HERPESVIRUS INFECTIONS IN WILD BIRDS

ERHARD F. KALETA

Clinic for Birds, Reptiles, Amphibians and Fish, Faculty of Veterinary Medicine, Justus Liebig University, Giessen, Germany

Avian herpesviruses (AHV) are widespread in domestic poultry (chickens, turkeys, Pekin ducks, geese and Muscovy ducks) and cause a variety of conditions in domestic and free-ranging wild bird species (Table 1.4).

TABLE 1.4 Avian herpesviruses, their natural hosts and predominant types of macroscopic lesions.

Lesions caused by AHV can be grouped into three cat­egories.

The taxonomic position of all currently known avian herpesviruses, order Herpesvirales, family Herpesviridae, subfamily Alphaherpesvirinae, has been reviewed⁽⁹⁸⁾. Atotal of nine herpesviruses are still unassigned to any genus. Recent publications contain descriptions of a further nine avian herpesviruses that are as yet incompletely character­ized and are not included in the list of assigned herpesvi­ruses (Table 1.5).

DUCK PLAGUE/DUCK VIRAL ENTERITIS

In Europe duck plague is an important herpesvirus disease of domestic Pekin and Muscovy ducks, many species of free-living ducks, geese and swans, and is occasionally found in other aquatic birds. Duck plague is also called duck viral enteritis (DVE). Both names are misleading, because not only ducks can be infected and enteritis is not the only lesion in clinical cases. Other names for duck plague are fowl plague and eendenpest (in Dutch).

AETIOLOGY

Duck plague virus is currently considered to belong to the family Herpesviridae as Anatid herpesvirus 1 (AnHVI) but is not yet assigned to any subfamily or species⁽⁹⁸⁾. However, placing it in the subfamily Alphaherpesvirinae has been proposed⁽¹⁰¹⁾.

TABLE 1.5 Avian herpesviruses in the family Herpesviridae, subfamily Alphaherpesvirinae.

‘See Kaleta, 2008⁽⁹⁹⁾

^bSee de Thoisy et al., 2009⁽¹⁰⁰⁾

nuclear membrane, particles enter the perinuclear spaces and the endoplasmatic reticulum of the cytoplasm.

EPIDEMIOLOGY

GEOGRAPHICAL DISTRIBUTION IN EUROPE

The majority of outbreaks have been described in North­ern and Central Europe, and North America. Clinical disease is predominantly seen in domesticated waterfowl.

Both migrant and resident species of waterbirds can be affected. The virus may be distributed from circumpolar regions of Eurasia and North America to the Southern regions of these continents by migrating birds.

HOST FACTORS

Subclinical and latent infections occur frequently in many species and are independent of age and sex. Many species are susceptible to infection (see Table 1.6), although Euro­pean teal (Anas crecca) and pintail (Anas acuta) appear resistant to experimental infection but still produce antibodies. During outbreaks a marked variation in species susceptibility is frequently observed. In a recent outbreak in domestic ducks and geese, many species that had not been considered susceptible before were affected⁽¹⁰²⁾. Like­wise, AnHV1 was isolated in Spain from common coots (Fulica atra) and crested coots (Fulica cristata), species that were previously considered resistant⁽¹⁰³⁾.

Natural infections have been described in ducklings as young as 7 days of age and in adult birds. Both sexes are equally susceptible. Stress due to physiological moulting,

TABLE 1.6 Hosts of duck plague virus of Eurasian anseriforms.

Degree of susceptibility: S = susceptible; M = moderately susceptible; R = resistant to infection

Some species of birds are of American or non-European origin but are kept in captivity in Europe

courtship, egg laying and incubation aggravates the clinical course of the disease.

ENVIRONMENTAL FACTORS

Most cases of duck plague in Europe occur during the winter, from January onwards, to early spring. The change from latency to clinically overt disease is regularly associ­ated with environmental stressors such as aquatic pollution and prolonged periods of freezing temperatures that result in the gathering of large flocks of susceptible birds on small areas of unfrozen water. Such conditions create environ­mental stress factors and facilitate virus transmission. The effect of stress has been studied experimentally; oral administration of cyclophosphamide, an immunosuppres­sant agent, resulted in decreased resistance following chal­lenge with a duck plague virus isolate, which did not cause mortality in immunocompetent mallards¹-¹⁰⁴). The practice of keeping large numbers of ducks and geese of different species in restricted and confined captivity to prevent exposure to avian influenza A virus, resulted in an out­break of duck plague in 2007 in Germany¹-¹⁰²).

