Oils, Chemicals and Repellents

Coconut oil was purchased from Swanson Health Products Inc. (Fargo, ND, USA). The coconut oil fatty acids were obtained from ACME HARDESTY (Blue Bell, PA, USA). Catnip essential oil was purchased from Bramble Berry Inc. (Bellingham, WA, USA). Fatty acids, methyl laurate, and DEET standards were purchased from Sigma-Aldrich (St. Louis, MO, USA), which all had a purity >98%. The tested chemicals were diluted to various concentrations using either ethanol or hexane that were also purchased from Sigma-Aldrich (99–100%). Genu pectin DD-slow set Z was obtained from CP Kelco (Atlanta, GA, USA). Waxy cornstarch (Waxy No. 1) was obtained from A.E. Staley Mfg. Co. (Decatur, IL, USA).

Insects

Stable flies used for laboratory repellency tests were from colonies maintained at the United States Department of Agriculture (USDA), Agricultural Research Service (ARS), Agroecosystem Management Research Unit (Lincoln, NE, USA). The flies were maintained at 23 ± 2 °C with variable humidity (30–50% RH) and a light:dark photoperiod of 12:12 (L:D). Adult stable flies were fed with citrated bovine blood (3.7 g sodium citrate/liter) by soaking blood in a feminine napkin (Stayfree®, McNeil-PPC Inc., Skillman, NJ, USA) and placing it on top of the cage. Horn flies were shipped as pupae from an insecticide susceptible laboratory colony maintained at the USDA-ARS Knipling-Bushland US Livestock Insects Research Laboratory in Kerrville, TX, USA. Emerged horn fly adults were maintained under the same environmental conditions as stable flies, and fed in the same manner, with the exception that the blood-soaked pads were placed inside of their cages.

The bed bug strains were collected from human dwellings in Cincinnati, OH and New Jersey city, NJ, USA40. They were fed on defibrinated rabbit blood (Hemostat, Dixon, CA, USA) through a Parafilm™-membrane covered feeder which was heated to 39 °C with a circulating water bath. The bed bugs were maintained at 26°, 65 ± 5% RH, and a photoperiod of 14:10 (L:D) hr. Insects were evaluated 7–25d after emergence, and they had not been fed (for both nymphs and adults).

Unfed adult female and male lone star ticks were obtained from an in vitro colony at the USDA, ARS, Knipling-Bushland US Livestock Insects Research Laboratory. The colony was established from a Tick Rearing Facility at Oklahoma State University. All unfed adult ticks were maintained in an aquarium held at 27 ± 2 °C, 14:10 (L:D) photoperiod and sustained at 85% RH using a saturated salt solution. Engorged females of brown dog ticks were collected from naturally infested dogs in Goiânia, Goiás state, Brazil. A laboratory maintained tick colony was fed on rabbits (Oryctolagus cuniculus) using an apparatus glued in their backs. All free-living stages were maintained in a climatic chamber (27 °C and >80% RH). The ticks used in the experiments were aged between 7 and 21 days. The use of rabbits in this study was approved by the Committee on Ethical Animal Use of the Federal University of Goiás (CEUA/UFG, protocol number 024/2014). The care and use of the animals during this study were undertaken according to bioethics and animal welfare guidelines required by CEUA/UFG.

Mosquitoes used in this study were USDA strain Aedes aegypti reared in insectaries maintained at 26.6 °C, 85 ± 5% relative humidity (RH), and a photoperiod of 14:10 (L:D) h. Batches of 500 eggs were hatched in larval pans in 2.5 l of reverse osmosis water. Larvae were fed 1–3 g of liver and yeast mixture at a 3:2 ratio. Adult mosquitoes were supplied with 10% sucrose solution and a separate supply of reverse osmosis water.

