SARS-CoV-2, the causative agent of the COVID-19 pandemic, presents a broad host range. Domestic cats and white-tailed deer (WTD) are particularly susceptible to SARS-CoV-2 with multiple variant strains being associated with infections in these species. The virus replicates in the upper respiratory tract and in associated lymphoid tissues, and it is shed through oral and nasal secretions, which leads to efficient transmission of the virus to contact animals. Robust cell-mediated and humoral immune responses are induced upon infection in domestic cats, which curb the progression of clinical disease and are associated with control of infection. In WTD, high levels of neutralizing Abs are detected early upon infection. In this review, the current understanding of the infection dynamics, pathogenesis, and immune responses to SARS-CoV-2 infection in animals, with special focus on naturally susceptible felids and WTD, are discussed.
Coronaviruses are a diverse group of viruses that infect different animal species and have long been recognized for their pandemic potential. Two highly pathogenic coronaviruses of zoonotic origin, SARS-CoV and Middle East respiratory syndrome (MERS)-CoV, causing fatal respiratory illness in humans emerged in 2002 and 2012, respectively, making coronaviruses a major public health threat (1). The most recent and notable coronavirus threat that emerged in humans is SARS-CoV-2, the causative agent of the COVID-19 pandemic, which was first detected in humans in Wuhan, China in December 2019 (2). Similar to SARS and MERS, COVID-19 patients also showed symptoms of viral pneumonia, including fever, cough, and chest discomfort, and in severe cases dyspnea and bilateral lung infiltration and death (2).
All previously described human coronaviruses (human CoV [HCoV]-HKU1, HCoV-229E, HCoV-NL63, HCoV-OC43, MERS-CoV, and SARS-CoV) have zoonotic origins resulting from spillover events associated with intermediate animal hosts (3). Epidemiological studies suggest that the Huanan market in Wuhan city was a major epicenter of SARS-CoV-2 infection, as 27 (66%) of the first 41 human cases of COVID-19 were linked to the market (4), which reportedly sold seafood and live animals for human consumption, including poultry, bats, marmots, cervids, raccoon dogs, and several other wild species (5). Viruses closely related to SARS-CoV-2 have been detected in bats and pangolins in multiple localities in South-East Asia, including in China, Thailand, Cambodia, and Japan (6, 7). A bat virus, named RaTG13, detected in Rhinolophus affinis in Yunnan, China, presenting 96.2% nucleotide similarity at the whole genome level with SARS-CoV-2, represents the closest relative to SARS-CoV-2 identified to date (6, 8). Other bat viruses such as RmYN02 from a Rhinolophus malayanus bat sampled in Yunnan, China (9), and bat coronaviruses ZC45 and ZXC21 previously detected in Rhinolophus pusillus bats in eastern China (10) also cluster with the SARS-CoV-2 lineage of the subgenus Sarbecovirus, suggesting that bats could be a reservoir for SARS-CoV-2–like viruses. Recent evidence suggests that raccoon dogs present at the Huanan market at the time the pandemic started may have been exposed to SARS-CoV-2 (11); however, whether this or any other animal species commercialized in the market served as an intermediate host before the virus jumped into humans remains unknown.
SARS-CoV-2 is a single-stranded, positive-sense RNA virus, belonging to the Coronaviridae family, Orthocoronavirinae subfamily, Betacoronavirus genus, and Sarbecovirus subgenus. The virus genome is ∼30 kb in length, and it shares ∼80% sequence homology with SARS-CoV. The genome contains 14 open reading frames (ORFs) that encode 29 viral proteins. About two thirds of the viral genome encode two overlapping polyproteins, pp1a and pp1ab (12). These polyproteins are cleaved by two viral proteases (NSP3 and NSP5) into 16 nonstructural proteins, which are essential for viral replication and transcription. The 3′ end of the genome encodes ORFs for four structural proteins, including the spike (S), envelope, membrane, and nucleocapsid (N) protein, which participate in virion assembly and entry and/or suppress host immune responses. Several accessory proteins such as ORF3a, ORF3b, ORF6, ORF7a, ORF7b, ORF8, ORF9b, ORF9c, and ORF10 are encoded at the 3′ end of the genome as individual ORFs and contribute to viral pathogenesis and host immune evasion (13, 14).