EPIDEMIOLOGICAL ROLE OF AFFECTED SPECIES

Free-living, diseased and subclinically infected (with entire, infective virus) or latently infected (with viral genome that is not necessarily infective) waterfowl are considered to be the source of virus for susceptible free- living birds and domestic waterfowl. This is supported by the observation of seropositive subclinical virus carriers among free-living waterfowl. Infected wild birds may access farms with highly susceptible domestic Pekin ducks, Muscovy ducks and geese, infecting these domestic waterfowl and causing significant mortality among them.

TRANSMISSION

Infected birds excrete large quantities of duck plague virus in faeces and saliva, which results in contamination of water and grazing grounds. Oral and nasal infection is the most likely route for acquiring natural infection. Egg (ver­tical) transmission of the virus has never been confirmed. Living vectors are not required for virus transmission.

AnHV-1 can persist throughout life in a latent form in the trigeminal ganglion (TG), lymphoid tissues and in peripheral blood lymphocytes. Conversion may then occur, promoting latency to subclinical and productive infection of infectious virus shed from the oropharynx and cloaca. The excreted virus remains infectious in contami­nated fresh water for several days. The contamination of feeding grounds and roosting sites provides opportunities for lateral transmission. The infective dose is unknown.

PATHOGENESIS, PATHOLOGY AND IMMUNITY

The first steps of virus multiplication occur following oro- nasal infection of the upper respiratory and digestive tracts. After the initial infection, the virus is phagocytosed by macrophages and transported to the bloodstream, result­ing in viraemia and colonization of internal organs, includ­ing the intestines. Parenchymatous organs develop focal haemorrhages followed by necrosis. The small intestinal lesions frequently comprise one or more ring-like areas of haemorrhages that can be seen from the serosal surface. These lesions are considered pathognomonic for duck plague. The oesophagal and cloacal mucosa contain haem­orrhages, which develop into large necrotic areas. Haemor­rhages and necrosis in the bursa of Fabricius are of diagnostic importance.

Owing to the rapid course of the disease, dead birds are generally in good body condition. Birds that have suffered for prolonged periods from detrimental environmental con­ditions may be in poor condition. As a result of extensive haemorrhages and loss of blood into the intestine, the body appears pale during necropsy. Prominent lesions are present in the proximal to distal parts of the intestine. The mucosa of the oesophagus, proventriculus, intestine, cloaca and bursa of Fabricius contain multiple haemorrhages that develop into extensive layers of necrosis of the mucosal surface and submucosa. The surface of the heart and the myocardium may show petechiae, ecchymotic or extended haemorrhages. The thymus shows initially a haemorrhagic inflammation, which changes during the course of the disease to necrosis, distinct atrophy and almost complete loss of thymocytes. The surface of the enlarged liver has a copper- like colour with intermingled small haemorrhages and pinpoint foci of necrosis. At later stages the liver appears dark bronze in colour and some areas are stained by bile. In more protracted cases these haemorrhages are replaced by large areas of necro­sis. The spleen is enlarged and contains pale foci. The kidneys are swollen. The respiratory tract is not altered.

Microscopically, during the acute phase, multiple haem­orrhages are prominent in almost all organs. These are subsequently replaced by necrosis in organs and ulcera­tions in the intestinal mucosa. INIB in the intestines and internal organs can be seen in the vicinity of the necrotic lesions.

The necrosis of lymphoid cells in the bursa of Fabricius and in the gut-associated lymphoid tissues (GALT) alters the immune responsiveness so that serum antibodies are either completely absent or only detectable in low titres in virus neutralization tests.

Death occurs as a result of anorexia and extended haem­orrhages in intestines and parenchymatous organs. Necro­sis of the intestinal mucosa results in invasion of bacteria with subsequent bacteriaemia. Infected ducks usually die after a few days of illness.

CLINICAL SIGNS AND TREATMENT

Clinical signs vary widely among species. The time interval between infection and the appearance of the first signs of disease is estimated to be between 3 and 7 days. Death usually follows 1 to 3 days later. Duck plague is clinically characterized by sudden onset of mortality without specific premonitory signs. Some susceptible birds appear listless, are reluctant to move and to fly, and the intake of food and water is reduced. Species-specific vocalization is absent in sick birds, even during handling. Occasionally, abnor­mal movements of the head and neck (torticollis) can be observed. Intestinal discharge may be watery, greenish and intermingled with fibrinous material. In severe cases, blood clots can be seen in the faeces. Other signs such as swollen eyelids, drooping of wings and incoordination are only rarely seen. Infection during egg laying results in smaller than normal clutch sizes but egg size and shell structure are not affected.

There is no known effective treatment of clinical duck plague. Palliative measures such as fluid therapy to com­pensate dehydration, provision of appropriate food enriched with vitamins and treatment of bacterial infec­tions and internal parasites can be attempted to aid recovery.

