Laboratory and field assays indicate that a widespread no-see-um, Culicoides furens Poey is susceptible to permethrin

doi:10.21203/rs.3.rs-4953430/v1

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Laboratory and field assays indicate that a widespread no-see-um, Culicoides furens Poey is susceptible to permethrin

https://doi.org/10.21203/rs.3.rs-4953430/v1

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The recent reemergence of Oropouche virus (OROV) highlights the need to better understand insecticide susceptibility in Culicoides (Diptera: Ceratopogonidae), which contains the vector of OROV and many other species that are biting nuisances and vectors of pathogens that affect humans, livestock, and wildlife. With adulticides as the primary method of Culicoides control, there is growing concern about insecticide resistance, compounded by the lack of tools to monitor Culicoides susceptibility. We adapted the CDC bottle bioassay and field cage trial methods, typically used to monitor insecticide susceptibility in mosquitoes and formulated adulticide efficacy, to evaluate permethrin susceptibility in the widely distributed coastal nuisance species, Culicoides furens. Permethrin caused complete mortality in C. furens in field and laboratory assays. We identified a diagnostic dose (10.75 µg) and time (30 minutes) that resulted in complete mortality in CDC bottle bioassays. Additionally, we determined that no-see-um netting is an effective mesh for field cage trials, allowing for accurate assessment of Culicoides susceptibility to ultra-low volume applications of formulated adulticides like Permanone 30–30, a widely utilized adulticide. These methodologies offer essential tools for assessing Culicoides susceptibility, which is crucial for managing populations of Culicoides and preventing the spread of OROV and other pathogens.

Biological sciences/Ecology

Health sciences/Diseases/Infectious diseases

Earth and environmental sciences/Ecology/Invasive species

Oropouche virus

vector control

IPM

IVM

The recent reemergence and rapid spread of Oropouche virus (OROV) in the Americas^1,2 highlights existing knowledge gaps in the ecology and control of Culicoides (biting midges, no-see-ums)³, including Culicoides paraensis Goeldi, the confirmed vector of OROV. Once considered endemic to the greater Amazonian Region of South America⁴, OROV is becoming increasingly prevalent in parts of South America, and recently spread to portions of the Caribbean, posing a significant public health risk outside of its endemic range^5,6. Vector control agencies in the areas of OROV expansion will benefit from understanding whether their mosquito control practices are effective against Culicoides spp. that bite humans.

The genus Culicoides (Diptera: Ceratopogonidae) includes species that are not only biting nuisances but also vectors affecting humans, wildlife, and livestock. Culicoides species transmit around 20 arboviruses and five nematode species of veterinary importance, affecting a wide range of hosts⁷. Notable arboviruses include African horse sickness virus (AHSV), bluetongue virus (BTV), and epizootic hemorrhagic disease virus (EHDV)^8,9. These viruses significantly impact livestock industries across multiple continents, including North America¹⁰, Central and South America⁸, Europe¹¹, Africa, Asia, and the Middle East¹².

Globally, vector control relies heavily on chemical adulticides, particularly pyrethrins and pyrethroids like permethrin due to their potency and relative safety for mammals^13,14. Permethrin is widely used for mosquito control in the United States¹⁵. Additionally, permethrin is the main active ingredient used for Culicoides control in Florida deer farms, where its frequent use has raised concerns about insecticide resistance¹⁶.

Monitoring insecticide susceptibility and resistance in target arthropods is a cornerstone of successful control program¹⁷. Insecticide resistance is often detected through laboratory bioassays¹⁸ such as the World Health Organization (WHO) tube assay and the Centers for Disease Control and Prevention (CDC) bottle bioassay¹⁷. These assays are extensively used to monitor resistance in mosquitoes and other vectors of public health significance ^17,19,20. However, standardized protocols for applying these methods to Culicoides are currently lacking. Field-based bioassays, such as “field cage trials”, are recommended to assess the susceptibility of insects to formulated insecticides like adulticides, particularly when resistance is detected in laboratory settings¹⁷.