The S protein functions in viral entry and mediates receptor binding and fusion. Receptor binding and membrane fusion are key steps for coronaviruses to achieve cross-species infection and establish efficient transmission pathways in new host species. The SARS-CoV-2 S protein is one of the main determinants of cellular susceptibility, as it mediates virus entry and cell fusion through its direct interactions with the cellular receptor (angiotensin-converting enzyme 2 [ACE2]) as well as host cellular proteases (furin and transmembrane serine protease 2 [TMPRSS2]) (15). Thus, at the cellular level susceptibility to SARS-CoV-2 is dependent minimally on the expression of a compatible cellular ACE2 receptor that allows efficient binding of the virus S protein and triggers subsequent cleavage of S1/S2 and S2′ sites by ubiquitous furin-like proteases and TMPRSS2, respectively, enabling fusion of the virus envelope and host cell membranes.
At the host level, SARS-CoV-2 susceptibility is also affected by efficient ACE2 receptor recognition. However, receptor binding is only the first step in virus infection. Host factors that enable virus replication in the cytoplasm making cells permissive to infection also play a critical role. Additionally, the expression levels of key entry factors (ACE2 and TMPRSS2) in susceptible/permissive cells and the localization and accessibility of these cells in target tissues are crucial for virus infection. Finally, the abilities of the virus to bypass intrinsic host restriction factors elicited to control the infection locally in target cells and tissues (16) or systemically are also very important determinants of susceptibility and host range to SARS-CoV-2.
Natural SARS-CoV-2 infections in animals
Natural SARS-CoV-2 infections have been reported in many species, including minks, ferrets, lions, tigers, cats, and white-tailed deer (WTD) (17). As of April 26, 2023, a total of 754 cases of SARS-CoV-2 infection have been recorded globally involving 32 animal species from 39 countries (18), with most SARS-CoV-2 variants being detected in animals. While most animal infections are subclinical, case fatality rates approaching ∼35–55% have been reported in mink (19).
Cats are highly susceptible to SARS-CoV-2 infection (17, 20–22). Most SARS-CoV-2 infections in cats involve natural human-to-cat transmission of the virus (23–32). Several variants including Alpha (B1.1.7), Delta (AY.3), and Omicron (B1.1.529) have been detected in cats (20, 29, 33–36). Experimental SARS-CoV-2 infections in cats confirmed efficient virus replication in the respiratory tract, prolonged virus shedding in respiratory and oral secretions, and development of lesions characteristic of SARS-CoV-2 infection in humans (22, 37–40). Additionally, the potential for cat-to-cat transmission of SARS-CoV-2 has been demonstrated (37–39, 41, 42).
Natural SARS-CoV-2 infections have also been reported in other felids including tigers and lions, which, similar to infections in domestic cats, have been linked to human-to-animal transmission events (18, 43–48). Affected animals may present subclinical infections or present with lethargy, anorexia, and mild respiratory signs (45–48). The first reported infected Malayan tiger presented necrosis in the trachea with viral RNA being detected in necrotic cells in tracheal wash fluid by in situ hybridization (48). Other felids such as cougars and snow leopards are also naturally susceptible to SARS-CoV-2 infection (18, 43).
The susceptibility of minks to SARS-CoV-2 was first reported during an outbreak affecting farmed mink in April 2020 in the Netherlands (49). The affected minks presented mild to severe respiratory distress with several animals dying of interstitial pneumonia. Importantly, during this outbreak mink-to-mink transmission was documented (49). After the initial outbreak in the Netherlands, several outbreaks were reported in mink farms globally including in Denmark (50), Poland (51), Spain (52), and the United States (53). Notably, infection and transmission of SARS-CoV-2 in mink resulted in adaptive mutations in the virus genome, and viruses with these mink-origin mutations were detected in humans, suggesting mink-to-human transmission of the virus (54, 55). These findings raised concerns about the potential establishment of a new reservoir of SARS-CoV-2 in mink.