DIAGNOSIS

Clinical signs are not specific. Gross pathology is of major diagnostic value.

Virus isolation is necessary to confirm a diagnosis of duck plague. Two to three days post- inoculation, a round-cell type cytopathic effect appears in susceptible cell cultures. Electron microscopy on purified and concen­trated gut content or faeces and ultrathin tissue sections can be useful for the detection of herpesviral particles. The application of PCR for accurate and rapid diagnosis is currently the method of choice(¹⁰5). PCR is performed on tissues (liver, kidney, spleen, intestines, cloaca) or on swabs from the cloaca or pharynx of live birds. The knowledge of the genome greatly supports the differentiation of AnHV-1 from other closely related viruses and is particu­larly useful for large-scale epidemiological studies.

Convalescent and immunized birds develop antibodies that can be detected in serum and egg yolk by a virus neutralization test. This test is useful for sero - epidemiological studies of all birds that are susceptible to duck plague virus. ELISA for detection of antibodies were successfully applied in commercial Pekin duck farms but have not been evalu­ated for testing of free-living Anseriformes.

MANAGEMENT, CONTROL AND REGULATIONS

Local outbreaks of mortality are not considered to pose a threat to any European waterfowl species, and no interven­tion is required. Birds that recover from natural infection appear resistant to reinfection.

An inactivated vaccine could potentially be applied to protect susceptible birds without the risk of introducing a modified live virus in free- living populations. However, formalin-inactivated adjuvanted vaccines had only a limited effect on subsequent experimental challenge. An inactivated vaccine is not commercially available in Europe. A live chicken embryo-adapted vaccine was developed to protect exposed juvenile and adult ducks¹-¹⁰⁶). Revaccina­tion at yearly intervals is necessary if breeding birds are kept. Unfortunately, this attenuated live- virus vaccine is considered suitable only for specific situations applicable to a small number of birds, and it is therefore currently difficult to obtain from European vaccine manufacturers.

Duck plague is a reportable disease in the USA but not in European countries. There are no specific regulations from the European Union or other countries in Europe for monitoring and control of duck plague.

PUBLIC HEALTH CONCERNS

Duck plague herpesvirus is not transmissible to mammals and is of no public health concern. However, humans who are involved in the health monitoring of free-living birds may act as important mechanical vectors of the virus.

SIGNIFICANCE AND IMPLICATIONS FOR ANIMAL HEALTH

In the USA, a die-off in the neighbourhood of the Lake Andes National Wildlife Refuge, South Dakota in 1923 resulted in the death of approximately half of the 100 000 wintering waterfowl (¹⁰7). However, duck plague outbreaks of similar dimensions have not been reported in Europe. Nevertheless, the Lake Andes disaster clearly demonstrates that duck plague can assume devastating proportions among wild birds. The source of duck plague virus in domestic ducks and geese is unknown in almost all out­breaks. Free-living European waterfowl, especially the mallard, are frequently implicated as the source of virus, but without definitive proof. Carnivorous free-ranging mammals are not susceptible to duck plague virus but may act as mechanical vectors.

Recovered birds should be tested and only released to the wild if AnHV- 1 and antibodies against it are not detected. Mixing of domestic waterfowl of unknown her­pesvirus disease status, and wild waterfowl at farms or rehabilitation centres should be avoided.

MAREK'S DISEASE

Marek’s disease (MD) (synonyms: polyneuritis gallinarum, fowl paralysis) is highly contagious and widespread in Europe in commercial and ornamental breeds of chickens, turkeys and quails. It is caused by Gallid herpesvirus 2 (Marek’s disease virus type 1) and Gallid herpesvirus 3 (Marek’s disease virus type 2). An additional member of the genus Mardivirus is the Meleagrid herpesvirus 1 (turkey herpesvirus 1), which is commonly isolated from turkeys and chickens. As turkey herpesvirus is avirulent for all gallinaceous birds, it is widely used as a live virus vaccine for chickens to prevent losses due to MD.

Virtually all countries and regions in Europe with domestic chicken and turkey populations are infected with MD viruses (MDV) of different virulence.

Chickens (Gallus gallus) are susceptible to MDV Genetic background (blood group alleles) influences the severity of the disease. Domestic Japanese quail ( Coturnix japonica) and probably free-living European Common quail ( Coturnix coturnix) can be infected under natural and experimental conditions and develop viraemia and tumor­ous lesions. The domestic turkey (Meleagrisgallopavo) may have visceral tumours but rarely neural lesions. Wild turkeys and other gallinacious species and birds of other orders resist infection. Very young chicks are more suscep­tible than juvenile or adult chickens. Gross pathological and histopathological lesions consistent with MD, but without demonstration of the virus, have been described in a large number of different avian species. In Europe these include the common buzzard (Buteo buteo), spar­rowhawk (Accipiter nisus), mallard (Anas platyrhynchos), eagle owl (Bubo bubo), little owl (Athene noctua), domestic goose (Anser anser), mute swan ( Cynus olor) and others⁽¹⁰⁸⁾.