Despite the medical, veterinary, and nuisance importance of many Culicoides spp., established protocols for monitoring susceptibility in no-see-ums are lacking. In North America, only a few studies have investigated the susceptibility of Culicoides to adulticides^21–23. These studies, conducted on Culicoides furens populations in Florida, tested the efficacy of ULV sprays containing organophosphates (malathion, naled) and a pyrethroid (resmethrin). Naled was found to be the most effective, achieving 90% mortality at distances up to 106 m, compared to 36 m for malathion and 25 m for resmethrin²³. However, a recent survey of Florida mosquito control programs documented higher usage of pyrethrin and the pyrethroid permethrin as adulticide active ingredients as compared to the organophosphates malathion and naled²⁴. Additionally, only one Florida mosquito control program reported using resmethrin in 2020²⁴.

Understanding Culicoides susceptibility to insecticides currently being used by vector control programs is essential for developing integrated vector control programs. However, studies are limited due to challenges in colonizing Culicoides and the lack of established testing protocols. While 151 Culicoides species are recorded in North America, only Culicoides sonorensis is currently colonized²⁵. Standard bioassays like the CDC bottle bioassay and cage field trials are commonly used for mosquitoes, but their applicability to Culicoides is uncertain. We adapted these methods to evaluate the susceptibility of wild C. furens from Indian River County, FL aiming to establish a diagnostic dose (DD) and diagnostic time (DT) for use in CDC bottle bioassays and identify a suitable mesh for cage field trials.

Insects. The challenges of Culicoides colonization have somewhat normalized the use of wild specimens²⁶. We tested wild C. furens, a nuisance species common in U.S. coastal areas, trapped at the University of Florida, Florida Medical Entomology Laboratory (FMEL) in Indian River County. Culicoides were captured using CDC miniature light traps (BioQuip products, Rancho Dominguez, CA, USA) connected to live midge collection chambers²⁷. Traps were baited with an incandescent light bulb, 2 mL of octenol (1-Octen-3-ol) placed in a 15 mL plastic tube with a piece of cotton wool, and ~ 1kg of dry ice contained in an insulated 1.89 L beverage container (Igloo Products Corp., Katy, TX).

Since reference (known susceptible control) C. furens are unavailable (no laboratory colony in existence), we used a laboratory-reared strain of Aedes aegypti (Orlando 1962) which originated from Orlando, FL (Orange County) as a susceptible control. This strain was originally obtained from colonies maintained at the USDA-ARS CMAVE. The Ae. aegypti were reared in an insectary at 26°C and 85% relative humidity. Egg papers were placed in plastic trays containing approximately 2 L of tap water at a density of 200–300 eggs per rearing tray. Larvae were fed a 1:1 mixture of lactalbumin and yeast ad libitum. Pupae were transferred from larval rearing trays to water-filled plastic cups in 30.5 x 30.5 x 30.5 cm Bug Dorm adult rearing cages (BioQuip, Rancho Dominguez, CA, USA). Cotton rounds soaked with 10% sucrose solution were placed inside each cage as carbohydrate source for emerging adults.

CDC Bottle bioassays. The CDC bottle bioassay is commonly used to monitor mosquito insecticide susceptibility¹⁹. We adapted this method to calibrate the assay for Culicoides by determining the DD and DT for permethrin. Following the CDC protocol¹⁷, we prepared permethrin solutions by diluting technical grade permethrin (100% Chem Service Inc., West Chester, PA, USA) in American Chemical Society (ACS) grade acetone (Fisher Scientific, Hampton, NH). Each bioassay used five 250-ml clear Wheaton bottles with lids (DWK Life Sciences Inc., Millville, NJ), four coated with 1.0 ml of the permethrin solution and one untreated control coated with acetone only. Bottles were coated by swirling the solution inside and then left to dry on a hot dog roller machine with heat turned off to ensure even coating. Control bottles were capped and rolled before preparing bottles containing insecticide. All bottles were then left open to dry in a dark environment to prevent permethrin photodegradation. Preliminary observations showed high mortality of C. furens in control bottles when the bioassay was performed within five hours after the bottles were coated. We tested various drying times (5, 10 and 24 h) and found that 24 hours of drying were necessary to eliminate control mortality.