One of the most remarkable findings regarding the SARS-CoV-2 host range was that WTD (Odocoileus virginianus) were shown to be highly susceptible and capable of transmitting SARS-CoV-2 to noninfected contact animals (56–58). Most importantly, multiple infections in free-ranging WTD have been reported in the United States and Canada. Genomic surveillance and sequence analysis of SARS-CoV-2 strains recovered from WTD indicate efficient deer-to-deer transmission of the virus (59–61), with recent evidence suggesting rare but possible spillback to humans (62).
SARS-CoV-2 pathogenesis and immune responses in cats
Although most of the natural outbreaks and experimental infection of SARS-CoV-2 are subclinical in cats, productive virus replication and pathology are observed mainly in the respiratory tract of affected animals. Infected cats transmit the virus to contact animals, and both primary and contact-infected cats seroconvert and resist reinfection with the virus. The infection dynamics and immune responses to SARS-CoV-2 infection in cats are reviewed based on several experimental studies and summarized in Fig. 1 (21, 22, 37, 40–42, 63, 64).
Clinical outcome of SARS-CoV-2 infection in cats
The clinical outcome of SARS-CoV-2 infection in domestic cats has been variable, depending on the virus strain, dose and route of inoculation, and age of the animals. Most experimental studies using the ancestral SARS-CoV-2 strains and natural routes of infection (intranasal and/or oral) have reported subclinical infection without notable changes in body temperature or body weight (21, 37, 39, 41, 42). No significant changes in hematological and serum biochemical parameters have been reported (37). In contrast, intratracheal inoculation of SARS-CoV-2 in cats has been shown to result in marked clinical disease with animals presenting lethargy, increased respiratory effort, fever, and weight loss (64). A comparative pathogenicity study with SARS-CoV-2 variants showed elevated body temperature, loss of body weight, lethargy, and respiratory signs for 1–3 d postinfection (dpi) in cats infected with the ancestral B.1 D614G and Delta (B.1.617.2) variants and subclinical infection in Omega BA1.1-infected cats (22).
Infection dynamics of SARS-CoV-2 in cats
Following replication of SARS-CoV-2 in the respiratory tract, viral RNA and infectious virus have been consistently detected in respiratory tissues (nasal turbinate, trachea, and lungs) and associated lymphoid organs including tonsil and retropharyngeal lymph nodes between 3 and 5 dpi (21, 22, 37, 40–42), with the highest viral loads detected at 3–4 dpi (21, 22, 37). Viral RNA is detected in the lungs between days 3 and 14 dpi with infectious virus being recovered between days 3 and 5 postinfection. Infected animals usually clear the infection from both upper and lower respiratory tissues by 10 dpi (22, 40). Limited viral RNA, but no infectious virus, has been detected in the upper respiratory tract (URT) and lymphoid organs up to 21 dpi (37, 42). Viral RNA has been detected in bronchoalveolar lavage fluid between 3 and 14 dpi with the highest viral loads detected at 3–5 dpi (22, 37, 42); however, infectious virus has only been detected between 3 and 5 dpi (22).
In addition to the respiratory tract, viral RNA has been detected in the heart, mediastinal lymph node, liver, spleen, kidney, small intestine, and mesenteric lymph node at 3–21 dpi (21, 22, 37, 41, 42). However, infectious virus was not detected in nonrespiratory tissues (21, 22, 40, 41). Interestingly, viral RNA or infectious virus is usually not detected in the blood or serum of infected cats (37), which suggests alternative routes of virus dissemination from the initial sites of virus replication in the respiratory tract to other organs and systems.
