turkey rhinotracheitis

Pictures

Picture	Title	Caption	Copyright
Title Symptoms Caption Turkey poult infected with avian metapneumovirus, showing swollen infra-orbital sinuses. Copyright David Cavanagh	Symptoms	Turkey poult infected with avian metapneumovirus, showing swollen infra-orbital sinuses.	David Cavanagh
Title Symptoms Caption Turkey poult infected with avian metapneumovirus, showing frothy eye. Copyright David Cavanagh	Symptoms	Turkey poult infected with avian metapneumovirus, showing frothy eye.	David Cavanagh
Title Symptoms Caption Domestic fowl infected with avian metapneumovirus, showing swollen head and nasal exudate. Copyright David Cavanagh	Symptoms	Domestic fowl infected with avian metapneumovirus, showing swollen head and nasal exudate.	David Cavanagh

Identity

Preferred Scientific Name

• turkey rhinotracheitis

International Common Names

• English: APV infection; avian metapneumovirus infection; avian pneumovirus infection; avian rhinotracheitis; swollen head syndrome

Pathogen/s

Overview

Rhinotracheitis in turkeys is perhaps the most economically important disease caused by avian metapneumovirus (aMPV) amongst poultry species, although avian metapneumovirus represents a number of related viruses that can replicate to a greater or lesser extent in other galliformes, including the domestic fowl, and also in wild birds (Gough and Jones, 2008). The virus belongs to the genus Metapneumovirus in the sub-family Pneumovirinae of the family Paramyxoviridae. aMPVs have been detected in poultry in Africa, Asia, Europe, North America and South America.

The first isolate of aMPV was the cause of rhinotracheitis in turkeys in South Africa in the late 1970s (Buys and du Preez, 1980; Buys et al., 1989a). Four subtypes of aMPV are recognised: A, B, C and D. Subtypes A and B have been detected in poultry in Asia, Europe and South America but not in North America, whereas subtype C was discovered in North America and later in Europe (Toquin et al., 1999; Toquin et al., 2006). Subtype D was isolated in France in 1985 (Bäyon-Auboyer et al., 1999, 2000). The subtypes differ substantially in their genome sequences, the genomes being a non-segmented, single-stranded, negative sense RNA of approximately 13 500 nucleotides.

In turkeys aMPVs may, depending on the country, be the most important of the viral respiratory pathogens, causing substantial economic loss. The aMPVs also cause disease in domestic fowl though the extent is variable, even within a region, other respiratory viruses also having an impact. Breeders, layers and meat-type turkeys and chickens are affected. The disease may be manifest more as a rhinitis than a tracheitis, and the virus can replicate in reproductive organs. In addition to general diminution of productivity the virus predisposes to secondary bacterial infection, which can be fatal. A number of live and killed vaccines are available to control infections by subtypes A and B.

aMPVs have been detected in many wild and semi-wild bird species on the same continents as aMPVs in poultry (see Hosts/species affected, below).

Host Animals

Hosts/Species Affected

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.

Systems Affected

Distribution

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.

Distribution Table

The distribution in this summary table is based on all the information available. When several references are cited, they may give conflicting information on the status. Further details may be available for individual references in the Distribution Table Details section which can be selected by going to Generate Report.

Pathology

Turkeys

Examination of nasal turbinates following experimental infection revealed clear to greyish exudates, first watery and then mucoid. This lasted 9 days. In the trachea there was excess mucous up to 7 days (Jones et al., 1988). In the field, serious secondary bacterial infection of poults is more likely, with effects similar to those of infection of domestic fowl described below.

In one study experimental infection of 7-week-old poults with type C virus did not reveal gross lesions in turbinates, infraorbital sinuses and trachea, but microscopic examination showed acute rhinitis, sinusitis and tracheitis (Panigrahy et al., 2000). In another experimental study using 3-week-old poults, Jirjis et al. (2000) observed gross changes of clear frothy fluid in the sinuses up to 10 days after inoculation. Microscopic changes were observed in nasal turbinates and infraorbital sinuses but not trachea or other respiratory tissues, including infiltration by lymphocytes, macrophages and plasma cells.

