turkey rhinotracheitis

All types of turkeys are susceptible – commercial meat-type birds, parents and grandparents (Pattison, 1998). Generally, turkey poults are more likely to exhibit high mortality than juvenile and older birds – though mortality in these birds can be high if they have not been vaccinated or naturally infected whilst young. Significant mortality does not always occur, even in poults, as mortality is dependent in part by secondary bacterial infection. Environmental and management factors will affect the outcome, as is the case for other diseases.

Experimental studies have shown that clinical signs exhibited by domestic fowl after aMPV infection are generally of less severity than in turkeys (Catelli et al., 1998). As in turkeys, the virus can adversely affect the performance of young birds and adults.

All four subtypes of aMPV have been variously detected in wild and semi-wild bird species, including: pheasants (Gough et al., 1988; Lee et al., 2007); guinea fowl (Litjens et al., 1989); sea gulls, including herring gull, Larus argentus argentus (Heffels-Redmann et al., 1998) and ring-billed gulls (Bennett et al., 2004); Muscovy duck (Toquin et al., 1999); house sparrows (Bennett et al., 2004; Gharaibeh & Shamoun, 2012); snow geese (Bennett et al., 2004); wild birds in the orders Psittaciformes, Anseriformes and Craciformes, and the Anas and Dendrocygma genera (Anseriformes Order) in Brazil (Felippe et al., 2011); pigeons (Gharaibeh & Shamoun, 2012). Thus aMPVs have a wide host range, and many avian species possibly pose a disease security threat to poultry in respect of aMPVs.

Following the discovery of aMPV in turkeys in South Africa in the late 1970s, the virus was isolated from domestic fowl in South Africa (reported by Buys et al., 1989b) and UK (Morely and Thomson, 1984) and then from turkeys and domestic fowl in many European countries in the mid to late 1980s (McDougall and Cook, 1986; Giraud et al., 1986; Wilding et al., 1986; Wyeth et al., 1987; Picault et al., 1987) and elsewhere in the 1990s, including North America (Cook et al., 1993a, 1999; Dani et al., 1999a,b; Panigrahy et al., 2000) and subsequently, including in Brazil (D’Arce et al., 2005; Chacón et al., 2011), Israel (Banet-Noach et al., 2005), Japan (Mase et al., 2003); Korea (Lee et al., 2007; Kwon et al., 2010); and Turkey (Ongor et al., 2010). It can be seen that aMPVs are widely distributed, both in poultry, game birds and in wild birds.

Clinical Diagnosis and Lesions

The observations described in Disease Course and Pathology for infection of turkeys would be indicative of infection by aMPV but not diagnostic. In domestic fowl the clinical signs and lesions seen could be associated with a number of viruses and bacteria. Laboratory diagnosis is, therefore, required for confirmation.

Differential Diagnosis

Turkeys

The clinical signs exhibited by turkeys infected by aMPV are similar to those caused by other pathogens, including avian influenza virus, Newcastle disease virus, mycoplasmas, Bordetella avium, Pasteurella spp. and Aspergillus spp. Indeed, many pathogens were incorrectly implicated as the primary aetiological agent of turkey rhinotracheitis before the discovery of aMPV. Clinically, infection by Bordetella avium cannot be distinguished from that of aMPV. The name 'turkey rhinotracheitis' is best used for infections by aMPV, 'turkey coryza' being used for Bordetella avium infection.

Domestic fowl

Infection with infectious bronchitis virus and mild forms of Newcastle disease virus and avian influenza virus could produce similar clinical signs and pathology as aMPV, as would infection by Mycoplasma spp. and bacteria such as Haemophilus paragallinarum and Escherichia coli.

Laboratory Diagnosis

Laboratory diagnosis is essential for confirmation of aMPV infections.

