SARS-CoV-2, a novel coronavirus with high nucleotide identity to SARS-CoV and SARS-related coronaviruses detected in horseshoe bats, has spread across the world and impacted global healthcare systems and economy 1,2 . A suitable small animal model is needed to support vaccine and therapy development. We report the pathogenesis and transmissibility of the SARS-CoV-2 in golden Syrian hamsters. Immunohistochemistry demonstrated viral antigens in nasal mucosa, bronchial epithelial cells, and in areas of lung consolidation on days 2 and 5 post-inoculation (dpi), followed by rapid viral clearance and pneumocyte hyperplasia on 7 dpi. Viral antigen was also found in the duodenum epithelial cells with viral RNA detected in feces. Notably, SARS-CoV-2 transmitted efficiently from inoculated hamsters to naïve hamsters by direct contact and via aerosols. Transmission via fomites in soiled cages was less efficient. Although viral RNA was continuously detected in the nasal washes of inoculated hamsters for 14 days, the communicable period was short and correlated with the detection of infectious virus but not viral RNA. Inoculated and naturally-infected hamsters showed apparent weight loss, and all animals recovered with the detection of neutralizing antibodies. Our results suggest that SARS-CoV-2 infection in golden Syrian hamsters resemble features found in humans with mild infections. SARS-CoV-2 was first detected from a cluster of pneumonia patients in Wuhan, Hubei Province, China in December 2019. Although 55% of the initial cases were linked to one seafood wholesale market where wild animals were also sold 3 , multiple viral (sustained human-to-human transmissibility by symptomatic and pre-symptomatic patients 4 ) and ecological factors (extensive domestic and international travel during Chinese Lunar New Year) have contributed to the rapid global spread of the virus. The clinical spectrum of patients with the novel coronavirus disease (COVID-19) is wide, 19% of 72,314 symptomatic patients in China progressed to severe and critical illness 5 with an estimated 1.4% symptomatic case fatality risk 6 . There is no approved vaccine or treatment against SARS-CoV-2, and the available interventions including country lock-down and social distancing have severely disrupted the global supply chain and economy. A suitable animal model is essential for understanding the pathogenesis of this disease and for evaluating vaccine and therapeutic candidates. Previous animal studies on SARS-CoV suggested the importance of the interaction between the viral spike protein and the host angiotensin converting enzyme 2 (ACE2) receptors 7–10 as well as age and innate immune status of the animals 11–14 in pathogenesis. As with SARS-CoV, the spike protein of SARS-CoV-2 also utilizes ACE2 receptors that are distributed predominantly in the epithelial cells of the lungs and small intestine to gain entry into epithelial cells for viral replication 1,15 . SARS-CoV-2 showed good binding for human ACE2 but limited binding to murine ACE21, which has limited the use of inbred mice for research. Macaques and transgenic ICR mice expressing human ACE2 receptor were shown to be susceptible for SARS-CoV-2 infection 16–18 ; however, there is limited availability of these animal models. Cynomolgus macaques and rhesus macaques challenged with SARS-CoV-2 showed pneumonia with limited 17 and moderate 18 clinical signs, respectively. The challenged transgenic mice showed pneumonia moderate weight loss, and no apparent histological changes in non-respiratory tissues 16 . Previously generated transgenic mice expressing human ACE2 receptor have been reported to support SARS-CoV replication in the airway epithelial cells but were associated with neurological-related mortality due to high ACE2 expression in the brain 7–10 . Golden Syrian hamster is a widely used experimental animal model and was reported to support replication of SARS-CoV 19,20 but not MERS-CoV 21 , which utilizes the dipeptidyl peptidase-4 (DPP4) protein as the main receptor