Experimental parameters affecting the outcomes of amitraz resistance testing in Varroa destructor

Abstract

Honey bee colony health is impaired by a large variety of biotic and abiotic factors. The severe impacts of Varroa destructor and viruses associated with it on honey bees are undeniable and universal. Beekeepers often utilize miticides to mitigate V. destructor populations, but resistance to many widely used synthetic miticides such as amitraz has developed. The Apivar® resistance test has proven to be a valuable tool to monitor for amitraz resistance in V. destructor. However, a more thorough understanding of the effects that changes in testing parameters have on the outcome of the Apivar® resistance is critical for an accurate and comparable interpretation of the results. In this project, the effects of temperature, product age, product exposure surface area, and container reuse on the Apivar® resistance test were studied. High temperature significantly increased control mortality while low temperatures significantly increased Apivar® resistance. The increased Apivar® resistance at low temperature was due to reduced amitraz toxicity at low temperature. There was no association of Apivar® age or manufacturing batch on the outcome of the Apivar® resistance test. The maintenance of Apivar® efficacy with age is likely because the amitraz breakdown product N‘-(2,4-dimethylphenyl)-N-methylformamidine (DPMF) is as toxic as amitraz. The rate and efficacy of the Apivar® resistance test were higher with a larger surface area of Apivar® strip. Reused testing containers or Apivar® strips from colony application yielded reduced efficacy and consistency. This research shows the standard conditions needed to produce comparable data from the Apivar® resistance test. Implications for amitraz resistance monitoring are discussed.

Keywords:

• Honey Bee
• Varroa destructor
• amitraz
• DMPF
• bioassay
• resistance monitoring

Introduction

Since its transition from its native host, the eastern honey bee (Apis cerana Fabricius) in southeast Asia in the late 1940s, the parasitic mite, Varroa destructor, has had a devastating impact on western honey bee (Apis mellifera Linneaus) colonies around the world (Rosenkranz et al., 2010; Traynor et al., 2020). V. destructor is found globally as its range expansion is facilitated by worldwide shipment of infested honey bees and beekeeping equipment. Its ability to evade stringent inspection and quarantine practices has allowed V. destructor to establish in highly regulated regions such as Hawaii (Rusert et al., 2021) and Australia (Calver, 2022). The cosmopolitan nature of V. destructor infestation of honey bee colonies is a critical global concern.

V. destructor injures honey bees through physical damage, physiological impairment, and viral transmission. V. destructor feeds on fat body of pupal and adult honey bees (Ramsey et al., 2019). The fat body performs a tremendous diversity of physiological processes such as lipid storage, hormone synthesis, xenobiotic detoxification, vitellogenin production, and protein metabolism (Arrese & Soulages, 2010). Fat body content and its associated physiological outputs are important indicators of overwintering success (Amdam et al., 2004; Döke et al., 2015). V. destructor infestation results in honey bees with compromised immune response, reduced weight, premature foraging, and shortened adult life span (Dainat et al., 2012; De Jong et al., 1982; Kralj & Fuchs, 2006; Yang & Cox-Foster, 2005). V. destructor also transmits many honey bee viruses such as Deformed wing virus (DWV) and Chronic bee paralysis virus (CBPV) which has significantly amplified their deleterious effects on honey bee colonies (Chen & Siede, 2007; Martin et al., 2012).

