Feline Morbillivirus infection

 

Edited April 2020

 

These guidelines were drafted by Maria Grazia Pennisi, Margaret J. Hosie, Séverine Tasker, Diane D. Addie, Karin Möstl et al.

 

 

Agent properties

 

The Morbillivirus genus (family Paramyxoviridae) includes important viral RNA pathogens of humans and animals including measles virus, canine distemper virus (CDV), rinderpest virus (globally eradicated in 2011), peste des petits ruminants viruses and viruses affecting marine mammals (Nambulli et al., 2016). CDV and related morbilliviruses have been shown to naturally infect wild and captive large felids (Appel et al., 1994; Roelke-Parker et al., 1996; Myers et al., 1997; Daoust et al., 2009; Meli et al., 2010; Terio and Craft, 2013) and they can be an important threat to wildlife conservation. In domestic cats, CDV infection has never been documented, but limited replication was observed in macrophages in asymptomatic experimentally infected cats (Bart et al., 2000). A paramyxovirus-like agent was isolated in 1981 from a cat with demyelinating lesions in the central nervous system (CNS) and intracytoplasmic inclusion bodies in glial cells (Cook and Wilcox, 1981). Also, cats are susceptible to highly pathogenic paramyxoviruses belonging to the Henipavirus genus (Hendra virus and Nipah virus) that infect humans and animals, but these agents have not been reported in Europe.

 

In 2012 a new paramyxovirus, subsequently named feline morbillivirus (FeMV), was first isolated from stray cats in Hong Kong (Woo et al., 2012). The virus was detected in renal tubular cells and lymph nodes in two cats affected by tubulointerstitial nephritis (TIN) and a small case-control study reported an association between FeMV and TIN (Woo et al., 2012). The virus is characterized by genetic diversity amongst isolates (Sakaguchi et al., 2014; Sieg et al., 2015; Park et al., 2016; Donato et al., 2019; Stranieri et al., 2019; De Luca et al., 2020) and recently whole genome sequencing of an isolate obtained in Germany revealed a new genotype, tentatively named feline morbillivirus genotype 2 (FeMV-GT2) (Sieg et al., 2019). Phylogenetic analysis demonstrated that FeMV-GT2 is genetically closely related, yet distinct, from previously described feline morbilliviruses.

 

Feline morbillivirus in vitro propagation in cell lines derived from 13 different mammal species (including humans) was evaluated and was observed only in feline cell lines or, less efficiently, in kidney cells from African green monkeys (Sagakuchi et al., 2015). Feline cell lines supporting viral replication included renal, fibroblastic, lymphoid and glial cells (Sagakuchi et al., 2015). FeMV-GT2 infected both renal cells and also primary epithelial lung cells, lymphocyte subsets, monocytes, and primary cells from cerebrum and cerebellum (Sieg et al., 2019). Recently, the tropism of FeMV-GT1 was investigated by IHC and immunoreactivity was detected in lung, kidney and brain sections (De Luca et al., 2020). These findings reveal new potential target tissues for FeMV infection in vivo, which might vary according to the virus genotype.

 

Evaluation of FeMV thermal stability was investigated in infected cell cultures; viral replication stopped after a few minutes of incubation at temperatures ≥ 60°C and progressively decreased, and was inhibited after 12 days, at 37°C. Virus replication was not inhibited at temperatures as low as 4°C, or following freeze-thawing (Koide et al., 2015).

 

Epidemiology

 

Following its initial discovery, FeMV was subsequently detected in Asia, in cats from Japan (Furuya et al., 2014), Thailand (Chaiyasak and Techangamsuwan, 2017) and Malaysia (Mohd Isa et al., 2019) and in the Americas in the USA (Sharp et al., 2016) and Brazil (Darold et al., 2017). In Europe, FeMV infection was documented in Italy (Lorusso et al., 2015; De Luca et al., 2018; Donato et al., 2019; Stranieri et al., 2019), Germany (Sieg et al., 2015, 2019), and the UK (McCallum et al., 2018), and isolates were obtained from the urine of cats in Italy and Germany (Lorusso et al., 2015; Sieg et al., 2018; Donato et al., 2019). Additionally, FeMV was reported in cats from Turkey (Yilmaz et al., 2017). Based on the above studies, it is clear that FeMV infection has been reported worldwide using PCR to detect virus in urine or kidney tissue. Urinary FeMV shedding (positivity range: 0.2-51%) and kidney infection ( positivity range: 7.4-80%) were variably found by PCR in both healthy and sick cats; such variation could be attributed to geographical differences in the level of endemicity and the different characteristics of the tested populations (in relation to age, husbandry, lifestyle or health status of cats) as well as analytical differences between the PCR techniques used (Woo et al., 2012; Furuya et al., 2014, 2015; Sieg et al., 2015, 2019; Sharp et al., 2016; Park et al., 2016; Darold et al., 2017; Yilmaz et al., 2017; De Luca et al., 2018; Mc Callum et al., 2018; Mohd Isa et al., 2019; Stranieri et al., 2019).