Intranasal infection of laying turkey hens, not previously infected with aMPV, resulted in infection of the oviduct (Jones et al., 1988). White masses of inspissated albumen were commonly observed up to 12 days after infection. Less frequently, there were deposits of solid yolk material in the abdomen. One bird had a folded shell membrane in the magnum whilst another had egg peritonitis. Misshapen asymmetric eggs were present in the uterus. There were signs of early stage ovary regression, with follicles showing shrinkage.

Domestic fowl

The lesions in broilers have been described by Cook and Pattison (1996) as a fine petechiation of the turbinate mucosa, progressing to a severe generalized red to purple discoloration of the mucosa. Yellow, oedematous subcutaneous tissue is revealed when the skin over the head is removed. Pericarditis and perihepatitis may be seen (Nakamura et al., 1997). Histological analysis reveals marked fibrinopurulent inflammation with oedema in the subcutaneous tissues of the head. Also seen in these tissues were marked exudation of fibrin, serum, heterophils, lymphocytes and macrophage with Gram-negative bacilli and sometimes vasculitis and thrombus formation were seen. Purulent inflammation in the air spaces of cranial spongy bones may be observed. Respiratory tissues exhibit rhinitis, infraorbital sinusitis and tracheitis.

McMullin (1998) has described field aMPV infections in broiler breeders. Sinusitis and rhinitis were seen but not tracheitis. Pus in subcutaneous tissue was observed less frequently than in broilers although pus with fluid accumulation was observed in cranial bones. Cook and Pattison (1996) report extensive peritonitis, initially as a moist inflammatory lesion in the ovarian region, often with yolk free in the peritoneum. Subsequently, there are advanced inflammatory lesions of the peritoneum with large amounts of yolk material.

Diagnosis

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).

List of Symptoms/Signs

Disease Course

Jones (1996) has written a good review of several aspects of aMPV infections in turkeys and chickens.

Turkeys

Turkeys infected by aMPV exhibit sneezing, snicking, rales and head shaking. There is a nasal discharge, foamy conjunctivitis and swelling of the infraorbital sinuses. Some birds develop pus cores within the infraorbital sinuses, large numbers of Escherichia coli and other bacteria being present. This can lead to mortality, especially in poults (Grant, 1996; Jones and Wilding, 1996; Alexander, 1997). Concurrent infection of turkey poults with Mycoplasma gallisepticum and Mycoplasma imitans (Naylor et al., 1992; Ganapathy et al., 1998) but not Mycoplasma synoviae (Khehra et al., 1999), may exacerbate disease.

Morbidity is frequently 100% in flocks that have not been vaccinated or previously infected. In the first year (1985) of clinical disease in Britain mortality varied between 3% and 15%, with overall losses in stags/toms of up to 25% to 20 weeks and 20% in hens to 16 weeks (Pattison, 1998). When type C virus first affected the state of Colorado in the USA, 80 to 100% of a flock developed a cough which persisted for about 10 days. Three to four days after the commencement of coughing, 10 to 40% of the birds developed severe sinusitis, nasal discharge and dried mucus around the nares. Mortality was only 1 to 5% when secondary bacterial infections were controlled by antibiotics but up to 30% and more when the bacteria were not controlled (Senne et al., 1998).

Mortality can occur in vaccinated flocks, depending in part on the nature of the bacteria present and environmental factors (poor ventilation, high levels of ammonia and dust) that enhance infection by these secondary pathogens. Surviving birds recover within 10 to 12 days.

Infection in laying turkeys can result in loss of egg production of 10 to 15% but sometimes much higher, and sometimes egg quality (Jones et al., 1988), recovery taking 3 to 5 weeks.