Scientific research and beekeeper surveys agree that V. destructor is a major cause of honey bee colony losses around the world (Kulhanek et al., 2017; Traynor et al., 2020). Beekeepers suffer significant financial and labor costs related to V. destructor management practices (USDA NASS., 2022). A variety of integrated pest management tactics such as use of V. destructor-resistant honey bee stocks, drone brood trapping of V. destructor, induced brood break via caging queens, powdered sugar grooming stimulation, and screened bottom boards are used by beekeepers to maintain low V. destructor infestation (Jack & Ellis, 2021). However, miticide application represents the most frequent and often exclusive method for V. destructor management. In the US, synthetic miticides such as tau-fluvalinate (Apistan®), coumaphos (CheckMite^TM), and amitraz (Apivar®) along with organic acids such as oxalic acid (Api-Bioxal), formic acid (Mite Away QuickStrips®), and plant-derived molecules such as thymol (Apiguard®) and hop beta-acids (HopGuard 3®) are available for V. destructor control. Historically, synthetic miticides are the most commonly applied type of miticide due to consistent formulation, ease of use, and efficacy. However, V. destructor has developed resistance to many of these compounds presumably due to frequent applications and exclusive use of these miticides without rotation of miticides with other modes of action or using the product inconsistent with labelled instructions. Resistance to tau-fluvalinate and coumaphos were reported shortly after their introduction (Elzen, 1998; Elzen & Westervelt, 2002; Pettis et al., 1998). Amitraz resistance was initially reported around 2000, but has mostly remained effective since the initial report of resistance (Elzen, 1999). Informal and unverified reports of amitraz resistance from beekeepers over the past 5 years provided the basis to formally study amitraz resistance in V. destructor. Initial evaluation of amitraz resistance from beekeeping operations showed that Apivar® resistance in field tests was due to reduced sensitivity to amitraz in laboratory bioassays, thus establishing the Apivar® resistance field test is a reliable indicator of bona fide amitraz resistance (Rinkevich, 2020). Annual surveys of amitraz resistance in beekeeping operations from across the US have used the Apivar® resistance test to evaluate amitraz resistance due to its low cost, ease of use, and rapid assessment of amitraz resistance (Rinkevich, Manuscript in Preparation). Due to variation in experimenter, sampling date, temperature, apiary location, sampling methods, etc., it is critically important to understand how variations in test parameters influence the outcomes of the Apivar® resistance test and affect the application of the results to make effective V. destructor management decisions.

Materials and methods

Honey bees and V. destructor

Honey bee colonies were maintained at the USDA-ARS Honey Bee Breeding, Genetics, and Physiology Lab (HBBGPL) in Baton Rouge LA (30.379143, −91.167020). A total of 50 V. destructor-susceptible Italian queens were purchased from a commercial queen producer and installed into queenless colony divisions housed in 8-frame Langstroth hive bodies on 2022-April-7. Additional brood boxes and honey supers were added based on the number of adult bees in each colony and during honey flow, respectively, at the beekeeper’s discretion. Colonies were maintained without supplemental nutrition or miticide treatments. V. destructor reared in honey bee colonies at the HBBGPL serve as the amitraz resistant reference population because they are not treated with any miticides including amitraz and there is very limited importation of honey bee colonies and associated V. destructor. The potential for V. destructor migration due to drift into colonies maintained at the HBBGPL is very low since there are no other beekeepers within a 6 km radius and proximity to suburban development and the Mississippi River act as physical barriers to migration. The suitability of this V. destructor population to serve as a susceptible reference stock is reflected in its high amitraz sensitivity, low Apivar® resistance, and susceptible genotype (Rinkevich, 2020; Rinkevich et al., 2023).