Repellency assays

Biting flies

The laboratory bioassay for testing repellent efficacy on fly biting/feeding consisted of a six-well feeding reservoir system similar to the K & D module21,22. Unlike assays used for mosquitoes, our bioassay with stable flies required no warm water. Adult flies were fed with blood once, then being starved, but water was provided 24–48 hours before the repellency test began. Stable flies were starved for 48 hours prior to testing, and 24 hours for horn flies. On the day of the test, small squares of the feminine napkin pad (3.75 × 4.75 cm) were cut to fit into wells of the module. When testing stable flies, the pads were soaked with ~5 mL of citrated bovine blood (local abattoir). The outer layers of the feminine napkin pads were cut and used for coating repellent candidates, which was made of 2 layers of 100% cotton flannel and a layer of ultra-thin nylon. Repellent candidates measured at three dosages, 2 mg, 10 mg and 20 mg, respectively, were dissolved in 300 µL of hexane (Burdick & Jackson High Purity Solvent, Muskegon, MI, USA), and then topically applied onto the outer layer evenly (4 × 5 cm). After air drying, it was placed on top of the blood-soaked pad. Approximately 3–5 starved flies were collected from the fly cages and transferred into each testing cell. After 4 hours, tested stable flies (anesthetized with CO 2 ) were checked for feeding status by rupturing their abdomen to determine the presence of blood after the trials. Repellent assays were conducted daily at room temperature for at least 4 hours. Flies in the repellent bioassay were exposed to randomized treatments (different repellents and dosages), and repeated until at least 10 replicates were completed. Percentiles of repellency [(number of flies fed on control − number of flies fed in treatment)/number of flies fed on control × 100] was determined and transformed to arcsine square-root values for analyses of variance (ANOVA). Significant differences at P < 0.05 (SAS version 10; SAS Institute, Cary, NC, USA) were determined by analyses performed on the Least-Square Means due to the unequal number of observations among the treatments. Replicate numbers were determined by the number of treatments tested per day, and controls were always run simultaneously.

Dose-response repellent tests of coconut fatty acids and lauric acid against stable flies and horn flies were conducted using three different dosages described above. Hexane was used as the control. Results were analyzed as described above. The comparative study of using different ratios of the major repellent acids (C 8:0 , C 10:0 and C 12:0 ) was also conducted using the same procedures as described above, but only tested at the 20 mg dosage. The relative ratios of binary and three-component acid blends were based on the GC analyses of coconut oil fatty acids. These repellent bioassays were repeated for at least 6 replicates.

The longevity tests using coconut fatty acids and its starch-based formulation were carried out under laboratory conditions using the same repellent bioassay described above. Repellent layers loaded with coconut fatty acids or starch formulations (20 mg) were prepared inside a laboratory ventilation hood, and aged by hanging from a metal rack (1 m long) with metal clamps until all aged repellent layers were produced (1st to 4th day) that were run simultaneously with a total of 5 replicates of each treatments (at ages of 1st to 4th day-old plus controls, a positive control catnip oil was also tested to make sure the assay worked properly).

The least repellency concentration tests (LR 50 and LR 90 ) using coconut fatty acids and lauric acid against stable flies were carried out using various dosages (0.2 mg, 2 mg, 10 mg, and 20 mg). The bioassay was conducted using the same procedures described in the blood-feeding assay (modified K&D module). Hexane was used as the control. The experiment was repeated at least 5 times. A POLO PC program was used for Probit analysis of concentration-repellency data.