Infected cats shed SARS-CoV-2 in nasal and oral secretions, with low levels of viral RNA also being transiently detected in feces. Viral RNA is detected in nasal secretions between 1 and 14 dpi, with peak viral RNA loads observed at 3–5 dpi (22, 37, 42). Importantly, a high level of infectious virus shedding in nasal secretions is detected up to 7 dpi with sporadic detection at 10 and 14 dpi (22, 41). A similar kinetics of virus shedding is also observed in oral secretions; however, the virus titers recovered are ∼1 log lower than those recovered from nasal secretions (22, 37, 41, 42). Shedding of viral RNA in feces is markedly lower than in respiratory and oral secretions (22, 37, 42).
Differences in the infection and virus shedding dynamics have been observed in cats inoculated with different SARS-CoV-2 variants (22). A study comparing the pathogenicity of three SARS-CoV-2 variants (the ancestral B.1 D614G, Delta [B1. 617.2], and Omicron [BA1.1]) reported significantly lower virus shedding in nasal, oral secretions, and feces in Omicron-inoculated cats than in cats inoculated with B.1 D614G and Delta viruses (22). Similarly, lower virus titers were detected in tissues of Omicron-inoculated cats than in B.1 D614G– and Delta-inoculated cats. Whereas D614G and Delta viruses present broad tissue tropism and abundant replication in nasal turbinate, palate/tonsil, trachea, retropharyngeal lymph node, and lungs at 3 and 5 dpi, the Omicron replication was mainly confined to the URT (nasal turbinate and trachea) with much lower viral loads detected in lungs (22).
Consistent with robust SARS-CoV-2 replication in the URT and shedding in respiratory secretions, the virus is efficiently transmitted between infected and contact cats (37, 41, 42). Current studies demonstrate efficient transmission from infected to contact animals as early as 1–2 d postcontact (37). Contact cats shed infectious virus through oral and nasal secretions by 1–2 d postcontact, with peak shedding occurring at day 7 postcontact (37).
Pathological changes following SARS-CoV-2 infection in cats
SARS-CoV-2 infection in cats produces pulmonary lesions that resemble lesions observed in humans with acute respiratory distress syndrome from COVID-19 (64).
Upon intranasal inoculation, the lungs of cats infected with the SARS-CoV-2 strain USA-WA1/2020 present various degrees of edema, discoloration, congestion, and atelectasis (37). Notably, intratracheal inoculation of cats with SARS-CoV-2 strain WA1/2020 results in a more severe outcome, and inoculated animals present marked pathology in lungs characterized by large multifocal to coalescing regions of dark red consolidation with edema at 4 dpi that become more pronounced at 8 dpi (64). Similarly, intratracheal inoculation of cats with the Delta variant results in severe consolidation, hemorrhages, and edema in lungs (63).
Histological changes are usually limited to URT and bronchial tree in lungs of infected cats. In the nasal passage, ulcerative neutrophilic rhinitis is accompanied by neutrophil infiltration of the lamina propria and accumulation of fibrin aggregates, cellular debris, degenerated epithelial cells, and leukocytes (22, 41, 42). Epithelial cells of the nasal mucosa are the main target of SARS-CoV-2 replication, with extensive viral RNA labeling observed by in situ hybridization at 3–5 dpi (22). SARS-CoV-2 nucleoprotein (NP) Ag and viral RNA are also detected within squamous (in rostral turbinate) and respiratory epithelial cells (in intermediate and deep turbinate) (42). Lesions in the trachea are characterized by epithelial degeneration, necrosis, and regeneration associated with a mixed inflammatory infiltrate (22). Multifocal lymphocytic and neutrophilic tracheobronchoadenitis of seromucous glands of the lamina propria and submucosa of the trachea and bronchi have been reported at 4 dpi, which progress to mild to moderate at 7 dpi (37). Viral staining is detected in cells within submucosal interstitial stroma and submucosal glands (22, 37, 42). Affected submucosal glands and associated ducts are variably distended, lined by attenuated epithelium, and contain necrotic cell debris and infiltrating lymphocytes, macrophages, and plasma cells (37).