 

A higher prevalence of urinary PCR positivity was reported in male cats compared to females (Mohd Isa et al., 2019) and in tomcats compared to neutered male cats (Park et al., 2016). Very high (53%) urinary PCR positivity was reported in cats in a cat shelter (Darold et al., 2017), but another study reported a higher prevalence in pet cats compared to shelter cats (Mohd Isa et al., 2019). The urine of cats from suburban/rural areas or from those with outdoor access more frequently tested FeMV PCR positive (Yilmaz et al., 2017; Donato et al., 2018). Similarly, a higher prevalence was detected in colony cats compared to household cats (De Luca et al., 2020).

 

Some studies investigated exposure to FeMV in cats by the detection of serum antibodies against viral N protein, using Western blot (WB) (Woo et al., 2012; Mc Callum et al., 2018) or indirect immunofluorescence antibody tests (IFAT) (Park et al., 2016; Donato et al., 2018; De Luca et al., 2020) . Antibody tests revealed a level of exposure in cats of between 18.5 and 28%, but in a study performed on geriatric cats in the UK antibody prevalence was much higher at 64% (Mc Callum et al., 2018), and it was concluded that this might have been associated with a chronic persistent course of FeMV infection.

 

Pathogenesis and clinical signs

 

Experimental studies are not available and there is no information about routes of transmission, initial phase nor the course of natural FeMV infection. The epidemiological data available do not clarify whether transmission requires close direct between cats.

 

Woo et al. (2012) investigated the presence of FeMV by PCR in blood and urine as well as in nasal and rectal swabs. Positive results were obtained in only 0.2% of blood samples and 0.9% of rectal swabs, compared to 11.6% of urine samples. Blood was rarely tested in clinical samples; however, viraemia is probably transient as EDTA blood tested positive far less frequently than urine (Furuya et al., 2014; De Luca et al., 2018) or did not test positive (Mohd Isa et al., 2019; De Luca et al., 2020). The persistence of viral shedding in urine was investigated in different studies. FeMV RNA was detected for 4 months in a cat who died with chronic kidney disease (CKD) (De Luca et al., 2018), and after 15 months in one healthy cat that showed stable viral loads and a high antibody titre pointing to chronic infection and persistent shedding in the urine (Sharp et al., 2016). Interestingly, another cat presenting with proteinuria and cylindruria when testing FeMV positive in urine, was diagnosed and treated for cholangiohepatitis; when rechecked eight months later, the urine tested negative and proteinuria and cylindruria were no longer present (Stranieri et al., 2019). Two  cats suffering from renal problems shed RNA of FeMV for more than six months and two years respectively, in spite of high virus-neutralising antibody titres (Sieg et al., 2019). Recently, five FeMV positive colony cats which died 8-10 months following the initial detection of FeMV RNA in urine samples still tested positive in urine (and at least one tested tissue) and no mutations were observed (De Luca et al., 2020).

 

When post mortem tissue samples (brain, cerebellum, heart, lung, intestine, stomach, liver, urinary bladder, spleen, mesenteric lymph nodes), in addition to the kidney, were tested by PCR, positivity was found in the urinary tract (kidney and bladder), lymphoid tissue (spleen and mesenteric lymph nodes) and brain (De Luca et al., 2018, 2020). In lymph node sections, positivity was associated with macrophages (Woo et al., 2012). A higher percentage of positive samples was found in kidney tissue (range: 7.4-80%) compared to any other tissue or urine (Woo et al., 2012; Furuya et al., 2014; Park et al., 2016; Yilmaz et al., 2017; De Luca et al., 2018, 2020; Mohd Isa et al., 2019; Stranieri et al., 2019).