Bed bugs

Petri Dish Assay was used to quickly evaluate the comparative repellency of the repellent candidates and DEET (Rutgers University tests)29. Plastic Petri dishes (11.4 cm diameter by 3.8 cm height) were used as experimental arenas. For each arena, the inner wall coated with a thin film of fluoropolymer resin, bottom covered with a piece of filter paper. Filter paper was cut into two equal parts, one half was treated with a repellent using a Potter spray tower at 2.16 mg/cm2 of ethanol solution, the other half was sprayed with equal volume of 95% ethanol. A small piece of filter paper also was treated with the same repellent and folded into a tent shape with the treated surface facing down. The paper tent was placed on the repellent treated side, and the dish was uncovered. For the control treatment, one half of the filter paper and the tent was treated with 95% ethanol, the other half was untreated. 95% ethanol was used as solvent. 10% DEET (v/v), and 10% coconut fatty acids (m/m) were used to evaluate their repellency against bed bugs. All filter papers were treated on the same day, each kind of treated papers (95% ethanol-treated, 10% DEET-treated, or 10% coconut fatty acids-treated) were divided into three groups. They were kept in our laboratory (25 ± 1 °C, 20% relative humidity (RH)) for 0, 3, and 7days before experiment. The repellency of chemicals against bed bugs was tested at 0, 3, 7 days after application. Each filter paper was used only once. Five nymphs (fourth-fifth instar) and five males (age was unknown) were released in the center of each dish, the number of bed bugs on each side of the dish was recorded after 24 hours. All treatments were tested simultaneously. Each treatment was replicated 5–8 times. The assays were started between 3–4 hours into the dark cycle. Experiments were conducted in a walk-in chamber at 25 ± 1 °C, 20% RH, with a photoperiod of 12:12 (L:D). Repellency indices were calculated according to the formula: Repellency index = (C − T)/C × 100, where C = the mean number of bed bugs on the treated filter paper halves in all control dishes, and T = the number of bed bugs on the treated half of the filter paper in one test dish29,41. Repellency indices between the two compounds were compared using independent-samples T-test. (IBM SPSS Statistics 22.0, IBM, Armonk, NY, USA).

Behavioral responses of bed bugs were also tested using an indoor arena bioassay by another laboratory group (University of Kentucky test). The test materials were carried out in 9.5 cm inner diameter arena (ClimbUp Insect Interceptor™, Memphis, TN, USA). The inner well of this arena was covered with white filter paper (9.0 cm diam. Whatman, No. 2) that was fixed in place with double-sided tape to prevent bed bugs from getting beneath it. Test materials consisted of 20 µl of a 10% (w/w) hexane solution of DEET, coconut fatty acids, or hexane only. Test materials were applied to a 1.75 × 1.5 cm piece of filter paper that was pleated along the shorter midline to form a tent. The hexane was allowed to evaporate for 1 h before two tents were placed into the arena. Three different choice experiments were conducted: 1) Coconut fatty acids versus control; 2) DEET versus control; and 3) coconut fatty acids versus DEET. These test arenas were positioned in a wind tunnel so that different treatments were located across the wind line, and thus different treatments were isolated from each other. Arenas were placed on three levels of wire mesh shelf that was 0.7 m wide, 0.25 m deep, with 0.3 m between levels. The wind tunnel was 1 m wide by 0.9 m high by 2.4 m in length. The shelf and thus the arenas were placed only at the downwind end of the tunnel. The wind speed in the tunnel was 0.3 m/s with air evacuated from the laboratory through a fume hood. Tents were held in a separate fume hood for 0, 3, 7, and 14 days before being introduced into an arena. At about 9 h into the photophase, each bed bug was placed at the center point to the arena; equidistant from each tent. Room temperature during the choice test remained at 24 ± 2 °C. The position of bed bugs was noted at 16 h after release, which was 1 h after the initiation of the second photophase of the test. A total of 20 bed bugs were released with each treatment combination (at 0 d, 3 d, 7 d, and 14 d). Neither bed bugs nor tents were reused. Thus, a total of 240 insects were evaluated. The number of responses was analyzed by a binomial test using the null hypothesis that the two tents were chosen with equal probability. Insects that did not make a choice between the two tents (i.e., were wandering in the arena) were not included in the statistical analysis, but are shown in the figures.

Ticks

Repellency against nymphs of the lone star ticks was determined by using the vertical paper assay described previously30. A 4 × 7 cm rectangle of Whatman No. 4 filter paper was prepared by treating the central 4 × 5 cm zone with a volume of 165 µL of test solution. After drying, the paper strip was suspended from a bulldog clip hung from a holder. Ten lone star tick nymphs were released from a glass vial on the lower untreated end of the paper strip. Locations of the nymphs were recorded at 1, 3, 5, 10 and 15 min. Ticks were considered repelled if they stayed on the lower untreated zone or fell off the filter paper without having crossed into the upper untreated zone. Each treatment/concentration included three replicates.