MDV-infected chickens, quails and turkeys are the natural reservoirs, and may shed MDV throughout life. MDV matures only in cells of the feather follicle epithe­lium. Large numbers of MDV are found in epithelial cells and feather dander, from where they may be adsorbed onto dust particles. Transmission is facilitated by inhala­tion of virus containing dust. MDV is not vertically trans­mitted. Inhalation and conjunctival infection are the dominant routes of infection.

No obvious gross lesions are present in subclinically infected chickens. Neural lesions are associated with macroscopically visible thickening and discolouration of peripheral nerves, especially the vagus nerve, brachial plexus and ischiadic plexus. The ocular form consists of a unilateral iridocyclitis and panophthalmy. Large tumours are most frequently present in the ovary or testes and in the proventriculus. Histopathologically, nerves are oede- matous, have focal accumulations of small lymphocytes (‘Marek’s cells’) and a proliferation of Schwann’s cells. Tumours are composed of small lymphoid cells of the T-cell type.

Live virus vaccination of newly hatched commercial chicks is common practice in hatcheries. In Germany, but not in other European countries, only the tumorous and neural forms of MD in chickens must be reported. Eradi­cation of the virus has never been attempted. Prevention of early exposure to MDV, improved hygiene and early vaccination (at 1 day old) are commonly practised in Europe to control this disease in poultry.

INFECTIOUS LARYNGOTRACHEITIS

I nfectious laryngotracheitis (ILT) is a respiratory disease in chickens, peafowl and captive and released pheasants caused by a herpesvirus, subfamily Alphaherpesvirinae, genus Iltovirus, type species Gallid herpesvirus 1^⁸. ILT is endemic in some European countries and occasionally causes substantial losses.

Outbreaks of ILT may occur in all European countries with an intensive chicken industry. ILT is mainly seen in adult chickens and peafowl (Pavo cristatus) of both sexes. Some species of pheasants are highly susceptible. Guinea fowl (Numida meleagris), turkeys, quails, pigeons, ducks, geese, swans and passerines are resistant. Climate, season and temperature do not appear to affect the course of ILT. Subclinically infected chickens and farm-raised common pheasants ( Phasianus colchicus) released for hunting are the source for pheasants in the wild.

Birds of some species develop mild signs such as bilat­eral serous conjunctivitis and rhinitis. Other species display bilateral serous conjunctivitis, respiratory rales and leth­argy. Severe signs of disease are noted in other species that consist of serous-purulent conjunctivitis and rhinitis, swelling of eyelids, abnormal movements of the head (tor­ticollis) and lethargy that ends in high levels of mortality. It is noteworthy to recognize that the degree of clinical signs does not correlate with the genus of these birds.

Birds suffering from ILT excrete large amounts of blood- stained tracheal mucus containing ILT virus, thus facilitating transmission to other susceptible species. Sub- clinically infected birds may excrete ILT virus from con­junctiva and oral secretions and can also be a source of infection by direct and indirect contact. Vectors play no role. No evidence exists for vertical virus transmission.

Following nasal and conjunctival infection and infec­tion of the respiratory epithelium, mainly the trachea, virus multiplication occurs in mucosal cells of the respira­tory tract. If diseased birds do not die of suffocation by mucus in the respiratory tract, recovery is likely. Virus transport to the TG occurs early during infection. ILT virus remains in a latent stage in the TG. Convalescent and immunized birds produce antibodies.

Most birds that die of ILT are in good body condition. The main lesions are restricted to the upper respiratory tract. Extended haemorrhages occur in the nasal cavity, conjunctival sac, periorbital sinuses, trachea and primary bronchi. The epithelium is oedematous and frequently detached from the submucosa. Histopathologically, oedema of the respiratory mucosa, haemorrhages and mucoid exudates are present, and epithelial cells frequently, but not in all cases, contain INIB.

The incubation period following natural exposure to virulent strains in chickens is 6—12 days. The disease in chickens, peafowl and some species of pheasants has a rapid course. Initial clinical signs consist of general depression, a reduction in egg laying, reduced food and water intake and difficulties in breathing. These non-specific signs are fol­lowed by severe expiratory rales, nasal discharge of blood- tainted mucus, swollen infraorbital sinus and haemorrhagic tracheitis and increased or significant mortality. Milder strains of ILT virus cause respiratory depression, gasping and expectoration of bloody mucus(¹⁰9). The clinical signs in pheasants differ markedly between genera and species. There is considerable variation in clinical severity and mortality among the different species of pheasant found captive and free living (feral) in Europe.