 

The relationship between infection or exposure to FeMV and CKD or TIN diagnoses was investigated retrospectively in live or dead cats. Woo et al. (2012) and Sieg et al. (2015, 2019) demonstrated an association between PCR positivity and CKD or other urinary pathologies, but no association was found by other authors (Darold et al., 2017; Mc Callum et al., 2018; Yilmaz et al., 2017; Mohd Isa et al., 2019; Stranieri et al., 2019; De Luca et al., 2020). However, such studies are sometimes limited if inappropriate criteria are used to select CKD and control cases, or few cats are tested. In cross-sectional studies there is a risk of false negative PCR results in case of intermittent viral shedding. Moreover, CKD can be clinically diagnosed months or years after a pathogen triggered the pathologic process leading to the development of TIN, but the infection may have resolved in the meantime. Additionally, many infectious and non-infectious factors are known to cause feline CKD (www.IRIS-kidney.com), and they are not easily investigated and recognized in field studies. Kidney histological and immunohistochemical (IHC) evaluation performed in some studies aimed to detect any association between the presence of lesions and the detection of FeMV, and possibly lesional occurrence of the virus (Woo et al., 2012; De Luca et al., 2018, 2020; Sutummaporn et al., 2019). A positive association was found between TIN and FeMV (either on urine PCR and/or WB) positivity in a controlled study that enrolled a limited number of cats. Additionally, the virus was demonstrated by immunohistochemistry (IHC) in renal tubular cells in two positive cats with aggregates of inflammatory cells in the renal interstitium, and renal tubular degeneration (Woo et al., 2012). In two other necropsied cats studied by IHC, viral antigen was demonstrated in the cortical tubules associated with small clumps of inflammatory mononuclear cells and in necrotic tubular cells surrounded by inflammatory infiltrate; additionally, strong and diffuse immunoreactivity was seen in tubules within the medulla with a mild inflammatory mononuclear infiltration (De Luca et al., 2018). Interestingly, in a recent study of 38 cats, FeMV IHC positivity was significantly associated with kidney lesions, particularly tubular and interstitial lesions (Sutummaporn et al., 2019). Moreover, in this study the tissue injury score of tubular lesions was higher in positive tubular sections and glomerulosclerosis was associated with FeMV positivity (Sutummaporn et al., 2019). Recently, IHC investigation of seven positive cats found immunoreactivity within epithelial cells of renal tubuli and lympho-plasmacytic cells infiltrating the tubular and interstitial areas in positive kidneys, but no association was found between FeMV positivity in kidney tissue (7/35) and evidence of renal lesions (23/35) or of TIN (14/35) (De Luca et al., 2020).

 

In conclusion, the impact of feline morbilliviruses on feline health has to be confirmed by further research, but a role in CKD development is strongly suspected and the potential damage of other organs cannot be excluded.

 

Diagnosis, prognosis, treatment and prevention

 

Urine and kidney tissue are the best samples for RNA detection by PCR and in most studies nested reverse transcriptase (RT)-PCR assays (targeting L, N or H genes) have been used (Woo et al., 2012; Furuya et al., 2015; Sieg et al., 2015, 2019; Park et al., 2016; Darold et al., 2017; Yilmaz et al., 2017; Mc Callum et al., 2018; Stranieri et al., 2019). Quantitative PCR (qPCR) for FeMV was developed by Sharp et al. (2016) and afterwards De Luca et al. (2018) found that this qPCR assay was more sensitive.

 

Western blot and IFAT assays were used in studies evaluating antibody response to FeMV, but cross-reactivity between CDV and FeMV was documented (Sakaguchi et al., 2014). An ELISA test detecting anti-FeMV P protein has been optimized (Arikawa et al., 2017). Virus neutralizing antibodies were measured by Sieg et al. (2019) to monitor chronically shedding cats. Histopathology with IHC evaluation of renal lesions associated with FeMV infection can be used, but renal biopsy is an invasive technique and cannot be recommended to confirm FeMV infection.

 

Currently FeMV diagnostic investigation is restricted to research laboratories and according to current knowledge ABCD does not recommend to routinely test cats for FeMV infection or exposure.

 

However, the prognosis and management of cats with urinary positivity to FeMV relies on addressing any concurrent CKD present and the use of the IRIS guidelines for the diagnosis, staging and management of CKD in cats (www.iris-kidney.com). Infected urine is probably the main source for transmission of FeMV infection between cats, particularly in catteries where cats are sharing litter trays.

 

Acknowledgement

 

ABCD Europe gratefully acknowledges the support of Boehringer Ingelheim (the founding sponsor of the ABCD) and Virbac.

 

References

 

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