Bioassays

Apivar® resistance test

The Apivar® resistance test was performed as previously described (née, Apivar® efficacy test (Rinkevich, 2020; Vu et al., 2020)). Briefly, bees from 2 frames containing sealed brood from each colony were shaken into an 18.9 L bucket. A 0.11 L plastic cup was used to scoop approximately 300 bees from the bucket into a 0.94 L plastic container with a 4 cm x 4 cm square of Apivar® glued perpendicular to the bottom. Control containers did not contain an Apivar® square. A lid with 0.64 cm mesh screen was used to seal the container. Four small binder clips were attached to the rim on the container that was then suspended over a 15.2 cm wide square plastic weighing dish coated with a thin layer petroleum jelly. The container was held at room temperature (22–24 °C) for 3 h, after which the container was turned over to stop the test. The number of V. destructor on the weighing dish was recorded. A drop of liquid dish detergent was added to the container through the screened lid. Bees were knocked to the bottom by pounding on a hard surface. Warm water was added through the screen lid to submerge the bees. The screened lid was replaced with a solid lid. The container was aggressively agitated by hand for 30 s and allowed to rest for 1 minute. This cycle of shaking was repeated twice for a total of 3 agitations. The solid lid was replaced with a screened lid. The contents of the container were poured through the screened lid through a 10 cm fine mesh strainer to collect V. destructor that were dislodged from the bees. The bees in the same container were rewashed and V. destructor collected again as described above but with one 30 s agitation until no more soap was visible in the sample. The total number of V. destructor in the fine mesh strainer was counted as the number of V. destructor in the wash. The number of V. destructor in the wash was added to the number of V. destructor on the weighing dish to determine the total number of V. destructor in the sample. The Apivar® resistance level was calculated by dividing the number of V. destructor dislodged in the soapy water wash (i.e., amitraz resistant V. destructor) by the total number of V. destructor in the sample. The Apivar® resistance level was corrected for control mortality using Abbott’s formula (Abbott, 1925). Detailed instructions, parts list, and data sheets can be found in the Supplemental Information. A video of the Apivar® resistance testing procedure can be seen at https://www.youtube.com/watch?v=-YUyj_RrtB0&ab_channel=USDA-ARS.

Amitraz toxicity

The toxicities of amitraz and its break down products N′-(2,4-dimethylphenyl)-N-methylformamidine (DPMF) N-(2,4-dimethylphenyl)formamide (DMF), and 2,4-dimethylaniline (DMA) were tested (Korta et al., 2001). Amitraz, DMF, and DMA were ordered from Sigma (Milwaukee WI). DPMF was ordered from ChemService (Westchester PA). All materials were > 97% purity. Stock solutions were made via serial dilution in acetone to create concentrations that resulted in > 0 but < 100% mortality (Table S1).

Glass vial bioassays were performed as previously described (Rinkevich, 2020). Twenty mL glass vials (Research Products International) were treated with 0.5 mL of stock solution and rolled for > 1 h to complete dryness in a fume hood on an unheated hot dog roller (HOTDG-V005, VIVO Electric). Control vials were treated with acetone.

V. destructor were collected by large-scale sugar shakes as previously described (Rinkevich, 2020). V. destructor were sampled from 4 or more colonies out of the 50 initial colonies that were set up each day the bioassays were performed in order to collect >300 V. destructor needed for the bioassays. Briefly, bees were shaken from 3 combs with sealed brood into a 22 L plastic bucket. The bees were coated with approximately 0.1 L of powdered sugar. The bucket was covered and gently inverted approximately 5 times and V. destructor were allowed to dislodge for 5 minutes. V. destructor were collected by dumping the bees and the powdered sugar through a large mesh screen to allow the V. destructor and powdered sugar to fall into a bucket. The bees where then returned to the colony. V. destructor were separated from the powdered sugar by passing the V. destructor and powdered sugar through a fine mesh sieve and consolidated in to a 4.5 L holding bucket in the field. V. destructor were collected from four or more colonies each day bioassays were conducted. Sugar was removed in the lab by rinsing the V. destructor in a sieve with water then placed on a tray covered with paper towels. A white-eyed honey bee pupa was added to each treated bioassay vial and then 10 V. destructor were added to the vial using fine hair brushes. Vials were sealed with parafilm and approximately 20 holes were punched through the film with a fine needle. Vials were placed on their side on a tray and secured with a rubber band. Under standard experiments, trays with treated vials were placed in an incubator (INR030, Darwin Incubators, St. Louis MO) at 33 °C (±0.5 °C with 75 ± 5% RH). Temperature and relative humidity in this and all other experiments were monitored with a Hobo MX2301A Temperature and Relative Humidity Logger (Onset Computer Corp., Bourne MA). V. destructor mortality was assessed 24 h later by emptying the contents of the vial in to a disposable 15.2 cm wide square plastic weighing dish. V. destructor that was not moving or unresponsive to prodding with a metal probe were recorded as dead. The LC₅₀ was calculated by probit analysis with a custom R script (Probit Analysis v6_1 (Silva, 2022),) using Abbott’s correction for control mortality (Abbott, 1925). LC₅₀ values were considered significantly different if there was no overlap in the 95% CI.