The affected host species, the clinical signs and macro­scopic lesions are suggestive of ILT. Confirmation is obtained by histopathological detection of lesions in the respiratory tract and the presence of INIB. Virus isolation is performed in embryonated chicken eggs or in chicken kidney cell cultures from samples of the respiratory mucosa. Inoculated embryos display pox-like foci on the chorioal­lantoic membrane. Large syncytia are present in cell cul­tures. Several PCR are applied to identify field and vaccine viruses⁽¹¹⁰⁾.

Local outbreaks of ILT in domestic chickens are elimi­nated by culling. Total eradication of ILT appears possible owing to its narrow host range, the detection of latently infected birds by PCR and the sensitivity of the virus to chemical disinfectants, ultraviolet light, dryness and ele­vated temperatures.

Vaccination of adolescent and adult chickens with live attenuated vaccines is conducted in endemically infected areas by conjunctival installation (eye-drop method). Owing to the residual virulence of attenuated live vaccine viruses, care must be taken to avoid mixing vaccinated and unvaccinated chickens. Circumstantial evidence suggests that attenuated live ILT vaccines may regain their original virulence by serial passages in chickens. Also, the duration and protective capacity of ILT vaccines is relatively limited.

Great caution is required to prevent spread of vaccine- derived virus from chickens to pheasants and peafowl. Vaccines should never be used for any species of pheasant. Severe post-vaccinal reactions, including mortality, are likely in pheasants and peafowl.

Formal reporting of ILT to governmental authorities is not required. Legal regulations do not exist.

Exact data on the prevalence of ILT in domestic bird populations are not available. Domestic and free- living waterfowl, pigeons and passeriform birds are not sus­ceptible. Mammals, including humans, are completely resistant.

SMADEL'S DISEASE OF PIGEONS

SmadeLs disease, pigeon herpesvirus infection or ingluvitis of pigeons, is a contagious disease of predominantly young pigeons of all breeds (racing and fancy) of worldwide dis­tribution. Single cases are also diagnosed in feral pigeons (Columbia livia) and of other birds of the family Columbi- dae. Generally, the pigeon herpesvirus can affect all species of the family Columbidae. Infection without subsequent development of clinical signs are frequently observed.

The aetiologic agent of SmadeLs disease is a member of the order Herpesvirales, family Herpesviridae, subfamily Alphaherpesvirinae, genus Mardivirus, species pigeon her­pesvirus, Columbid herpesvirus 1, CoHV1(9⁸).

Exact data on the prevalence of SmadeLs disease is not available. However, numerous reports provide evidence for the presence of the pigeon herpesvirus in all European countries and many pigeon lofts. Pigeon herpesvirus has been detected in all breeds of domestic pigeons ( Columba livia f. domestica), feral pigeons, and other members of the family Columbidae. Young pigeons (squabs) are more sus­ceptible to disease than adults. Free-living but also domes­tic pigeons of various breeds are frequently co-infected with a large variety of other infectious agents, which increases the severity of the clinical course of SmadeLs disease. These agents include the pigeon circovirus, reovi- rus, adenovirus, Salmonella typhimurium var. copenhagen, Chlamydia psittaci, Trichomonas gallinae, yeasts and intes­tinal parasites. The disease in domestic pigeons occurs more frequently in the presence of environmental stressors. Feral pigeons in urban areas suffer frequently from chronic, immunosuppressive lead intoxication, which promotes the frequency and severity of the disease. Since the complete ban in Europe of the gasoline additive tetraethyl lead in 1997, the numbers of clinically overt forms of Smadel’s disease in pigeons has fallen. Infected domestic and feral pigeons provided as food for captive birds of prey may result in lethal infections of these birds, so this practice is not recommended, or, if done, falconers should remove the head and neck of pigeons before feeding. Chronically infected female and male breeding pigeons transmit the herpesvirus to their squabs by feeding regurgitated crop milk during the first weeks of life of the squabs. Contact during courtship, preening and mutual feeding of adult pairs during mating does not result in virus transmission. Egg transmission of the virus has not been recorded.

Ingested virus replicates in the oropharynx region, fol­lowed by short-term viraemia and virus multiplication in all internal organs. Squabs succumb as a result of severe epithelial lesions in the pharynx, oesophagus and crop and as a result of co-infections. The body development of clini­cally infected squabs is poor. Diphtheroid pharyngitis, oesophagitis and ingluvitis are prominent. The enlarged liver and spleen contain numerous pin- point white foci. Additional lesions caused by secondary infections are fre­quent. INIB are frequently present in epithelial cells of the upper digestive tract, liver and spleen. Surviving squabs develop antibodies.

