Anesthetic Effect

The mice brains were collected in a separate suite at the same time of the day during their active cycle following four different anesthesia and euthanasia methods: (1) Ketamine/Xylazine: mice were decapitated 45 min following an intraperitoneal injection of 100 mg/kg (#VINB-KET0-7021, Henry Schein Animal Health, Dublin, OH, USA) with 10 mg/kg xylazine hydrochloride supplement (#X1251-1G, Sigma-Aldrich, St. Louis, MO, USA). Serum ketamine levels are highest at 10–20 min post-injection (Ganguly et al., 2018 (link)); therefore to reduce the impact of ketamine on MAPK activity we chose the 45 min time-point, also because that is around the time a perfused brain would be collected. The anesthetic effect of a mixture of 100/10 mg/kg ketamine/xylazine is known to last up to 80 min with reflex suppressions and produces stable heart rates 40 min post-injection in mice (Erhardt et al., 1984 (link); Xu et al., 2007 (link)); thus the mouse is still sedated at our chosen time point; (2) Isoflurane: mice were placed in a plexiglass chamber with 5% isoflurane, USP (#NDC 13985-046-60, VetOne, Boise, ID, USA) for 5 min, and decapitated when fully sedated, as measured by a lack of active paw reflex; (3) Carbon Dioxide Asphyxiation: mice were placed in a new cage with corn cob bedding, and immediately euthanized by displacement of air with 100% carbon dioxide, within 5 min and decapitated for tissue collection; and (4) Decapitation: mice were gently restrained and decapitated in a new cage to minimize the exposure to blood from conspecific mice.

Ticks were acquired from the Oklahoma State Tick Rearing Facility (OSU) (Stillwater, OK, USA). Equal numbers of each sex and species (I. scapularis and A. americanum) were obtained. For each lot of I. scapularis and A. americanum and prior to shipment to the study site, OSU screened a subsample of ticks (n = 10) for pathogens using standardized PCR assays. Ixodes scapularis were screened for B. burgdorferi and Anaplasma phagocytophilum. Amblyomma americanum were screened for the presence of Ehrlichia chaffeensis, Francisella tularensis and Rickettsia rickettsii. All PCR-screened ticks were negative for the above pathogens. Once ticks arrived at the study site, they were housed in an industry-standard desiccator with the relative humidity maintained at > 90% until enclosed in a feeding capsule for attachment to deer.
The feeding capsules utilized in this study were specifically designed for holding blood-feeding I. scapularis and A. americanum. Feeding capsules allow for the containment and localization of ticks and aid in facilitating blood-feeding [40 (link)]. The traditional stockinet sleeve method for feeding ticks on cattle [41 (link)–43 ] was determined to be inadequate for white-tailed deer. We instead developed a feeding capsule for deer application, which was in part based upon feeding capsules for ticks (referred to hereafter as tick feeding capsules) previously designed for tick-feeding on rabbits and sheep [44 ]. To make each capsule, sheets of ethylene–vinyl acetate foam were cut into three square pieces. Each square had a different outside area, allowing for flexibility (base, approx. 12 × 12 cm; middle, approx. 9 × 9 cm; top, approx. 7 × 7 cm), and had a combined depth of approximately 18 mm. The center of each square was cut away, creating an opening. The inner surface areas of the base and middle piece openings were each approximately 7 × 7 cm; the top piece had a smaller opening (approx. 1.5 × 1.5 cm) through which the ticks were to be inserted, which decreased the probability that ticks would escape through the top of the capsule (Additional file 3: Figure S2).
Deer were anesthetized using an intramuscular injection of telazol and xylazine at dosages of approximately 3 mg/kg and approximately 2.5 mg/kg, respectively. Once fully anesthetized, deer were weighed to the nearest 0.1 kg using a certified balance. Prior to blood collection and capsule attachment, large patches of fur on the neck were trimmed using electric horse clippers (Wahl®; Wahl Clipper Corp., Sterling, IL, USA). Prior to capsule attachment, 10 ml of blood was collected from the jugular vein of each deer using a 20-gauge needle. The blood from each individual deer was immediately placed into a vacutainer containing EDTA and was centrifuged for 10 min at 7000 revolutions/min. The plasma was transferred to 1.5-ml centrifuge tubes, which were then stored at − 20 °C until analysis.
Two identical tick feeding capsules were attached to opposing sides of the neck of each deer using a liberal amount of fabric glue (Tear Mender, St. Louis, MO, USA). Each capsule was held firmly in place for > 3 min to allow it to adhere to the skin and fur. For each deer, 20 I. scapularis mating pairs were placed within one capsule, and 20 A. americanum mating pairs were placed within the second capsule. Prior to tick attachment, 20 ticks (all same species and sex) were placed into a modified 5-ml syringe. Ticks were chilled in ice for approximately 5–10 min to slow movement. The 20 mating pairs were then carefully plunged into the capsules and a fine mesh lid was applied and reinforced with duct tape. Representative photos and video of the tick attachment process are presented in Fig. 2 and Additional file 4: Video S1, respectively. The capsules were further secured to deer by wrapping the neck with a veterinary bandage (3 M Company, St. Paul, MN, USA).Fig. 2