for viral entry. Previous study of SARS-CoV (Urbani strain) in 5-weeks-old golden Syrian hamsters showed robust viral replication with peak viral titers detected in the lungs on 2 dpi, followed by rapid viral clearance by 7 dpi, but without weight loss or evidence of disease in the inoculated animals 20 . A follow up study reported testing different strains of SARS-CoV in golden Syrian hamsters and found differences in virulence between SARS-CoV strains; lethality was reported in hamsters challenged with the Frk-1 strain, which differed from the non-lethal Urbani strain by the L1148F mutation in the S2 domain 19 . Hamsters are permissive for infection by other respiratory viruses including human metapneumovirus 22 , human parainfluenza virus 3 23 and influenza A virus and may support influenza transmission by contact or airborne routes 24,25 . Alignment of the ACE2 protein of human, macaque, mice, and hamster suggest that the spike protein of SARS-CoV-2 may interact more efficiently with hamster ACE2 than murine ACE2 (Extended Data Fig. 1). Here, we evaluated the pathogenesis and contact transmissibility of SARS-CoV-2 in 4–5 weeks old male golden Syrian hamsters. Hamsters were infected intranasally with 8 × 104 TCID50 of the BetaCoV/Hong Kong/VM20001061/2020 virus (GISAID# EPI_ISL_412028) isolated in Vero E6 cells from the nasopharynx aspirate and throat swab of a confirmed COVID-19 patient in Hong Kong. On 2, 5, 7 dpi, nasal turbinate, brain, lungs, heart, duodenum, liver, spleen and kidney were collected to monitor viral replication and histopathological changes. Peak viral load in the lungs was detected on 2 dpi and decreased on 5 dpi; no infectious virus was detected on 7 dpi despite of the continued detection of high copies of viral RNA (Fig. 1a). Infectious viral load was significantly different between 2 and 7 dpi (P= 0.019, Dunn’s multiple comparisons test) but not the RNA copy number (P= 0.076). No infectious virus was detected in the kidney although low copies of viral RNA were detected on 2 and 5 dpi (Fig. 1b). Histopathological examination detected an increase in inflammatory cells and consolidation in 5–10% of the lungs on 2 dpi (Fig. 1c, 1d) and 15–35% of the lungs on 5 dpi (Fig. 1e, 1f). Mononuclear cell infiltrate was observed in areas where viral antigen was detected on 2 and 5 dpi. Immunohistochemistry for SARS-CoV-2 N protein demonstrated viral antigen in the bronchial epithelial cells on 2 dpi (Fig. 1d) with progression to pneumocytes on 5 dpi (Fig. 1f). On 7 dpi, there was an increased consolidation in 30–60% of the lungs (Fig. 1g); however, no viral antigen was detected (Fig. 1h) and type 2 pneumocyte hyperplasia was prominent (Extended Data Fig. 2a). CD3 positive T lymphocytes were detected in the peri-bronchial region on 5 dpi, which may facilitate the rapid clearance of the infected cells (Extended Data Fig. 2b). There was moderate inflammatory cell infiltration in the nasal turbinate (Fig. 1i), and viral antigen was detected in the nasal epithelial cells (Fig. 1j) and in olfactory sensory neurons at the nasal mucosa (Fig. 1j). Infection in the olfactory neurons was further confirmed in cells expressing both SARS-CoV-N protein and neuron-specific beta-III tubulin (Extended Data Fig. 2c). Compared to mock infection (Extended data Fig. 2d and 2e), infection lead to a reduction in the number of olfactory neurons at the nasal mucosal on 2 dpi (Extended Data Fig. 2f), prominent nasal epithelial attenuation on 7 dpi (Extended Data Figure 2g), followed by tissue repairing on 14 dpi (Extended data Figure 2h). Though no inflammation was present (Fig. 1k), viral antigen was detected from the epithelial cells of duodenum on 2 dpi (Fig. 1l). This resembles the detection of SARS-CoV virus replication in the epithelial cells of terminal ileum and colon of SARS-CoV patients without observing apparent architectural disruption and inflammatory infiltrate 26 . No apparent histopathological change was observed from brain, heart, liver, and kidney on 5 dpi (Extended Data Fig. 