As recommended by Brogdon and Chan¹⁸, when calibrating the CDC bottle bioassay to determine DD and DT for a new insect, we prepared bottles with a range of permethrin concentrations to determine a possible DD. Based on the CDC’s recommended DD of 43 µg permethrin/bottle for susceptible Ae. aegypti¹⁷, and the small size of C. furens in comparison to Ae. aegypti, the permethrin concentrations we tested as possible DD were 43, 21.5, and 10.7 µg permethrin/bottle. The permethrin concentration that killed 100% of the wild C. furens between 30 and 60 minutes would be selected as the DD (dose that kills 100% of insects¹⁸.

Bottle bioassays were performed at the same time but in separate bottles for susceptible Ae. aegypti and wild C. furens. Mosquitoes and C. furens were aspirated by mouth into each bottle with the goal of 20 individuals per bottle. Mortality was recorded every five minutes for the first 15 minutes and every fifteen minutes until 2-h post-exposure¹⁸. After the 2-hrs of exposure, insects were transferred into 8-oz paper food cups with lids and generic mesh to prevent all insects from escaping. Insects were provided with a cotton round with water and mortality was then assessed at 24-h post-exposure²⁸.

Field cage trials. Two field trials were conducted between March and April, 2023 in Indian River County, FL (Table 1), targeting caged susceptible Ae. aegypti and/ or wild C. furens (Table 1). Permanone 30–30 (30% permethrin, 30% piperonyl butoxide) was applied at maximum rate (0.007 lb permethrin/acre) (Table 1) using a truck-mounted ULV aerosol generator that was operated by personnel of the Indian River Mosquito Control District. Spray missions were performed around dusk (1700h and 1800h) on a five-acre plot of low-cut grass at the Indian River County Fairgrounds.

Bioassay cages (Fig. 1) were made of cardboard rings (15.2 cm diameter, 2.5 cm width) covered with mesh^26,29. Three mesh types were tested (Fig. 1): stainless steel (SS) McMaster-Carr, Douglasville, GA), no-see-um netting (NN) (Seattle Fabrics, Inc., Seattle, WA), and fine nylon tulle fabric (Hobby Lobby Stores, Inc., Oklahoma, USA), hereafter called mosquito mesh (MM). SS and NN mesh were chosen because preliminary observations indicated they could prevent escape of C. furens (Cooper, unpublished data). Although the MM mesh has openings too large for Culicoides, it was included as it has been used successfully in mosquito cage trials³⁰.

Table 1

Summary of ULV permethrin adulticide field cage trials to evaluate the susceptibility of *C. furens* in Indian River County, Florida, USA. (SS) stainless steel, McMaster-Carr, Douglasville, GA; (NN) no-see-um netting, Seattle Fabrics, Inc, Seattle, WA.; (MM) mosquito mesh (fine nylon tulle fabric), Hobby Lobby Stores, Inc, Oklahoma City, OK.
Field Trial	Date	Species tested	Mesh type tested	Number of passes		Wind speed
1	March 1	Ae. aegypti (susceptible)	SS, NN, MM	2	10 and 5 mph, respectively	3–5 mph
2	April 6	Ae. aegypti (susceptible)	SS, NN, MM	1	10 mph	6–10 mph
		C. furens (wild)	SS, NN

The mosquitoes-only field cage trial aimed to compare different mesh types for field bioassay cages. Approximately 20 susceptible Ae. aegypti were placed in each cage. After confirming that the mesh types allowed insecticide passage and caused mortality in these mosquitoes, a combined mosquitoes and Culicoides trial was conducted. Each cage contained about 30 wild C. furens and 20 Ae. aegypti, except for the MM cages, which only had Ae. aegypti (Table 1). This design allowed us to evaluate the susceptibility of wild C. furens to permethrin relative to susceptible Ae. aegypti.