I nfection by pigeon herpesvirus only rarely results in clinically overt forms of disease. Important co-factors such as poor hygiene, overcrowding in lofts, and concurrent infections increase the likelihood and severity of clinical disease. Adult pigeons usually do not show obvious signs of disease, whereas young squabs are depressed and anaemic with poor growth and poor plumage. Mortality can reach 30 to 50%.

A presumptive diagnosis is based on young age, clinical signs, gross and microscopic pathology. Virus isolation confirms the aetiologic diagnosis. Antibody assays (neu­tralization tests) have no value for the diagnosis of indi­vidual cases, because most healthy pigeons possess circulating antibodies¹-⁹⁹).

Squabs can be raised free of the infection if crop milk from virus-free parents is used as the only source of food. Chemotherapy using thymidine-kinase inhibitors (aciclo­vir or ganciclovir) has been tried with limited success. Vaccines are not available. The treatment of the prevailing secondary pathogens ameliorates the clinical course of the disease. Improved hygiene, including repeated cleans­ing and disinfection of the lofts, reduces the risk of exposure.

Pigeon herpesvirus 1 is not transmissible to mammals, including humans. As herpesvirus-infected pigeons are fre­quently concurrently infected by Chlamydia psittaci, special care is needed to prevent transmission of this zoonotic pathogen to people. There is also a potential risk of spill over of pigeon herpesvirus and also of chlamydia from domestic pigeons, to free-living columbiforms during racing competitions. However, definite proof for this assumption is not available.

INCLUSION BODY HEPATITIS OF OWLS, EAGLES AND FALCONS

Inclusion body hepatitis of owls (synonyms Hepatosplenitis infectiosa strigum (HSiS), inclusion body disease of owls), eagles and falcons (synonyms Inclusion body disease of falcons and eagles) are caused by unassigned viruses in the family Herpesviridae^”'.

The herpesviruses of owls, eagles and falcons are Owl herpesvirus 1 (hepatosplenitis virus or strigid herpesvirus 1), inclusion body disease virus of eagles (Eagle herpesvirus 1, acciptrid herpesvirus 1), and Inclusion body disease virus of falcons (Falconid herpesvirus 1 ), respectively.

Biological and virological properties of herpesvirus iso­lates from eagle owls, falcons and eagles are very similar, if not identical, to the Pigeon herpesvirus 1⁽¹¹¹). These viruses cause an almost identical gross and microscopic pathology(¹¹²), possess cross-reacting neutralizing antibod- ies⁽¹¹³⁾, form similar bands in restriction endonuclease pat- terns(¹¹⁴) and yield similar sequence data of a fragment of the highly conserved herpesviral DNA polymerase gene using degenerate PCR primers(¹¹¹). These herpesviruses are readily inactivated by chemical disinfectants and by expo­sure to ultraviolet light.

The domestic pigeon (Columba livia f. domestica) and the ubiquitous feral pigeon (Columba livia) are con­sidered the natural reservoirs¹-¹¹⁵). Detailed, contemporary data on the prevalence of herpesvirus in Strigi-, Falconi- and Accipitriformes in Europe is not available.

Owl herpesvirus 1 induces a highly lethal disease or subclinical infection in free-living and captive birds of: the order Strigiformes, family Strigidae, subfamilies Asioninae, genus Asio; subfamily Striginae, genera Strix, Megascopi, Otui, Bubo, Nyctea; subfamily Surniinae, genus Athene. Owls of all these genera present very similar gross, his­topathological and ultrastructural lesions in liver, spleen, bone marrow and oesophagus. Most of these viruses were isolated from dead birds that lived free or were maintained in breeding and rehabilitation centres in Europe(¹¹⁶).

There is evidence of different susceptibility of owl species to herpesvirus isolated from an eagle owl (B ubo bubo). Experimentally, the European barn owl ( Tyto alba) resisted infection, whereas nine identically infected owls of other species (five Asio otus, three Athene noctua, one Aego- lius funereus) died with typical lesions in the liver, spleen and bone marrow(¹¹⁷). Unfortunately, isolates from Euro­pean barn owls are not available for molecular characteri­zation. A herpesvirus was isolated from a barn owl but detailed characterization of this virus was not reported. Serologic studies in Germany showed neutralizing anti­bodies against an isolate from an eagle owl in 24 of 111 eagle owls but not in 61 barn owls(¹¹⁸). Falcon herpesvirus induces lesions similar to the owl herpesvirus in falcons, Falconiformes, genera Hierofalco, Chiquera, Aesalon, Tin- nunculus'^v,'. Eagle owl herpesvirus was isolated from eagles in Germany (Accipitriformes, genera Haliaeetus, Accipiter, Buteo^).