2i, 2j, 2k, 2l). To assess the transmission potential of the SARS-CoV-2 in hamsters, three donor hamsters were inoculated intra-nasally with 8 × 104 TCID50 of the virus. At 24h post-inoculation, each donor was transferred to a new cage and co-housed with one naïve hamster. Weight changes and clinical signs were monitored daily and nasal washes were collected every other day from donors and contacts for 14 days. In donors, the peak infectious viral load in nasal washes was detected early post-inoculation followed by a rapid decline, although viral RNA was continuously detected for 14 days (Fig. 2a). Hamsters inoculated with the SARS-CoV-2 showed the maximal mean weight loss (mean ± SD, −11.97 ± 4.51%, N=6) on 6 dpi (Fig. 2b). Transmission from donors to co-housed contacts was efficient, and SARS-CoV-2 was detected from the co-housed hamsters on day 1 post-contact (dpc), with the peak viral load in nasal washes detected on 3 dpc (Fig. 2c). The total viral load shed in the nasal washes was approximated by calculating the area under the curve (AUC) for each animal. The contact hamsters shed comparable amount of virus in the nasal washes compared to the donor hamsters (P= 0.1, two-tailed Mann-Whitney test). Contact hamsters showed the maximal mean weight loss (mean ± SD, −10.68 ± 3.42%, N=3) on 6 dpc; all animals returned to the original weight after 11 dpc (Fig. 2d). Neutralizing antibody were detected using plaque reduction neutralization (PRNT) assay from donors on 14 dpi (titers at 1:640 for all) and from contacts on 13 dpc (titers at 1:160, 1:320, and 1:160). As viral RNA was continuously detected from the donor’s nasal washes for 14 days while infectious virus titers decreased rapidly, we repeated the experiment and co-housed naïve contacts with donors on 6 dpi. Low quantity of viral RNA was detected in the nasal washes in one contact on 3 and 7 dpc without detection of infectious virus in the nasal washes (Fig. 2e), and none of the contact hamsters showed weight loss (Fig. 2f). PRNT assay detected no neutralizing antibody ( 90% (PRNT90) reduction in the number of plaques. Histopathology and immunohistochemistry. Tissue (hearts, livers, spleens, duodenums, brains, right lungs and kidneys) were fixed in 4% paraformaldehyde and were processed for paraffin embedding. The 4-μm sections were stained with hematoxylin and eosin for histopathological examinations. For immunohistochemistry, SARS-CoV-2 N protein was detected using monoclonal antibody (4D11) 34 , CD3 was detected using polyclonal rabbit anti-human CD3 antibodies (DAKO), and the neuron-specific beta-III tubulin was detected using monoclonal antibody clone TuJ1 (R&D Systems). Images were captured using a Leica DFC 5400 digital camera and were processed using Leica Application Suite v4.13. Statistics and reproducibility. Kruskal-Wallis test and Dunn’s multiple comparisons test were used to compare viral loads in the lungs and kidney on 2, 5, 7 dpi. Area under the curve was calculated from the nasal washes of the donor and contact hamsters followed by Mann-Whiteny test. Data were analyzed in Microsoft Excel for Mac, version 16.35 and GraphPad Prism version 8.4.1. For the detection viral replication in hamsters, 9 hamsters were inoculated and tissues were collected from animals on 2 (N=3), 5 (N=3), 7 (N=3) dpi; the results from the three animals were similar (Fig. 1a and 1b). Inoculation of the donor hamsters was independently performed twice and the inoculated hamsters showed comparable weight loss and shed comparable amount of virus in the nasal washes (Fig. 2a, 2b, 3a, 3b). Transmission by direct contact, via aerosols or fomites were performed with three pairs of donor: contacts at 1:1 ratio. Extended Data Extended Data Figure 1. Sequence alignment of ACE2 proteins (1–420) from human, macaca, hamster, and mouse. Amino acid residues of human ACE2 that are experientially shown to interact with the receptor binding domain (RBD) of SARS-CoV-2 35 are denoted by *. Amino acid residues that are important for the interaction between human