Modified testing parameters

Temperature

To study the effects of temperature on the Apivar® resistance test, incubators were used to house containers at 35 °C, 33 °C, 30 °C, 25 °C, 20 °C, 15 °C, and 10 °C for the 3-hour duration of the test in complete darkness. Bees and V. destructor were sampled from 10 to 15 colonies for each experiment. Samples were processed after the end of the experiment as described above.

The effects of temperature on amitraz toxicity was studied by placing vials treated with amitraz in incubators set at 35 °C, 33 °C, 30 °C, 25 °C, 20 °C, 15 °C, and 10 °C in complete darkness for the 24-hour exposure duration. All other aspects of the amitraz toxicity bioassays were the same as described above.

Apivar® strip age

The age of Apivar® strips on efficacy was studied using batches of Apivar® strips manufactured on different dates that were open and stored in their original packaging inside of a plastic storage container in the lab at 22 °C–24 °C. Information on Apivar® batch number, manufacturing date, testing date, and strip age can be found in Table S2.

To test the efficacy of reused strips that were previously installed into a colony, five Apivar® strips that were manufactured on 2020-Sept-21 were installed into colonies on 2021-July-11 to treat for V. destructor. Strips were removed from the colonies on 2021-August-20 and left on that ground at the apiary until they were collected on 2022-May-5. The strips were stored in the lab in a zip top bag until used in the Apivar® resistance test under normal conditions during the week of 2022-Sept-19. Two 4 cm x 4 cm squares were cut from each of the five strips. The Apivar® resistance test was performed at 22 °C–24 °C as described above using bees and V. destructor collected from 10 different colonies.

Reused containers

The effect of container reuse in the Apivar® resistance test was measured by reusing test containers after the bees were washed with soapy water. Containers were reused twice for a total of three uses. Bees and V. destructor were collected from 15 colonies for tests in cups used once and 10 colonies were used for tests in cups reused 2 and 3 times. Apivar® resistance tests with these reused containers were performed at 22 °C–24 °C as described above.

Surface area

The impact of the surface area of the Apivar® strip was evaluated by using Apivar® strips of varying size. The normal test conditions call for a 4 cm x 4 cm square of an Apivar® strip (i.e., the strip is cut as long as it is wide, 5 squares per Apivar® strip). This size of Apivar® strip is referred to as the full-size strip and provides 32 cm² of surface area (i.e., 2 sides of 4 cm × 4 cm) when glued perpendicularly to the bottom of the test container. The double size strip was 8 cm long and 4 cm wide to provide 64 cm² of surface area. The half size strip measured 2 cm long and 4 cm wide to provide 16 cm² of surface area when glued perpendicularly to the bottom of the test container. Quarter size strips were 2 cm long by 2 cm wide and provided 8 cm² of surface area. Bees and V. destructor were collected from 15 different colonies for all treatments except the quarter size strip was evaluated from 10 colonies. The Apivar® resistance test was performed at 22 °C–24 °C as described above. The number of V. destructor on the trays was recorded every 15 minutes over the 3-hour exposure duration. Apivar® resistance was calculated at each 15-minute interval for each area of exposure. The LT₅₀ for each size of Apivar® surface area was calculated by plotting time against Apivar® resistance in SigmaPlot v11.2.0.5 (Systat Software, Palo Alto, CA) and fitting the data points with a 2-parameter logarithmic function and using the equation of the line to calculate the time at which Apivar® resistance equals 50%.