In lungs, mild bronchitis with lymphoid hyperplasia, moderate to severe histiocytic bronchiolitis with partial to complete occlusion of lumina, and moderate to severe thickening of alveolar septa are reported at 3 dpi (22, 40). The interstitial inflammation seems to resolve quickly, whereas alveolar septal thickening persists up to 10 dpi (40). The BALT becomes hyperplastic, containing a mixture of CD3+ and CD20+ lymphocytes (65). Persistent lung lesions are observed as late as 28 dpi and include histiocytic bronchiolitis with luminal plugs and thickened alveolar septa coupled with perivascular fibrosis and vascular proliferation within the thickened interstitium (40). The specific cell types that contribute to the interstitial thickening include CD3+ T lymphocytes, CD20+ B lymphocytes, CD204+ macrophages, and fewer plasma cells (65). The perivascular lymphoid aggregates are also MHC class II positive. Additionally, abundant type II pneumocytes are scattered within the alveolar septa. The bronchiolar plugs consist of round to epithelioid cells with foamy to slightly fibrillar eosinophilic cytoplasm consistent with viable and degenerative Iba-1+ and CD204+ macrophages (65). At later time points (28 dpi), increased numbers of bronchioles show variable remodeling with occasional erosion, robust epithelial proliferation, and segmental bronchiolar constriction (65). Hypertrophy of smooth muscle of terminal bronchioles and bronchioles is also noticed at 28 dpi (65). Staining of viral RNA in the lung is more frequent in the interstitial regions, especially in cells of the bronchiolar glands at early time points (3–5 dpi) (22). No viral RNA or Ags are detected within lining epithelial cells or elsewhere in the pulmonary parenchyma, including smaller airways and alveoli (22, 37). A comparative pathogenicity of three SARS-CoV-2 variants, that is, B.1 D614G, Delta (B1.617.2), and Omicron (BA1.1), revealed contrasting pathology in respiratory tissues (22). The degree and duration of pathological changes observed in the respiratory tract were higher in B.1 D614G– and Delta-infected cats than in Omicron-infected cats. Striking differences are noticed in the lungs across variants, with the B.1 D614G– and Delta-infected cats showing marked bronchial and bronchiolar necrosis, degeneration, and mixed inflammation in lungs, whereas no necrotizing lesions are observed in Omicron-infected cats (22).
Immune responses to SARS-CoV-2 in cats
Studies addressing the immune responses of SARS-CoV-2 in cats are limited. Expression of innate immune cytokines, divergent lymphocyte phenotypes, and neutralizing Ab responses during SARS-CoV-2 infection in cats are summarized below.
High levels of proinflammatory cytokines such as IL-2, GM-CSF, and CXCL1 (neutrophil chemoattractant) at 3 dpi and IL-1β, TNF-α, IL-2, IL-4, IL-6, and CXCL1 at 7 dpi were observed in B.1 D614G– and Delta variant–inoculated cats when compared with Omicron-inoculated animals. In contrast, increased levels of anti-inflammatory cytokines IFN-γ and CCL5 were observed in Omicron-inoculated cats, which mirrored the lower pathogenicity of this variant in cats (22). However, more detailed functional studies, particularly their expression in the highly susceptible upper and lower respiratory tracts of cats, are needed to elucidate the role and contribution of innate immune cytokines in the pathogenicity of SARS-CoV-2 in this species.
A recent study reported a significant shift in systemic lymphocyte immunophenotypes during SARS-CoV-2 Delta variant infection in cats (63). A steady increase in the proportion of CD4+ T cells through 8 dpi was observed in infected cats suggesting a T cell–mediated proinflammatory (Th1) response that may contribute to immunopathology (63). Although CD8+ T cells that are essential for virus clearance during acute infection remained unaltered throughout the course of infection (63), the proportion of CD21+ B cells was reduced in infected cats throughout the study period, resembling observations in untreated human COVID-19 patients, possibly due to B cell exhaustion and exacerbated immune activation (63, 66). Further studies are required to elucidate the protective mechanisms of CD8+ T cells and the contribution of these and other lymphocyte subtypes in the control of infection and viral clearance from tissues in domestic cats.