Reverse transcriptase polymerase chain reaction (RT-PCR) tests are now the preferred means of laboratory diagnosis of active infections, and many have been described, including: (Naylor et al., 1997a,b; Bäyon-Auboyer et al., 1999; Cavanagh et al., 1999; Banet-Noach et al., 2005; D’Arce et al., 2005; Kwon et al., 2010; Chacón et al., 2011; Falchieri et al., 2012. This approach has been used directly on virus recovered on mouth, nose and tracheal swabs, avoiding the need for virus isolation. The identity of the PCR product may be confirmed by sequencing.

Antibody detection is also used to confirm aMPV infections. Several ELISAs are commercially available for this purpose. Ideally, paired sera should be examined for evidence of a rise in titre. A single live vaccination with aMPV generally stimulates a poor antibody response in contrast to infection with virulent virus. ELISAs made with either A or B types of aMPV have been used to detect infection with both types of virus but it is generally accepted that sensitivity is greater when the homologous antigen is used (Eterradossi et al., 1995; Toquin et al., 1996; Bäyon-Auboyer et al., 1999). Antibodies to type C virus were not detected by ELISAs with types A and B antigen (Senne et al., 1998; Toquin et al., 1999). Choi et al. (2010) have demonstrated that diagnosis of aMPV infection in laying hens by yolk ELISA to detect anti-aMPV antibodies may be a suitable alternative to testing serum.

Antibodies can also be detected by indirect immunofluorescence applied to sections of tissue; trachea has been most widely used. Neutralization tests may be applied, though less frequently than RT-PCR tests, using virus grown in tracheal organ cultures (ciliostasis being used to determine the endpoint) or, after adaptation, to chick embryo fibroblasts or Vero cells (absence of cytopathic effect or antigen detectable by fluorescence/immunoperoxidase staining being used to determine the endpoints). aMPVs do not cause haemagglutination, hence haemagglutination-inhibition tests cannot be used.

Primary isolation of aMPV is carried out using chicken tracheal organ culture (TOC) or chicken embryonated eggs with subsequent adaptation in chicken embryo fibroblasts (CEF) or Vero cultures for further investigation (Coswig et al., 2010). Virus isolation should be sought as soon as possible after development of clinical signs. Indeed, it is advisable to collect samples from birds exhibiting the earliest clinical signs. Sources of virus are: nasal turbinates and trachea; ocular, nasal or tracheal exudates; swabbings of these. Jirjis et al. (2000) were able to detect type C virus (by virus isolation and RT-PCR) in experimentally infected 3-week-old turkeys from nasal turbinates, but not from trachea, up to 6 days after infection by the ocular/nasal route.

Virus isolation from domestic fowl has been more difficult than from turkeys. A contributory factor may be that other viruses interfere with the replication of aMPV. For example, both vaccinal and virulent strains of infectious bronchitis virus interfere with the replication of aMPV (Jones et al., 1998). Also, if other viruses are present these may outgrow aMPV during virus isolation in, for example, tracheal organ cultures.

Although aMPVs can be adapted to grow in chick embryo fibroblasts (CEFs; Collins et al., 1993), Vero cells (Buys et al., 1989a; Cavanagh and Barrett, 1988), chick embryo rough cells (CER) (Dani et al., 1999a, b) and QT35 quail cells (Goyal, 1998) these are not generally regarded as being optimal for primary isolation. In many laboratories this has been done for aMPV types A and B using tracheal organ cultures from chick or turkey embryos, the presence of aMPV being suggested by ciliostasis (McDougall and Cook, 1986; Jones et al., 1991). Ciliostasis takes much longer to occur than with, say, infectious bronchitis virus. Cultures should be observed for up to 11 days and further passages may be necessary. Alternatively, one might culture the virus for 2 or 3 days, passage it once for the same period and then passage a third time, observing for up to 11 days. Confirmation that the ciliostatic agent is aMPV must be sought, for example, by immunofluorescence. This can be done without fixing the tracheal rings (Bhattacharjee et al., 1994). The Colorado strain of aMPV was poorly ciliostatic (Cook et al., 1999). A pneumovirus in Muscovy ducks was isolated using Vero cells (Toquin et al., 1999).