The viruses have been isolated from dead captive and wild birds and antibodies have been detected in live, apparently healthy birds, indicating infection in both young and adult hosts⁽¹²⁰). Infected European birds include eagle owl, long-eared owl (Asio otus), snowy owl (Nyctea scandiaca), little owl (Athene noctua), Tengmalms owl (Aegoluius funereus) and great horned owl (Bubo virgin- ianus). Tawny owl (Strix aluco^) and barn owl proved resist­ant to a high dose ofvirus during experimental infections ⁽¹¹⁷⁾. It has been noted that the susceptible species have yellow and orange irises, whereas the resistant ones have brown irises, although the relevance of this observation is not clear.

Circumstantial evidence suggests that raptors may become infected following consumption of infected pigeons and infected birds of prey. A German study yielded 12 strigid herpesviruses from 95 dead eagle owls and iden­tified virus neutralizing antibodies in 116 of 695 serum samples derived from apparently healthy adult eagle owls. Consequently, breeders of eagle owls were advised to use only virus- and antibody-negative birds to produce young for release.

Oral transmission via consumption of herpesvirus- infected pigeons is the most likely mode of infection. Horizontal transmission from bird to bird by sharing prey or by mutual aggression may also be possible. Evidence for vertical or egg transmission has not been published.

Experimental studies showed spread of HSiS virus in the body of infected birds to palate, the choana, oesopha­gus, liver, spleen, bone marrow, thymus, trachea, lung, intestines. Virus was not detected in brain, heart, proven­triculus or gizzard(¹¹⁷). Virus-positive cells included hepa­tocytes and Kupffer cells, cells of connective tissues, lymphatic cells and epithelial and mesenchymal cells⁽¹²¹⁾. The progression of the disease is associated with rapid virus multiplication and organ dysfunction¹-¹¹⁶). Dead owls were in relatively good body condition, suggesting a short course of the disease. The mortality rate is not known. Serologic data suggested that infected birds could recover.

Lesions in owls, falcons and eagles are uniform and consist of liver enlargement and small foci of necrosis in the liver, spleen and bone marrow. Intranuclear, eosinophilic inclusions in cells of these organs are characteristic.

Natural infection in owls was followed by an incubation period of approximately 1—2 weeks. However, after experi­mental infection general malaise and lethargy was noted after 3—4 days. The most frequent finding was sudden death⁽¹²¹). Some owls had millet-seed-sized yellow nodules in the buccal palate and oesophagus. Similar signs were noted in falcons and eagles(¹¹²).

The gross and microscopic lesions are characteristic. Virus isolation in cell cultures of avian origin form the basis for the aetiologic diagnosis. Virus neutralization tests are used to demonstrate antibodies that indicate previous infection¹-^113,122). PCR can be applied to detect the DNA polymerase gene of herpesviruses¹-¹¹¹).

Breeding of birds for subsequent release must be carried out using individuals that are free of virus and antibodies. Inactivated vaccines to protect healthy owls and falcons are of limited success. Cell-culture-adapted falcon herpes­virus live vaccine⁽¹²³⁾ did not provide effective protection against challenge with a homologous virulent virus obtained from a kestrel (Falco mexicanus). Treatment has no effect against the viral infection.

Release of herpesvirus-infected or antibody-positive birds must be avoided.

INCLUSION BODY DISEASE OF CRANES

Inclusion body disease of cranes, or hepatitis of cranes, is caused by Gruid herpesvirus 1 (GrHV-1), an unassigned virus in the family Herpesviridad⁹'¹. The virus caused fatal hepatitis in 12 grey-crowned cranes (Balearica pavonina) and in seven demoiselle cranes (A nthropoides virgo) in a safari park in Austria and additional losses in a zoo in Morbihan, France, in the winter of 1973—1974. In March and April 1979 a die-off in several crane species occurred in the International Crane Foundation, Baraboo, Wiscon­sin, USA; 18 out of 51 birds died suddenly. This appar­ently new disease in three locations has so far only been seen in captive cranes. It is not known whether any rela­tionship exists between outbreaks in Austria, France and the USA as the origins of the birds in the respective col­lections is not known. The currently known spectrum of susceptible cranes comprises the sandhill crane (Grus canadensis), red-crowned crane (Grus japonensis), hooded crane (Grus monacchus) and Stanley crane (Anthropoides paradisea). All diseased birds were mature and both sexes were involved. The maintenance of cranes in overcrowded enclosures may have facilitated virus spread. The disease is frequently lethal, but seropositive convalescent birds have been observed. Experimental infections provide evidence for susceptibility of white Pekin ducklings and adult coots (Fulica americana), whereas white leghorn chicks (Gallus gallus) and Muscovy ducks ( Cairina moschata) were resist­ant. Crane herpesviruses from the outbreaks in Austria and France cross-react with a herpesvirus isolated from a bob­white quail (Colinus virginianus)(¹²⁴) and yield identical bands in restriction enzyme analysis¹-¹¹⁴).