Cages were hung approximately 1.3 m above ground on shepherd hooks in five clusters, each with the three mesh types (Fig. 1). Clusters were spaced every five meters along a north-south transect, 20 m downwind of the spray truck path. Two untreated (control) cages of each mesh type were placed 20 m upwind. Mortality was assessed in the field 10 minutes post-exposure, and subsequently recorded at 1, 12 and 24 h post-exposure in the laboratory. Mortality was based on knockdown (inability to stand on legs or have coordinated flight¹⁷). Insects were provided a cotton round with tap water and maintained in the cages for 24 hours, before being placed in dry ice to kill any remaining live insects.

Data Analysis. Results from the cage field trials were analyzed by calculating the mean mortality of mosquitoes and Culicoides at various times post-exposure. For each trial, mean mortality and standard deviation were plotted³⁰. For CDC bottle bioassays, time-response survival curves were created for each permethrin dose. Insect survival was assessed using Cox proportional hazards models with the ‘survival’ package in R version 4.2.2³¹. Clustered Cox regression was performed to compare mean mortality across the different permethrin doses for both mosquitoes and Culicoides³².

CDC Bottle Bioassays. Susceptible Ae. aegypti mosquitoes reached complete mortality at 30 minutes post-exposure in CDC bottle bioassays performed with the high and medium permethrin doses (43 and 21.5 µg /bottle), while these same doses caused complete mortality of C. furens at five minutes post-exposure (Fig. 2). Significantly greater mortality was observed in C. furens exposed to the high dose and medium doses in comparison to mosquitoes (P < 0.0001). The low permethrin dose (10.75 µg /bottle) caused complete mortality in Ae. aegypti and C. furens at 45 and 30 minutes, respectively (Fig. 2). No significant differences were observed in the mortality of C. furens and Ae. aegypti mosquitoes exposed to the low dose (P = 0.3699).

The susceptible Ae. aegypti mosquitoes and wild C. furens did not recover at 24 hours with any of the three permethrin doses, with the exception of C. furens from bottles treated with the low dose, in which five percent recovery was observed at 24 hours. However, mortality in untreated control bottles was significantly lower in comparison to treated bottles (P < 0.0001). We did not observe significant differences in the mortality of susceptible mosquitoes in comparison to wild C. furens (P = 1.0000). Abbot’s formula was not used to correct percent mortality in any of the bottle bioassays because control mortality at two hours did not exceed 3%¹⁷.

Mosquito-only field cage trial. All three mesh types allowed the passage of Permanone 30–30, resulting in mosquito mortality (Fig. 3A). In the mosquito-only trial, two adulticide passes were performed because no mortality was observed within 10 minutes after the first pass. We were unaware whether the insecticide had not gone through the mesh during the first pass or there was another reason why the insects were not experiencing mortality. A second pass performed within 20 minutes after the first application resulted in immediate mortality of the exposed mosquitoes.

At one-hour post-application, mortality rates were 89% (SS), 97% (NN), and 98% (MM) (Fig. 3A). Complete mortality was observed at 12 hours in SS and MM cages (Fig. 3A), while NN cages only reached 96% at 12 hours, with no further increase by 24 hours. Mosquitoes did not recover from the adulticide effects within the 24-hour period. Control cages showed minimal mortality at 24 hours, with 15% in MM cages and 5% and 6% in SS and NN, respectively. Since control mortality was only observed after 24 hours, Abbot’s formula was not applied to correct the percent mortality in treatment cages¹⁷.