For brown dog ticks, petri-dish bioassays were performed under controlled environmental conditions (at 27 °C and 70% RH) in complete darkness, based on the methodology described by Bissinger et al.31. The coconut fatty acid-treated filter papers were dried for 10 min under a fume hood prior to use in the assays. Six ticks (three males and three females) were placed in each arena along the line formed by the junction of treated and untreated papers. Control assays were made using clean paper versus clean paper. The positions of the ticks were evaluated at 24 h, 48 h, 72 h, 96 h and 1 week after the beginning of each experiment. Each experiment was replicated 10 times, with new ticks, for each individual compound.

In the Petri-dish bioassay repellency rates were determinate as the mean percentage ticks located on the untreated side of the Petri dish. The chi-square test was used for comparison of the tick choices, taking the significance level to be p < 0.05. When a higher significant proportion of ticks were found in the control side, the compound/concentration was considered as repellent.

Mosquitoes

Repellency was determined as the minimum effective dosage (MED, the minimum threshold surface concentration necessary to prevent mosquitoes from biting through the treated surface) of the coconut oil fatty acids to prevent bites through the fabric was first carried out at the USDA-ARS laboratory in Gainesville, FL. A 0.15 g sample of coconut oil fatty acids and DEET standard were added to prepare in 2 ml of solvent (acetone). Serial dilutions of the fatty acids and DEET were performed and each dilution was held in a separate vial. A 50 cm2 section of muslin cloth was added to each vial. Each volunteer wore each treated cloth to pinpoint the cloth which was treated with a concentration that failed (greater than or equal to 5 bites in one minute) and was next to an adjacent higher concentration passed (less than 5 bites in one minute). The lowest concentration passed was the MED for that test subject. Additional details on the bioassay methodology can be found in Carroll et al.32. There were three human volunteers in this study and all three provided written informed consent to participate in this study as part of a protocol (636–2005) approved by the University of Florida Human Use Institutional Review Board (IRB-01). A second repellent bioassay was conducted at Anastasia Mosquito Control Station in Florida. The three human volunteers aged from 30–60 involved in the second bioassay also followed the similar protocol approved by the Florida Human Use Institutional Review Board. An informed consent document was introduced to each individual volunteer in this study and received their approval for participation. Repellent treatments consisted of 1.0 mL of coconut fatty acids, lauric acid and DEET (control was also included), which was pipetted onto the forearms of the volunteers and applied from the wrist to the elbow by a gloved person to ensure full coverage of repellent. Approximately every 30 min from the start of the experiment each volunteer held their arm in a mosquito cage for 3 min. Protection failure was indicated by 2 mosquitoes landing and probing for more than 3 min on the treated area of the volunteer’s arm. Control volunteers were rotated with the other volunteers of the repellency test for 10 secs to 1 min to ensure that the mosquitoes still demonstrated attraction to hosts. The experiment was concluded after 6 h and 48 min when the last volunteer was probed for more than 3 min by 2 or more mosquitoes.

All methods related to mosquito repellency studies related to human participants from two laboratories were performed in accordance with the relevant guidelines and regulations approved by each institute.

Chemical analyses of coconut fatty acids

The fatty acids from the coconut oil and coconut free fatty acids were identified by Agilent gas chromatography (GC) as well as with a GC combined with mass spectrometry (GC/MS) to confirm the identification of the acids. A 30-m FFAP column (0.25 mm i.d., 0.25 µm df (Sigma-Aldrich Inc.)) was used. Helium was used as the carrier gas. For analyzing the relative ratios of all acids, an Agilent GC system (6890 N) equipped with an FID detector and SP-2380 column (30 m × 0.25 mm i.d.). Parameters for SP-2380 analysis were: column flow 1.0 ml/min with a helium head pressure of 136 kPa; split ratio 50:1; programmed ramp 120 to 135 °C at 20 °C/min, 135 to 265 °C at 7 °C/min, hold 5 min at 265 °C; injector and detector temperatures set at 250 °C. For structure confirmation using GC-MS, the same temperature program was used as those of GC system. Saturated C 8 − C 30 FAME provided standards used to make FAME assignments. The FID results were standardized for the individual fatty acids and reported as w/w%. Relative proportions of free fatty acids from the coconut fatty acids were determined by acid methanolysis. Coconut fatty acids samples for GC were prepared by heating a 10 mg sample of coconut oil in 0.5 ml of 0.5 M KOH/MeOH to reflux on a heating block for 60 min in a sealed vial. After cooling to room temperature, 2 ml of 1 M H 2 SO 4 /MeOH was added to the vial, and the vial was resealed and heated to reflux on a heating block for 15 min. The solution was cooled and transferred to a small separatory funnel with hexane (1 mL) and washed with water (2 mL), dried over sodium sulfate, gravity filtered, placed in a GC vial with hexanes, sealed, and injected onto the GC.