Results

Temperature affected the outcomes of the Apivar® resistance test in the control and experimental containers. Control mortality increased up to nearly 22% mortality with temperatures >30 °C while control mortality was <2% at temperatures <30 °C (Figure 1A). There was a significant exponential relationship of temperature and control mortality (Exponential Growth, Single, 2-parameter, R²=0.711, F_1,69=137.731, p < 0.0001). Apivar® resistance significantly increased at temperatures <20 °C while amitraz resistance was < 1% at temperatures >20 °C (Figure 1B). There was a significant linear correlation of temperature with Apivar® resistance (Linear Regression, R²=0.267, F_1,84=30.482, p < 0.0001). Data on the effects of temperature on control mortality and Apivar® resistance can be found in Table S3).

Figure 1. Temperature significantly affects control mortality and Apivar® resistance in the Apivar® resistance test. (A) Control mortality increased significantly at temperatures ≥30 °C. (B) Apivar® resistance increased significantly at temperatures ≤20 °C. In both figures, different letters indicate significant differences.

Temperature influenced amitraz toxicity in the glass vial bioassays. The amitraz LC₅₀ was significantly higher at temperatures ≤15 °C, resulting in approximately 1.6 to 6.2-fold decrease in amitraz sensitivity (Figure 2A, Table S4). There was a strong correlation of amitraz LC₅₀ with the log of temperature (Linear Regression, R² = 0.549, F_1,6 = 6.081, p = 0.056). There was a significant correlation of the amitraz LC₅₀ with Apivar® resistance along the range of temperatures tested (Linear Regression R² = 0.233, F_1,84 = 26.30, p < 0.0001).

Figure 2. Toxicity of amitraz at different temperatures and comparative toxicity of amitraz and DMPF. (A) Temperature significantly affected amitraz toxicity in the glass vial bioassay. The amitraz LC₅₀ was significantly higher at low temperatures. Different letters indicate significant differences. (B) DMPF is a toxic breakdown product of amitraz. There was no significant difference in the toxicity of amitraz and DMPF. The LC₅₀ of DMA and DMF are >1000 μg/vial and 181.7 μg/vial, respectively (data not shown). A summary of bioassays with amitraz, DMPF, DMA, and DMF can be found in Table S5.

Apivar® strips stored in their original but open package had similar levels of efficacy in the Apivar® resistance test. There was no correlation of Apivar® resistance with the age of the Apivar® strip that were new from their package (Linear Regression, R²=0.0008, F_1,48=0.042, p = 0.839). While there were some statistical differences between the age of some strips, the magnitude of the differences is trivial (i.e. < 1.5% range in Apivar® resistance among groups; Wilcoxon Each Pair, p < 0.05; Figure 3). Apivar® strips that were reused after application at the colony level yielded highly variable outcomes in the Apivar® resistance test that were significantly different from all new strips tested regardless of age (Figure 3). A table of the statistical output for all of the comparisons of Apivar® strip age mentioned above can be seen in Table S2.

Figure 3. Age of Apivar® strips does not dramatically affect the outcome of the Apivar® resistance test. All new strips showed <1.75% Apivar® resistance regardless of age. Strips that were reused after being placed in the colony for treatment were not reliable in the Apivar® resistance test. Different letters indicate significant differences.

The toxicity of amitraz and its breakdown products yielded various outcomes. The LC₅₀ for amitraz was not significantly different than the LC₅₀ for DMPF (Figure 2B). The LC₅₀ for DMF was more than 21,000-fold higher than the LC₅₀ for amitraz. DMA did not induce any mortality at concentrations up to 500 mg/vial. Therefore, DMA is >116,000-fold less toxic than amitraz to V. destructor. A data table of bioassay results with amitraz, DMPF, DMA, and DMF can be found in Table S5.

Reusing containers in the Apivar® resistance test showed inconsistent outcomes and increased variation. Reusing the cups for the Apivar® resistance test significantly increased Apivar® resistance from 0% in the first use to 1.9% and 3.2% in the second and third use, respectively (Kruskall-Wallis, X²=12.946, df = 2, p = 0.0015). Raw data and statistical outputs from the reused container tests can be seen in Table S6.