Mosquitoes and Culicoides field cage trial. The second trial confirmed that bioassay cages constructed with SS and NN could be used to evaluate Culicoides susceptibility to ULV adulticides. Complete mortality of Ae. aegypti occurred in all bioassay cages at 1-hr post-application (Fig. 3B), while 100% mortality of C. furens was observed at 10 minutes. Mosquitoes and C. furens did not recover from the effect of the adulticide within the 24-hour period. Mortality in untreated control cages was only observed at 24 hours for Ae. aegypti placed inside SS cages and biting midges in SS cages (6%) and NN cages (16%). Because 100% mortality was observed in all treatment cages 1-hr post-exposure and no mortality was observed in the control cages until after 24 hours post-exposure, Abbot’s formula was not used to correct percent mortality in treatment cages.

In this study, we demonstrated that both the CDC bottle bioassay and field cage trials can be effectively adapted to assess insecticide susceptibility and efficacy of ULV treatments against Culicoides species. Obtaining this information is advantageous for protecting the health of humans and other animals. Our findings include the first records of DD and DT for Culicoides in CDC bottle bioassays, which can serve as reference for other species, such as C. paraensis, the main vector of OROV. We also identified two suitable mesh types for field cage trials with Culicoides. We observed mortality in C. furens in CDC bottle bioassays using technical grade permethrin and in the field cage trial with Permanone 30–30, which suggests that wild C. furens from Indian River County are susceptible to permethrin.

Despite the widespread use of adulticides to control Culicoides spp., their susceptibility to these treatments remains poorly understood. This study highlights the utility of field cage trials in assessing the effectiveness of adulticide applications against both vector and nuisance species. Although C. furens is not a vector of pathogens, this nuisance species can significantly impact the economy. For instance, in Australia, Culicoides nuisance is estimated to reduce residential property values by 8,300 USD per residence based on actual sale price³³. Ratnayake and co-authors also suggest that Culicoides prevent urban development in coastal areas, with over half of surveyed residents of Hervey Bay, Australia considering the impact of Culicoides a major factor in housing investment decisions³³.

We were able to determine a potential DD and DT using wild Culicoides. Typically, DD and DT used in CDC bottle bioassays are established using susceptible populations⁶. However, despite numerous attempts to establish Culicoides colonies since 1921³⁴, only C. sonorensis is colonized in the United States²⁵. Moreover, the susceptibility of this colonized C. sonorensis to permethrin remains undocumented. In the absence of a susceptible Culicoides population for calibration, we adapted the CDC bottle bioassay for wild C. furens. We found that 10.75 µg permethrin per bottle resulted in 100% mortality of C. furens from Indian River County, FL, within 30 minutes. Therefore, we recommend using 10.75 µg permethrin per bottle and 30 minutes as the DD and DT for this and potentially other wild Culicoides species.

We expect that the time to reach complete mortality in CDC bottle bioassays and cage field trials will vary between studies due to various factors. For example, different mortality rates may be observed in cage field trials using different permethrin formulations, even when applied at the same dose. We anticipate that differences in the time to reach complete mortality in CDC bottle bioassays could occur even within the same species when examining different populations. Although we observed complete mortality of C. furens from Indian River County, FL within 30 minutes in CDC bottle bioassays, different results could be observed in tests conducted with C. furens collected in locations where selection pressure is different.

Culicoides furens from Indian River County, FL, were more susceptible to permethrin than the Ae. aegypti used as a reference for susceptibility in both CDC bottle bioassays and field cage trials. The permethrin DD for Ae. aegypti has been established as 43 µg permethrin per bottle, which is expected to cause complete mortality at 10 minutes¹⁷. In our study, C. furens exposed to 43 µg permethrin per bottle were knocked down within seconds after being placed inside the bottle. Similarly, in our field cage trials, C. furens experienced complete mortality within 10 minutes, in comparison to one hour for Ae. aegypti. These disparities in the time to complete mortality is likely due to the differences in size between the Ae. aegypti from our colony (3.03 mm wing length)³ and the wild C. furens (0.91 mm wing length)³⁴. The efficacy of ULV insecticide applications against adult mosquitoes is related to the droplet size³⁵. Therefore, application rates on mosquito control product labels are based on the dose needed in a droplet to kill a mosquito based on its size³⁶. In this case, C. furens is a smaller insect and the dose needed to kill it is much lower, resulting in the high mortality we observed in C. furens using a mosquito-specific ULV adulticide application rate.