Robust neutralizing Ab responses are developed by SARS-CoV-2–infected cats (42). Virus-neutralizing Abs are detected in serum as early as 7 dpi (22, 37, 41) and remain elevated for up to 28–42 dpi (41). Abs against NP and S-RBD are detected as early as 5 dpi in infected cats by ELISA (37). Serum IgG Ab responses against full-length S protein and the RBD as well as NP are detected at 7 dpi, which increase at 14 and 21 dpi (41). Seroconversion to the S protein seems to be fast and robust, and the specificity of the response to the RBD exceeds that of NP (41). Serum IgM Abs against the RBD are detected at 7 and 14 dpi but not at 28 dpi, and the IgM responses are less robust than the IgG responses (41).
Of note, cross-neutralization assays revealed cross-neutralizing Ab activity in serum from B.1 D614G–inoculated cats against the Delta but not against the Omicron variant. Interestingly, sera from Delta-inoculated cats presented neutralizing activity to both the B.1 D614G and Omicron variants. Animals infected with the Omicron variant, however, presented lower and delayed neutralizing Ab responses (35).
SARS-CoV-2 pathogenesis and immune responses in WTD
WTD are one of the most abundant large ruminant species in North America. The ACE2 protein of WTD shares high amino acid homology with that of human ACE2, thus allowing efficient binding of SARS-CoV-2 S protein and entry into cells (67). Such ACE2 homology makes WTD highly susceptible to diverse SARS-CoV-2 lineages of human origin (61). Detailed experimental studies in WTD revealed some of the basic aspects of the virus infection dynamics and pathogenesis in this species (56–58). A summary of SARS-CoV-2 infection dynamics in WTD is presented in Fig. 2 and discussed in further detail in the sections that follow.
Infection dynamics of SARS-CoV-2 in WTD
Infections of WTD with SARS-CoV-2 are usually subclinical (56–58). However, a slight and transient increase in the body temperature has been observed between 1 and 3 dpi in infected fawn and adult deer (56–58).
SARS-CoV-2 productively infects and replicates in the URT and associated lymphoid organs in WTD. During the acute stage of infection (2–5 dpi), infectious virus is consistently detected in a broad range of tissues, including nasal turbinate, tonsil (palatine and pharyngeal), lymph nodes (medial retropharyngeal, mandibular, and tracheobronchial), and in the trachea, bronchus, and lungs (58). Of note, infectious virus has also been detected in the olfactory lobe, caudate nucleus of brain, cerebrum, and cerebellum at 2 dpi in infected WTD (57). However, no infectious virus has been detected in any tissues after 5 dpi (56).
Virus shedding has been detected in nasal and oral secretions and less consistently in feces of infected WTD (56–58). In an experimental infection in fawns with the SARS-CoV-2 ancestral B.1 lineage, shedding of viral RNA has been detected in the nasal and oral secretions between 2 and 21–22 dpi, with higher viral RNA loads detected between 2 and 7 dpi and decreasing thereafter (56, 58). Infectious virus is detected in the nasal and oral secretions of infected animals between 2 and 5 dpi (56, 58), whereas no infectious virus was detected in feces (58). Shedding of viral RNA in feces was markedly lower and was characterized by intermittent detection of low amounts of viral RNA (56, 58). Consistent with active virus replication and shedding through respiratory and oral secretions, SARS-CoV-2 is efficiently transmitted between inoculated and contact WTD (56–58). Transmission to contact fawns was observed on 3 dpi, a time point when infectious virus shedding in nasal and oral secretions was abundant, but not on 6 or 9 dpi when no infectious virus was recovered from inoculated animals (58). Taken together, these results demonstrate that WTD enable active SARS-CoV-2 replication in the respiratory tract, with virus shedding and efficient transmission of the virus likely occurring through respiratory or oral secretions.