The size of the Apivar® strip used in the Apivar® resistance test significantly affected the LT₅₀ and final efficacy. There were significant differences in the LT₅₀ values among the surface area of Apivar® exposure (Kruskall-Wallis, X²=28.07, df = 3, p < 0.0001, Figures 4A,B). There was a significant negative correlation of the LT₅₀ with the log of surface area of the Apivar® strip (Linear Regression, R² = 0.440, F_1,52=40.89, p < 0.0001, Figures 4A,B). The Apivar® resistance at the end of the 3-hour exposure was significantly different among the surface area of Apivar® exposure (Kruskall-Wallis, X²=16.17, df = 3, p = 0.001, Figure 4C). There was a significant negative correlation of the surface area of the Apivar® strip with Apivar® resistance at 3 h (Linear Regression, R² = 0.210, F_1,54 = 14.05, p = 0.0004). The data summary and statistical outputs for the Apivar® LT₅₀ and 3-hour Apivar® resistance for different sized Apivar® strips are found in Table S7.

Figure 4. Size of the Apivar® square affects the rate and outcome of the Apivar® resistance test. (A) Apivar® resistance over time with different sizes of Apivar® strip. (B) Comparison of the Apivar® LT₅₀ with variable sizes of Apivar® strips. Letters above bars indicate significant differences in the Apivar® LT₅₀ between different sized Apivar® strips. (C) Apivar® resistance at 3h endpoint with Apivar® strips of different sizes. Letters above bars indicate significant differences in the Apivar® resistance at 3-hour endpoint between different sized Apivar® strips.

Discussion

Temperature can significantly impact the outcomes and interpretation of the Apivar® resistance test. At high temperatures, high control mortality would make truly amitraz resistant V. destructor appear susceptible, thereby underestimating amitraz resistance. This is supported by the observation that V. destructor with a resistant genotype infrequently appear with a susceptible phenotype (Rinkevich et al., 2023). When the temperature was too low, there was an increase in amitraz resistance, leading to an overestimate of amitraz resistance. Therefore, it is important to conduct the Apivar® resistance test at temperatures between 20 °C and 30 °C for the most accurate and comparable results.

The influence of temperature on V. destructor falling off of bees is a well-documented phenomenon. Nearly all V. destructor fell off bees held in cages held in an incubator at 40 °C for 48 h (Harbo, 2000). High V. destructor fall (85%–90%) can also be induced when V. destructor on bees were exposed to 46 °C–48 °C for as little as 2 minutes (Komissar, 1987). The impact of temperature on V. destructor extends to V. destructor reproduction and survival in brood cells where reproduction is impaired at temperatures above 36.5 °C and mortality significantly increases above 38 °C (Le Conte et al., 1990). Heat induced V. destructor mortality is the basis for the development of a number of devices that use heat to control V. destructor (Goras et al., 2015; Huang, 2001; Porporato et al., 2022). While little peer-reviewed research on the effectiveness of commercially available hyperthermic devices exists, the use of heat as a non-chemical means of V. destructor control remains an exciting prospect (Jack & Ellis, 2021).

Temperature had a significant influence on amitraz toxicity where amitraz was less toxic at lower temperatures. This shows that the higher Apivar® resistance at low temperatures is due to reduced amitraz toxicity and not reduced honey bee activity at lower temperatures where bees tend to cluster on the Apivar® square. While this positive temperature coefficient for amitraz is the first report of such a phenomenon in V. destructor, the influence of temperature on pesticide toxicity is a widely studied field of research, at least historically (Scott, 1995). This reduced amitraz toxicity may lead to reduced treatment efficacy with amitraz over the winter when temperatures within the colony may dip below 20 °C (Meikle et al., 2017). However, more research is needed to verify this potential amitraz inefficacy at the colony level.

There was a strong correlation of Apivar® resistance with the amitraz LC₅₀ at the respective temperatures. This confirms previous results of the correlation between these parameters when compared across V. destructor populations with varying levels of Apivar® resistance and amitraz toxicity (Rinkevich, 2020). This reproducibility shows that Apivar® resistance and amitraz toxicity are intimately related along environmental and genetic gradients and lends a high level of confidence that the Apivar® resistance test is a reliable indicator of amitraz sensitivity.