ACE2 and RBD of SARS-CoV are highlighted in red boxes 36 . Extended Data Figure 2. Haemotoxylin and eosin (H&E) staining and immunohistochemistry on SARS-CoV-2 challenged hamster tissues. a, Hyperplasia of the pneumocytes detected on 7 dpi. b, Detection of CD3 positive cells (using rabbit anti-human CD3 polyclonal antibody) in the lungs on 5 dpi. c, Detection of SARS-CoV-2 N protein (red staining, using monoclonal antibody 4D11) and olfactory neurons (brown staining, using monoclonal antibody TuJ1) from the nasal turbinate on 5 dpi. d, Detection of olfactory neurons (using monoclonal antibody TuJ1) from the nasal turbinate of a mock infected hamster (N=1). e, Nasal epithelial cells from the nasal turbinate of a mock infected hamster (N=1) showed negative staining for TuJ1. f, Detection of olfactory neurons from nasal turbinate on 2 dpi. g, Detection of olfactory neurons from nasal turbinate on 7 dpi. h. Detection of olfactory neurons from nasal turbinate on 14 dpi. i, H&E staining of the brain tissue on 5 dpi. j, H&E staining of the heart on 5 dpi. k, H&E staining of the liver on 5 dpi. l, H&E staining of the kidney on 5 dpi. Hamsters were intra-nasally inoculated with PBS (mock infection, N=1) or with 8 × 104 TCID50 of SARS-CoV-2 (N=9) and the tissues were collected on 2 (N=3), 5 (N=3), 7 (N=3) dpi. H&E and immunohistochemistry with tissues from three animals showed similar results and the representative results were shown. Extended Data Figure 3. Experimental layout for the aerosol transmission experiment in hamsters. To evaluate SARS-CoV-2 transmissibility via aerosols, one naïve hamster was exposed to one inoculated donor hamster in two adjacent stainless steel wired cages on 1 dpi for 8 hours. DietGel®76A (ClearH2O®) was provided to the hamsters during the 8-hour exposure. Exposure was done by holding the animals inside individually ventilated cages (IsoCage N, Techniplast) with 70 air changes per hour. Experiments were repeated with three pairs of donors: aerosol contact at 1:1 ratio. After exposure, the animals were single-housed in separate cages and were continued monitored for 14 days. Extended Data Table 1. Detection of SARS-CoV-2 in the soiled cages.To evaluate transmission potential of SARS-CoV-2 virus via fomites, three naïve fomite contact hamsters were each introduced to a soiled donor cage on 2 dpi. The fomite contact hamsters were single-housed for 48 hours inside the soiled cages and then were each transferred to a new cage on 4 dpi of the donors. The soiled cages were left empty at room temperature and were sampled again on 6 dpi of the donor. Surface samples and corn cob bedding were collected from the soiled cages on different time points to monitor infectious viral load and viral RNA copy numbers in the samples. Days post-inoculation Animal cage info Sampled area Material log10 TCID50/ mL log10 RNA copies/ mL Day 2 donor cage A 1.79 6.70 donor cage B bedding corn cobs < 5.18 donor cage C < 5.79 Day 4 fomite contact cage A cage side (in directcontact with theanimals) < 6.89 fomite contact cage B plastic < 5.21 fomite contact cage C 1.79 6.33 fomite contact cage A < 3.76 fomite contact cage B cage lid plastic < 4.33 fomite contact cage C < 4.10 fomite contact cage A < 5.26 fomite contact cage B pre-filter paper-based < 5.27 fomite contact cage C < 5.31 fomite contact cage A < 3.64 fomite contact cage B water bottle nozzle stainless steel < 4.20 fomite contact cage C 2.21 6.06 fomite contact cage A < 4.84 fomite contact cage B bedding corn cobs < 5.27 fomite contact cage C < 6.06 Day 6 fomite contact cage A cage side (in directcontact with theanimals) < 5.70 fomite contact cage B plastic < 5.61 fomite contact cage C < 6.51 fomite contact cage A < 4.75 fomite contact cage B cage lid plastic < 3.46 fomite contact cage C < 4.24 fomite contact cage A < 5.48 fomite contact cage B pre-filter paper-based < 5.23 fomite contact cage C < 5.36 fomite contact cage A < 5.12 fomite contact cage B bedding corn cobs < 6.24 fomite contact cage C < 5.58 Supplementary Material 1