Our results are the first published information comparing SS and NN to perform field cage trials to quantify susceptibility of Culicoides to permethrin adulticide products in the United States. Linley and Jordan²¹ evaluated the efficacy of thermal fog applications with formulated products contained naled against C. furens. However, the authors highlight that this product was not the most effective compound against Culicoides and that pyrethroid applications including a permethrin formulated product were likely to be more effective¹⁹. Complementary studies to these tests, showed that naled applied with a vehicle mounted ULV machine was not effective against C. furens and that this was possibly due to the fine mesh used to confine Culicoides, which did not allow the insecticide to pass through²⁰. Although we found that cages made with SS and NN allowed the passage of ULV permethrin droplets, resulting in C. furens mortality, we recommend using NN since this material is easier to handle and at least 11 times cheaper than SS (NN: USD 5.44/m²; SS: USD 62.20/m²).

Understanding whether or not Culicoides biting midges are susceptible to the adulticides that are commonly applied for their control provides the opportunity to evaluate current control practices and to potentially replace ineffective methods with more successful, affordable, and sustainable alternatives. We suggest using the methodologies adapted for biting midges in this study for CDC bottle bioassays and field cage trials to evaluate permethrin susceptibility of C. paraensis, the OROV vector. This information will inform whether permethrin applications that may be performed by vector control agencies can also offer protection against the OROV vector. These methodologies could also be used to evaluate insecticide susceptibility in EHDV and BTV vector species on livestock farms. Wild Culicoides species of medical or veterinary importance could be targeted with ULV permethrin applications, similar to those commonly used for controlling Culicoides on deer farms¹⁶. These field-collected Culicoides spp. could also be exposed to a CDC bottle bioassay with technical grade permethrin at the DD and DT we suggest (10.75 µg /bottle at 30 minutes). Although CDC bottle bioassays are generally performed in laboratory settings, the protocol described here is a relatively simple, rapid, and economic test that can be performed in temperature-controlled areas outside the laboratory.

Our results highlight the opportunity of monitoring insecticide resistance as part of an integrated vector management (IVM) program against Culicoides. Effective insecticide resistance management should accompany ULV adulticide applications to ensure long-term control of Culicoides. IVM programs for blood-feeding dipterans often require using multiple control strategies to relieve the biting pressure³⁷. Given the limited alternatives for Culicoides control, effective management relies on accurate susceptibility assessments to ensure that adulticide applications remain effective and to prevent the development of insecticide resistance.

Data Availability

The dataset used and analyzed during the current study are available from the corresponding author upon reasonable request.

Competing Interests:

The authors declare no competing interests.

Author Contribution

V.C., N.B.C, and E.B. designed the experiments. Y.J coordinated and provided resources for field trials. V.C., N.B.C, and E.B carried out the experiments and conducted the data analysis. V.C wrote the manuscript. All authors reviewed and provided feedback for the manuscript.

Acknowledgement

The authors would like to thank Sarah McInnis, Victor Recendez, Heather Whitehead (Indian River County Mosquito Control District), and Morgan Rockwell (UF/IFAS Florida Medical Entomology Laboratory) for their support during field cage trials. We thank Ana Romero-Weaver for training VMC in CDC bottle bioassays and Tanise Stenn for rearing mosquitoes used in this study. Funding for this work was provided by the Florida State Legislature, through the Cervidae Health Research Initiative and NIFA Project FLA-FME-006106.

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No competing interests reported.

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Laboratory and field assays indicate that a widespread no-see-um, Culicoides furens Poey is susceptible to permethrin

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