Primary isolation may also be attempted using chick or turkey embryos inoculated via the yolk sac (McDougall and Cook, 1986). The Colorado aMPV was isolated by inoculating 6-day-old embryonated domestic fowl eggs via the yolk sac and incubating for 9 days. After a second 9-day passage in embryos, a yolk sac membrane homogenate was inoculated on CEFs and passaged again on CEFs, whereupon a cytopathic effect was observed. Goyal used seven to eight blind passages in CEFs and QT35 cells for primary isolation of the Colorado strains (Goyal, 1998). Passage of the Colorado virus first in chick embryos, via the yolk sac, and then in CEFs has also been described by Panigrahy et al. (2000).

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Immunology of the disease

Live aMPV vaccines tend to induce a poor serum antibody response. Certainly the absence of detectable antibody after vaccination is not a cause for concern. Field infection is likely to result in detectable antibodies. Vaccinated poults without detectable aMPV antibody are protected against challenge, immunity taking a week or more to develop (Cook et al., 1989; Jones et al., 1992). Vaccination or challenge of laying hens results in progeny with maternally derived antibody (MDA). Experiments indicated that MDA did not prevent the development of clinical disease in poults infected at up to 10 days of age (Naylor et al., 1997b). There is evidence of temporary immunosuppression – impaired humoral and cell-mediated immunity – after natural infection (Timms et al., 1986).

Immunization and Vaccines

Vaccination is the main form of control in those countries where aMPV infection is widespread. Live and killed vaccines are available to manage infections by subtype A and B viruses. Although the vaccines are effective, it has been demonstrated that live aMPV vaccines have a tendency to revert to virulence (Lupini et al., 2011). Also, there is evidence for the mutation of field virus in response to sustained vaccination (Cecchinato et al., 2010). Some variants have an increased capacity to break through the immunity induced by available commercial vaccines (Catelli et al., 2010).

A number of vaccination protocols have been used.

In meat-type (commercial) turkeys it is common to apply live virus vaccine by eye drop or coarse spray in the hatchery. Sometimes it is applied at 7-10 days of age. Where field virus of both types A and B are present some growers have applied a mixture of type A and B vaccines. Others have vaccinated at 1-day-old with one type and vaccinated with the other type later. Turkey breeders have a live primer vaccination applied by coarse spray at about 2 weeks of age, followed by injection of a killed vaccine at about 22 weeks of age (Cook et al., 1996).

Where aMPV is considered to be an economic problem in broiler chickens the birds may be given live vaccine at the hatchery or at 7 days of age. Broiler breeders and commercial layers may be given live vaccine at 10 to 12 weeks of age followed by killed vaccine at 16 to 20 weeks of age. Depending on the dominant type of aMPV in a region, one might use a live type B vaccine and a killed type A vaccine in an attempt to maximize protection. Infectious bronchitis virus vaccines have been shown to interfere with the replication of vaccinal and field aMPVs although immunity to aMPV was still induced (Jones et al., 1998). Ganapathy et al. (2010) compared the induction of protection in chickens following vaccination by three different routes: spray, drinking water and oculo-oral application. The authors concluded that accurate vaccination by spray and drinking water achieved levels of protection equal to that induced by the oculo-nasal application.

Farm-Level Control

In addition to vaccination (where appropriate) good biosecurity should be practised, as always, including attention to temporary storage and disposal of carcasses at a farm. Multi-age sites are particularly at risk. As with other viral diseases, air quality is important to reduce stress on the respiratory mucosa which, together with aMPV infection, can predispose the birds to secondary bacterial infections. Thus control of ventilation, heating and possibly misting if litter is very dry is important. Overstocking of poults up to 6 weeks of age is to be avoided.

After the first instances of aMPV infection in Colorado, USA, the disease was eliminated from that state by depopulation, cleaning, a good interval before repopulation, marketing restrictions and good biosecurity in general.