Domestic fowl

The consequences of infection of domestic fowl by aMPVs can, as in turkeys, be very variable although usually they are less severe than in turkeys. Indeed, infected flocks of chickens may not always exhibit clinical signs. Secondary bacterial infections and environmental factors (poor ventilation, high levels of ammonia and dust) are important contributors to clinical disease following infection with aMPV (Cook and Pattison, 1996). Assessment of the impact of aMPV infections in domestic fowl is complicated in broilers by infections at around the same time with other viral respiratory pathogens, such as infectious bronchitis (Cavanagh et al., 1999).

Infected broilers may exhibit coughing and sneezing with nasal discharge. A small proportion, perhaps 5% or so, may have head swelling, which has given rise to the name 'swollen head syndrome'. This is associated with secondary bacterial infection and may lead to mortality.

Swollen heads are exhibited to a lesser extent by infected broiler breeders, perhaps 1% or less (Wyeth et al., 1987; Pattison et al., 1989; Jones et al., 1991) although Maharaj et al. (1994) observed swollen heads in 10% of breeders. The breeders sit with their neck arched, the head resting on the back, sometimes referred to as ‘star gazing’. There may be discharge from eyes or ears and swelling around the eyes and on top of the head. Such birds, when disturbed, lose coordination. Some may exhibit head rolling. Soiling around the vent may follow production of green diarrhoea. Egg production is adversely affected.

Females are affected more than males and commercial egg layers are affected by aMPV to a lesser extent than broiler breeders.

Strains of aMPV may differ in their virulence although on occasion this may be a consequence of partial attenuation during growth in laboratories. Notwithstanding, Cook et al. (1993a, b) reported that a chicken isolate grew to higher titres in specific pathogen free (SPF) chickens than did a turkey isolate. Strains also differ with respect to the severity of clinical signs within a given species.

Epidemiology

Following infection of turkeys and domestic fowl virus is released within 24 h, reaching a peak 3 to 5 days after infection and declining precipitously thereafter. Clinical signs may be evident at day three although they would probably be minor at this stage so that maximum release – and spread – of the virus may occur before the disease is fully recognized. Although the virus is undoubtedly released in aerosols, spread appears to require fairly close contact between birds. Spread is believed to be oral rather than faecal.

No specific biological vectors are known, the virus probably being spread by such agents as people, their vehicles and non-specifically by vermin. However, there is evidence for replication of aMPVs in game birds and many species of wild birds. Thus many avian species might be considered as potential carriers of aMPVs, with consequences for disease security. There is no evidence for vertical transmission.

Impact: Economic

When aMPV-related rhinotracheitis was first observed in turkeys in South Africa and Europe morbidity was usually >90% and mortality about 30% (Buys and du Preez, 1980; Buys et al., 1989a). Of course, as none of the birds had been vaccinated, economic losses were very high. Vaccines were subsequently developed and their use is widespread. The consequences of field infection were thereby diminished although some economic losses are still sustained despite vaccine application. The effects of aMPV in vaccinated flocks may be irregular; not all houses on a site will exhibit disease and sites will not necessarily exhibit disease with every successive crop. In the USA, where there is no vaccination against aMPV, aMPV-C caused 50-100% morbidity and 3-30% mortality in meat-type turkeys (Jirjis et al., 2000).

The economic effects of aMPV infection on domestic fowl have been more difficult to assess, in part because of widespread infections with other respiratory viruses. When aMPV vaccines have been applied to broilers some farms have yielded better economic performance whilst others have not. This may be interpreted as being indicative that certain, unspecified conditions apply to some farms with the consequence that infection by aMPV has significant effects on performance. In those cases vaccination may be beneficial.

Zoonoses and Food Safety

There is no known zoonotic or food safety issue regarding aMPVs.

Disease Treatment

There is no treatment for the viral infection. Antibiotics, administered in drinking water, have been used in attempts, sometimes successful, to minimize losses caused by secondary bacterial infections. Severely affected birds should be culled but moderately affected ones might be moved to a hospital pen and injected with antibiotics. Ventilation should be improved and ammonia and dust levels minimized. In-feed medication could be continued for a further two weeks.

Prevention and Control

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.

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