Xylazine

One-month-old female Syrian hamsters (Japan SLC Inc.) and 7- to 8-mo-old female Syrian hamsters (Envigo) were used in this study. Baseline body weights were measured before infection. Under ketamine−xylazine anesthesia, four hamsters per group were inoculated with 10^5.6 PFU (in 110 μL) or with 10³ PFU (in 110 μL) of UT-NCGM02 via a combination of the intranasal (100 μL) and ocular (10 μL) routes. Body weight was monitored daily for 14 d.
For virological and pathological examinations, two, four, or five hamsters per group were infected with 10^5.6 PFU (in 110 μL) or with 10³ PFU (in 110 μL) of the virus via a combination of the intranasal and ocular routes; 3, 6, and 10 d postinfection, the animals were killed, and their organs (nasal turbinates, trachea, lungs, eyelids, brain, heart, liver, spleen, kidneys, jejunum, colon, and blood) were collected.
For the reinfection experiments, three hamsters per group were infected with 10^5.6 PFU (in 110 μL) or with 10³ PFU (in 110 μL) of UT-NCGM02 or PBS (mock) via a combination of the intranasal and ocular routes. On day 20 postinfection, these animals were reinfected with 10^5.6 PFU of the virus via a combination of the intranasal and ocular routes. On day 4 after reinfection, the animals were killed, and the virus titers in the nasal turbinates, trachea, and lungs were determined by means of plaque assays in VeroE6/TMPRSS2 cells.
For the passive transfer experiments, eight hamsters were infected with 10^5.6 PFU (in 110 μL) or with 10³ PFU (in 110 μL) of UT-NCGM02 via a combination of the intranasal and ocular routes. Serum samples were collected from these infected hamsters on day 38 or 39 postinfection, and were pooled. Control serum was obtained from uninfected age-matched hamsters. Three hamsters per group were inoculated intranasally with 10³ PFU of UT-NCGM02. On day 1 or 2 postinfection, hamsters were injected intraperitoneally with the postinfection serum or control serum (2 mL per hamster). The animals were killed on day 4 postinfection, and the virus titers in the nasal turbinates and lungs were determined by means of plaque assays in VeroE6/TMPRSS2 cells. All experiments with hamsters were performed in accordance with the Science Council of Japan’s Guidelines for Proper Conduct of Animal Experiments and the guidelines set by the Institutional Animal Care and Use Committee at the University of Wisconsin–Madison. The protocol was approved by the Animal Experiment Committee of the Institute of Medical Science, the University of Tokyo (approval no. PA19-75) and the Animal Care and Use Committee of the University of Wisconsin–Madison (protocol no. V00806).
Detailed materials and methods for this study are described in SI Appendix.

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.

At the end of experimental period, rats were placed in individual metabolic cages and 24 h urine was collected to measure albumin and creatinine concentration. Then, animals were deeply anesthetized with ketamine (50 mg/kg) and xylazine (5 mg/kg) and blood samples were collected via cardiac puncture for urea and creatinine measurement. Kidneys were immediately taken and washed with cold isotonic saline. The left kidney was fixed in 10% formalin for histopathological examinations and the right kidney stored at -80℃ for biochemical analysis.