Pathology of SARS-CoV-2 in WTD
Infections with SARS-CoV-2 in WTD have not been associated with gross pathological lesions (56–58). Histologically, WTD infected with the ancestral B.1 lineage of SARS-CoV-2 presented with rhinitis characterized by submucosal lymphoplasmacytic infiltrates and frequent mucosal exudation of neutrophils in the nasal turbinate, and moderate to abundant staining of viral RNA and NP was detected at 2 and 5 dpi. Of note, viral RNA and Ag distribution in nasal turbinate epithelial cells is consistent with detection of high levels of expression of the virus receptor ACE2 and the serine protease TMPRSS2 in these cells (58). In lungs, diffuse congestion of alveolar capillaries and multifocal hemorrhage in the submucosa of larger airways as well as multifocal perivascular and interstitial lymphohistiocytic infiltrates are observed. Moderate to abundant viral RNA and Ag staining in the bronchial epithelial cells is observed at 2 dpi and is less evident at 5 dpi (58). In lymphoid organs (tonsil and lymph nodes), follicles exhibit moderate lymphoid depletion and lymphocytolysis. Tonsils also contain multifocal hemorrhages in crypt epithelium, crypts with variable numbers of neutrophils and cell debris, and congested vasculature filled with neutrophils. Numerous tonsillar follicles and overlying epithelium are positive for virus RNA and N Ag at 2 and 5 dpi (58). Notably, high ACE2 and TMPRSS2 expression levels have been detected in crypt epithelial cells and in germinal centers in the tonsil where active virus replication is detected (58). In addition, viral staining in the mandibular and/or retropharyngeal lymph nodes has been observed at 2 and 5 dpi, with the interfollicular and paracortical regions of these lymph nodes containing large foci of virus replication (58).
Immune responses to SARS-CoV-2 in WTD
SARS-CoV-2 infection in WTD results in the development of neutralizing Abs to the virus as early as 7 dpi (56–58). The geometric mean titers of neutralizing Abs detected in WTD infected with the ancestral B.1 lineage of SARS-CoV-2 vary from 37 to 107, 85 to 214, and 85 to 256 on 7, 14, and 21 dpi, respectively (56). Abs against S RBD and NP are also detected upon infection, and titers of Abs against the NP are lower and appear at later time points (14–18 dpi) (56, 57). Development and kinetics of neutralizing Abs parallel resolution of viral replication. No studies have been performed to date to assess T cell responses or define immunological cells and effector functions controlling infection in WTD.
SARS-CoV-2 presents a broad host tropism with a diverse array of susceptible domestic and wild animal species. Among these species, domestic cats and WTD are exquisitely susceptible to infection and highly efficient in virus transmission. The infection dynamics of SARS-CoV-2, including virus replication kinetics and tissue tropism, transmission dynamics, cellular and humoral immune responses, and pathological changes in the respiratory tract in SARS-CoV-2–infected domestic cats resemble SARS-CoV-2 infections in humans. These initial studies elucidated basic aspects of SARS-CoV-2 pathogenesis and immune responses in domestic cats; however, functional details of the host innate and cellular immune responses are still missing. Similarly, WTD are highly susceptible to SARS-CoV-2 infection and can transmit the virus and mount robust neutralizing Ab responses. Recent studies suggesting that WTD may serve as a new wildlife reservoir for diverse ancestral SARS-CoV-2 lineages deserve special attention. Furthermore, the immune responses and viral factors leading to clearance or to possible virus persistence in WTD should be further studied, as this may allow implementation of measures to effectively prevent or reduce infection rates in this animal population.
The authors have no financial conflicts of interest.
This work was supported by the Division of Microbiology and Infectious Diseases, National Institute of Allergy and Infectious Diseases Grant R01AI166791-01 and by National Institute of Food and Agriculture Grant 2023-70432-39463.