinities of productive areas exerts a strong selection pressure that increases resistance development (Scott et al 2000). Conventional (pyrethroids and organophosphates) and newer chemistries (spinosyn, oxadiazine, and avermectin) are commonly used for the management of house fly in livestock farms worldwide, including Pakistan. The fluctuations in insecticide resistance patterns observed in our study suggest that the susceptibility decreased during July and November for chlorpyrifos, cypermethrin, lambda-cyhalothrin, spinosad, and acetamiprid. Susceptibility decreased in the months of July and March for indoxacarb, abamectin, and emamectin benzoate, but increased for nitenpyram. Resistance to imidacloprid remained stable throughout the months in which strains were collected. For some insecticides, resistance increased in late summer to early winter and decreased in early spring. In contrast, resistance increased in early spring and late summer and decreased in early winter for some insecticides. This might be due to resistance genes, unmeasured fitness costs, hybridization of susceptible individuals, and the selection pressure of different insecticides in poultry farms. The low level of resistance to some insecticides during a certain period of the year is due to the cessation of insecticide use in the off-season.

In this study, resistance to conventional and newer chemistry insecticides significantly varied from season to season in M. domestica. The highest use of insecticides occurs from March to August because of the high population of the house fly in these months. It is expected that an increase in resistance occurs during these months because resistant adults would have mated and oviposited. Resistance to most of the newer chemistry insecticides tested increased in July and March. However, resistance to organophosphates and pyrethroids decreased in March. Previously, the fluctuation in resistance to different insecticides has been observed in insect pests like Plutella xylostella L. (Sayyed et al 2005). The fluctuation of resistance in our study might be due to the decreased usage of organophosphate and pyrethroid insecticides in March, but increased resistance for newer chemistry insecticides is due to their frequent applications for house fly control. Higher resistance to pyrethroids and organophosphates in November may be because of cross-resistance between these insecticides. Cross-resistance between pyrethroids and organophosphates could be due to the involvement of esterases (Ahmad et al 2007). The house fly might have developed resistance through different mechanisms, but if the resistance is completely or incompletely dominant, then a minimum of two different mechanisms of resistance can occur (Rahardja & Whalon 1995, Sayyed et al 2005). Previously, an autosomal, incompletely dominant resistance and also the involvement of glutathione-S-transferase, esterase, and microsomal monooxygenase esterase mechanism of pyrethroid resistance were reported for the house fly (Abbas et al 2014b).

The variation in resistance could also be due to the involvement of fitness costs in the absence of selection pressure from insecticides. Selection pressure with isolan (1-isopropyl-3-methyl-5-pyrazolyl dimethyl carbamate) led to the development of a relatively stable resistant phenotype in this house fly strain, suggesting fitness costs are not involved. In contrast, selection pressure with m-isopropylphenyl methyl carbamate on the house fly strain produced an unstable phenotype, suggesting high fitness costs are involved (Georghiou 1964). In a previous experiment, a house fly strain collected from the Multan region showed a dose-dependent incomplete dominance of resistance to lambda-cyhalothrin that remained unstable in the absence of selection pressure, which may reveal a high fitness cost (Abbas et al 2014b). The results of our study may have important implications for house fly resistance management to pyrethroids and organophosphates.

A simple alternate insecticide use strategy can be easily practiced in tropical and sub-tropical climatic conditions. Applications of insecticides at a specific season when susceptibility is higher can lead to a greater change in frequency of resistance alleles. Therefore, alternate use of insecticides belonging to different groups should be recommended and inhibit the application of insecticides of the same group repeatedly. To prevent diarrheal and avian influenza epidemics, house fly outbreaks should be controlled by using appropriate insecticides suited to the prevailing environmental situations. Moreover, alternate use of the most effective insecticides in the most suitable season of the year will reduce selection pressure and ultimately delay the development of insecticide resistance (Abbas et al 2014a, Khan & Akram 2014). The house fly collected from Khanewal showed high susceptibility to spinosad, indoxacarb, abamectin, and emamectin benzoate, so these compounds could be used for this strategy.

Poultry farmers have major problems with insecticide use for the management of poultry pests in Pakistan. Inappropriate chemical use at incorrect doses, bad spray methods, and schedule are common. Moreover, adulterated chemical formulations and poor extension services are of significant importance. If the earlier mentioned problems are addressed in a proper manner and farmers are trained for proper use of insecticides and motivated to implement newly developed strategies and techniques, then these problems can be solved in the future. On the basis of our data, we recommend that chlorpyrifos, lambda-cyhalothrin, cypermethrin, spinosad, and acetamiprid should be used in March, while indoxacarb, abamectin, and emamectin benzoate in November and nitenpyram in July for house fly control.