Figure 1

Figure 1. Study design for experimental infection of reindeer with JCV. JCV, Jamestown Canyon virus; PBS, phosphate-buffered saline.

In May and December 2020, we screened 30 reindeer from the Large Animal Research Station (University of Alaska Fairbanks; UAF) for CSV antibodies by using cELISA (4,7) (Appendix Figure 1). After a summer of insect activity, seroprevalence remained the same among the adults (91%; n = 10/11), but 84% (n = 16/19) of young animals had seroconverted. We housed a control group (1 infected and 1 naive female) in a separate pen from experimentally exposed naive reindeer (2 females, 1 male) and a superinfected group (2 females, 1 male) (Figure 1). Exposed animals received 10⁶ PFU of JCV in 1 mL of sterile phosphate buffered saline (PBS), pH 7.2, administered subcutaneously in the right shoulder. The control group received 1 mL PBS without virus, administered identically.

We disinfected handling facilities daily with a 10% bleach solution and collected rectal temperatures, weights, and jugular blood samples daily. We manually differentiated and quantified leukocytes on EDTA blood smears stained with EASY III (Azer Scientific, https://www.azerscientific.com). We anesthetized animals with intramuscular ketamine (7–10 mg/kg) and xylazine (0.6 mg/kg) before we euthanized them with intravenous pentobarbital (60–80 mg/kg) (Euthasol; Virbac, https://us.virbac.com) (Figure 1). During necropsy, we collected cerebral spinal fluid in red top BD Vacutainer tubes (https://www.bd.com) and placed oral, nasal, and conjunctival swab samples in tubes containing 1.0 mL sterile PBS, stored at −80°C until shipping. We subdivided tissues (liver, kidney, spleen, lung, gonad, brain, spinal cord, quadriceps, heart, muscle under the injection site, tonsils, parotid lymph nodes, retropharyngeal and mesenteric lymph nodes) and stored them frozen (−20°C) and fixed in 10% neutral buffered formalin. We also collected heart blood, chest fluid, abdominal fluid, urine, and feces if present. We disinfected instruments between each animal. We transferred all samples to the Zoonotic Parasite Research Unit (Western College of Veterinary Medicine, Saskatoon, SK, Canada) and stored at −20°C until testing.

We extracted RNA from tissues and fluids by using the RNeasy Mini Kit (QIAGEN, https://www.qiagen.com) and performed reverse transcription PCR with JCV primers and probe (4). We considered samples at <30 quantification cycles (Cq) to be positive. Positive controls were gBlocks gene fragments (Integrated DNA Technologies, https://www.idtdna.com) created from the small (S) segment of JCV (GenBank accession no. MN135989.1).

Tissues were trimmed, embedded in paraffin, cut, and stained with hematoxylin and eosin after 30 days in formalin at Prairie Diagnostic Services Inc. (Saskatoon). Slides were read by a blinded, board-certified, veterinary pathologist. Paraffin-embedded sections of brain (cerebrum, cerebellum, and obex), cranial lymph node, tonsil, and a gastrointestinal section with myenteric plexus were sent to Brigham and Women’s Hospital and Harvard Medical School (Boston, MA, USA) for in situ hybridization (ISH) and immunohistochemistry (Appendix Table 1). RNA ISH was performed on a Leica BOND-III system (Leica Biosystems, https://www.leicabiosystems.com) with RNAScope probes V-JCV-LMS-O1 (1187658-C1) (Advanced Cell Diagnostics, https://www.acdbio.com) according to manufacturer protocols and 14 ZZ probes targeting JCV isolate 11497-03; segments tested were 52–814 bp of the S segment (GenBank accession no. EF681845.1), 2–462 bp of the medium (M) segment (accession no. EF687121.1), and 227–337 bp of the large (L) segment (accession no. EF687059.1). We performed immunohistochemistry by using the Leica-BOND-III system with rabbit polyclonal GFAP antibody (1:3,000 dilution; abcam, https://www.abcam.com) and rabbit polyclonal IBA1 antibody (1:100 dilution; FUJIFILM Wako Chemicals Corp., https://www.fujifilm.com). Slides were reviewed by a blinded, board-certified, medical neuropathologist.

Figure 2

Figure 2. Average antibody response for naive and superinfected groups of reindeer experimentally infected with Jamestown Canyon virus. The titer of the naive control animal is provided for comparison. SD (error bars)...

Throughout the experiment, we observed no changes in behavior or body temperature of the reindeer (Appendix Table 2). The naive exposed group seroconverted on postinfection day 5, and antibody titers for all exposed reindeer peaked at postinfection day 7 (Figure 2). Lymphocytes were consistently elevated for reindeer in the naive and superinfected exposed groups but not in the control group. Viral RNA was detected in blood on days 1 (Cq = 13.80; naive animal) and 5 (Cq = 10.08; superinfected animal). Viral RNA was detected in chest fluid (Cq = 11.47), urine (Cq = 20.26), spleen (Cq = 22.04), parotid lymph node (Cq = 16.26), obex (Cq = 28.03), and uterus (Cq = 11.66) from exposed but not control animals (Appendix Table 1).

Figure 3

Figure 3. Histopathology of samples from reindeer experimentally infected with JCV. Representative brain sections from naive control (RD985), naive infected (RD977), and superinfected (RD978) animals shows H&E-stained sections of cerebral cortex and...

Histopathology revealed subtle, nonspecific neuronal changes in the brains of all animals except the naive control. Mild congestion and lymphoplasmacytic inflammation were detected in kidneys (n = 8), urinary bladder (n = 1 naive control), liver (n = 4, including previously infected control), and conjunctiva (n = 1, previously infected control). In situ hybridization did not reveal viral RNA within lesions. Immunohistochemistry for GFAP (glial fibrillary acidic protein) showed mild to moderate astrogliosis and for IBA1 (allograft inflammatory factor 1) showed mild microgliosis in brain sections (Figure 3).

We demonstrate JCV experimental infection in reindeer with no clinical signs of illness and minimal, nonspecific histological changes, similar to what has been reported for other animal arbovirus infections (8). Our results are consistent with those of a previous experiment in which white-tailed deer were infected with JCV and were viremic for 2–4 days but showed no clinical illness (9). We detected RNA in blood in the chest cavity of 1 animal at postinfection day 11, suggesting that that reindeer may be viremic for longer. The high rate of seroconversion in the naturally exposed herd of 30 reindeer suggests that reindeer and caribou may play a role in the ecology of CSVs in northern ecosystems, especially given that most young animals seroconverted within a single summer (11%–84%). Further studies focusing on isolation of live virus from experimentally infected animals in addition to vector competence would provide more information about the potential for reindeer to be maintenance hosts of the virus (10).

In humans, neuroinvasive disease is a strong component of JCV disease (11). We note that genetic material was found in the obex of 1 reindeer, suggesting potential infection, but ISH and immunohistochemistry results did not demonstrate neuronal invasion, despite a potential glial response. Although JCV was detected in the uterus, the implications for potential vertical spread or other effects are unknown. Single-stranded RNA viruses are not stable in the environment, and multiple freeze–thaw events probably affected genetic detection of virus in our study (12).

Out detection of JCV in blood and tissues of experimentally exposed reindeer, together with high seroprevalence in free-ranging animals in North America, suggests that Rangifer spp. caribou and reindeer could play a role in disease ecology of JCV in Arctic and sub-Arctic North America (4). Caribou and reindeer have substantial cultural and subsistence value for Arctic peoples; our findings also highlight that those animals may play a role in emergence and dissemination of vectorborne zoonoses in Arctic regions.