Repellent efficacy and longevity of coconut fatty acid formulation against biting flies on pasture cattle

A starch coconut fatty acid composite was prepared in a 4-L stainless steel Waring blender (Dynamics Corporation of America, New Hartford, CT, USA). A mixture of hot (80–90 °C) deionized water (1500 mL) and coconut fatty acids (152.0 g) was stirred to crudely emulsify the mixture. To the hot slurry, a mixture of waxy starch (200.5 g; moisture content 8.70%) and pectin (3.99 g; moisture content 12.09%) was added and stirred vigorously. The resulting slurry was delivered to the jet cooker utilizing a Moyno progressing cavity pump (Robbins Meyers, Springfield, OH, USA) at a flowrate of 1 l/min. The slurried mixture and steam were combined in a Penick and Ford hydroheater (Penford Corp, Cedar Rapids, IA, USA). Cooking temperature was 140 °C using steam supplied at 448 kPa, and the hydroheater backpressure set at 275 kPa. Approximately 2100 ml of a white opaque aqueous starch-coconut fatty acid mixture was collected and then cooled to room temperature while stirring (solids content ranged between 14–16%, as determined by freeze-drying accurately weighed amounts of the solution in duplicate). The solids content varied between experiments due to dilution of the cooked dispersion with variable amounts of condensed steam. The final composite had a solids content of 14.20% and was composed of 45.0% coconut fatty acids, 55.0% starch, and 85.80% water. The actual amount of coconut fatty acids contained in the formulation was 14.6% × 0.45 = 6.6 wt.%. The starch encapsulated coconut fatty acid composite was warmed in hot water bath and stirred well before the application. The aqueous starch-coconut fatty acid composite was stored at room temperature and subsequently brought to the field before being r topical applied on cattle.

The repellency against biting flies of the starch coconut fatty acid composite was tested on heifers under field conditions during the summer of 2017. The repellency tests were carried out in North Platte (University of Nebraska, West Central Research and Extension Center), NE, USA. Tests were conducted using criteria specified by the American Society for Testing and Materials (ASTM, 1980) and protocols approved by the Institutional Animal Care and Use Committee of the University of Nebraska (IACUC protocol no. 06–12–053 C). To test the effectiveness of the coconut fatty acid-based formulation, we used 12 heifers randomly assigned to two groups of six. Each cattle were an ear-tag number. Around 500 ml of the formulation was topically applied onto the whole-body surface of each cattle (including 4 legs as well). Since starch based formulation without coconut fatty acids added showed no repellency against biting flies, the control group cattle were not treated to reduce animal stress caused by treatment. Testing was done in two separate pastures of equivalent carrying capacity ranging in size from 10 to 17 hectares. Pastures were randomly assigned to each treatment that will enable precise estimates of treatment and treatment by period effects. Battery driven Fimco® sprayers were calibrated and used to make the application to the legs and belly of each animal. Cattle were individually restrained using a cattle chute during spray application and then released into the test pasture for exposure to ambient biting fly populations. The total number of biting flies on all four legs and belly of each cow were counted and expressed as the total number of stable flies per animal. Counts will be made between 1300 and 1600 during predetermined intervals by the same individual. The counts started from day 1 through day 4 (starting from Tuesday till Friday each week). These counts were confirmed using Microsoft Image Viewer from a window-based computer to examine photographs taken from a Nikon digital camera (D60) during the observations. Comparisons between treated and control animals for numbers of flies observed were performed using Student t-test. Results with P < 0.05 were considered statistically significant.

All methods related to biting fly repellency studies involved cattle from university of Nebraska North Platte research center were performed in accordance with the relevant guidelines and regulations approved by UNL animal committee.