Apivar® strips stored in their open and original package in the lab showed no significant losses of efficacy more than 8 years since the strips were manufactured. There were no dramatic variations in the limited number of batch numbers of Apivar® that were tested. Therefore, the loss of Apivar® efficacy due to strip age or variation among batches is not supported by this data. It is likely that amitraz resistance explains a significant proportion of amitraz treatment failures.

A common explanation about the loss of Apivar® efficacy is that amitraz breaks down over time. This is especially true in humid conditions within a colony where amitraz breaks down into DMPF, DMF, and DMA to the point where amitraz is undetectable (Chaimanee et al., 2022; Korta et al., 2001). However, DMPF is just as toxic as amitraz to V. destructor as seen in the amitraz toxicity test in this study. Therefore, amitraz is a pro-miticide in that it needs to be converted into DMPF to exert its toxic effects in V. destructor and is consistent with previous observations in other arthropods (Salgado & David, 2017; Schuntner & Thompson, 1978). This pro-miticide concept was validated in studies with heterologously expressed octopamine receptors that showed DMPF was more potent and efficacious at activating octopamine receptors than amitraz (Guo et al., 2021). While it is true that amitraz does break down over time, the DMPF breakdown product is toxic to V. destructor.

Apivar® strips that were reused either after application at the colony or after washing the bees in the Apivar® resistance test led to inconsistent outcomes in the Apivar® resistance test that were significantly different from new, unused strips. These inconsistent results strongly discourage the reuse of Apivar® strips in the Apivar® resistance test and colony level treatments.

The larger surface area of the Apivar® strip increased the rate and efficacy of the Apivar® resistance test as demonstrated by the lower Apivar® LT₅₀ and Apivar® resistance at 3 h, respectively. Due to the relationship of Apivar® surface area with the rate and efficacy of the Apivar® resistance test, the Apivar® resistance test should be conducted for 3 h with a 32 cm² Apivar® square (i.e., 2 sides of 4 cm × 4 cm) so results are comparable across tests.

The effect of the Apivar® strip surface area on the Apivar® resistance test is important because previous methods used to test for miticide resistance recommended using a significantly lower surface area. Commonly referred to as “the Pettis test,” a section of Apistan® strip with 2.4 cm² of surface area was used (3/8“x1” or 0.95 cm × 2.54 cm strip of Apistan® stapled to an index card) and bees infested with V. destructor were exposed for 24 h to measure resistance to tau-fluvalinate (Pettis et al., 1998). This methodology was adapted for testing for Apivar® resistance and there is a wealth of historical data on Apivar® resistance using the Pettis test. A side-by-side comparison of the Pettis test and the Apivar® resistance test described here will validate if these methods yield similar conclusions about amitraz resistance in V. destructor.

This research shows the importance of standardized conditions on the outcomes and interpretation of the Apivar® resistance test. The established Apivar® resistance protocol should be followed consistently by using 4 cm x 4 cm square of Apivar® with a 3-hour exposure duration held between 20 °C and 30 °C without any modification to yield results that can be compared across studies. This research also demonstrates that Apivar® strips retain their efficacy for a very long duration when stored in their original package. While amitraz does degrade, the DMPF degradation product is the toxic molecule to V. destructor. Therefore, amitraz resistance and not Apivar® product integrity is the more likely cause of V. destructor control failure with amitraz. Reusing Apivar® strips is strongly discouraged by this research and prohibited by the label instructions.

Acknowledgements

The author would like to thank David Dodge and Nathan Egnew of the USDA-ARS Honey Bee Breeding, Genetics, and Physiology Lab for technical assistance in colony maintenance and laboratory assays. No external funds were acquired for this study. Mention of trade names or commercial products in this publication is solely for the purpose of providing specific information and does not imply recommendation or endorsement by the U.S. Department of Agriculture. USDA is an equal opportunity provider and employer.

Disclosure statement

No potential conflict of interest was reported by the author(s).

Additional information

Funding

Funding-No external funds were provided for this research.