Tick capsule attachment and tick attachment. a Female ticks being plunged into capsule, b plunger being removed prior to mesh lid being secured, c completed, secured capsule being checked to ensure all corners are adhered to the neck, d closeup of completed capsule containing 20 Ixodes scapularis mating pairs

After completion of capsule and tick attachment, deer were given tolazine via intramuscular injection at a dose of 4 mg/kg to reverse the effects of the anesthetic. Deer were then housed in individual pens, observed closely until they were mobile and moving normally and monitored routinely for the remainder of the day.

The maxillary molars were extracted under anesthesia with triple anesthesia (medetomidine hydrochloride 0.75 mg/kg (Domitol, Nippon Zenyaku Kogyo Co.,Ltd. Fukushima, Japan), midazolam 4.0 mg/kg (Dormicum, Sandoz K. K., Tokyo, Japan), butorphanol 5.0 mg/kg (Vetorphale, Meiji Seika Pharma Co., Ltd. Tokyo, Japan)) using hooked-end forceps. After surgery, mice were injected with Atipamezole 0.75 mg/kg (antisedan, Nippon Zenyaku Kogyo Co.,Ltd.) to antagonize the anesthesia effect. Two weeks later, the tissue from the extraction socket was collected and used for analysis.

Prior to the behavior test, mice were adapted to the behavior recording environment by placing the mouse in the behavior testing arena with the power amplifier turned on. During the behavior recording, mice were lightly anesthetized with isoflurane (1% induction and maintenance). The base plate on the mouse and the wearable ultrasound transducer were both sufficiently filled with degassed ultrasound gel (Aquasonics). The wearable transducer was then securely plugged into the base plate of the mouse, and the mouse was then placed in a circular arena on a heating pad for 30 min to allow the mouse body temperature to recover from any possible anesthesia effects. The heating pad was then removed, and the mouse was allowed to habituate for 15 min in the actual behavior test arena.
During the recording period, FUS was applied at a frequency of 1.5 MHz, duty cycle (DC) of 40%, pulse repetition frequency (PRF) of 10 Hz, and 15 s total sonication duration with 185 s inter-stimulation interval (ISI) for a total of five stimulations. The onset and offset of the ultrasound pulse was smoothed to avoid possible auditory effects [29 (link)]. The acoustic pressures used in the study were 0, 0.7, and 1.1 MPa (measured in a water tank) to investigate the effect of pressure on locomotor behavior outcomes. These parameters corresponded to mechanical indices of 0, 0.57, and 0.90 (in water), respectively. Custom MATLAB software was used to control when ultrasound was applied via an Arduino Uno. A red LED attached to the Arduino Uno would turn on when ultrasound was applied to precisely synchronize mouse behavior to each FUS stimulation. In each group, mice were given five consecutive FUS stimulations at one pressure.