Crane herpesvirus is excreted with faeces. Transmission via eggs was ruled out. The infective viral dose is unknown. It is likely that infection occurs by the oral route, but it is not clear if it results in an initial virus replication in the upper digestive tract. Postmortem data provide evidence for viraemia with subsequent dissemination in internal organs. The most prominent lesions are seen in liver and spleen, which are enlarged and with numerous grey foci. Necrosis is seen in the gastrointestinal tract, thymus and bursa of Fabricius(¹²⁵). Enteric lesions are occasionally observed. Histologically, numerous intranuclear inclusions are present in hepatocytes.

The course of the disease is rapid: birds succumb within 2 days. Although most infected cranes succumb, recovery and seroconversion is possible. Initial signs consist of depression, anorexia, lack of preening, enteritis and sitting with eyes closed(^126,127).

The aetiological diagnosis is obtained by virus isolation in the cell culture yielding cytopathic effects, followed by characterization of the virus. Monoclonal antibodies that enable specific detection of crane virus by immuno­fluorescence and antibody assays in a competitive ELISA have been produced¹-¹²⁸).

Separation of newly acquired birds in quarantine and serological monitoring should reduce the risk of introduc­tion and subsequent spread. There is no vaccine or effec­tive treatment.

So far, spread of crane virus from infected premises to free- living birds or white Pekin ducklings has not been reported. The role of the quail virus as a possible source of infection for cranes has been proposed but the relation­ship, if there is one, is not clear.

HERPESVIRUS INFECTIONS IN PASSERIFORMES

Although the order Passeriformes contains approximately half of all avian species, the isolation and characterization of herpesviruses from these birds are rarely described and there are no published reports in free-living European pas­serine birds. The few publications refer to captive pet birds such as canaries (Serinus canaria f. domestica'f¹²’'. In Austria(¹³⁰), Switzerland(¹³¹), Canada(¹³²) and Illinois, USA(¹³³), lethal diseases that are associated with conjunc­tivitis and respiratory distress were seen in gouldian finches (Chloebia gouldiae). Herpesvirus isolations were obtained from healthy appearing sharp-tailed mannikin (Lonchura striata), bronze mannikin ( Spermestes cucullatus), common cardinal (Cardinalis cardinalis) and zebra finch ( Taeniopy- gea guttata)(¹²⁹). Also, a herpesvirus was isolated from a disease outbreak in newly imported superb starlings (Lam- protornis superbus)(¹³⁴) that is genetically related to a psit- tacid herpesvirus of the genotype 1⁽¹³⁵). So far, there are no publications providing evidence for lateral spread of these exotic passerine herpesviruses found in captive passerines, to endemic wild European species.

HERPESVIRUS INFECTION OF STORKS

The white stork (Ciconia ciconia) is a common bird in many parts of Europe. The causes of decline and recovery of white and black storks (Ciconia nigra) are carefully documented, and dead birds are usually comprehensively examined. Herpesviruses (Ciconiid herpesvirus 1) have been isolated and tentatively assigned to the family Her- pesviridae(⁹ⁱ'¹. The herpesvirus causes necrotic lesions usually in the liver and spleen(¹³⁶). Additionally, haemor­rhagic enteritis was described in Spain⁽¹³⁷⁾. Follow-up studies in rehabilitation centres in Germany provide evi­dence for a long-lasting, possibly life-long, cell-associated viraemia in disabled but otherwise normal adult white storks. It appears that storks can live with such viraemia for prolonged times, frequently for years, and produce healthy offspring(¹³⁸). The stork herpesvirus is antigeneti- cally unrelated to any of the other avian herpesviruses.

PACHECO'S disease

In 1931 Genesio Pacheco and Otto Bier(¹³⁹) described, for the first time in great detail, a highly lethal disease in Brazilian large parrots and differentiated this apparently new disease from psittacosis.

The causative virus of Pacheco’s disease (PD) is desig­nated Psittacid herpesvirus 1 (PsHV1), it is classified as a member of the family Herpesviridae, and stands as an unas­signed virus in the subfamily A lphaherpesvirinae, genus Iltovirus(^,'¹. At least four, possibly more, major genotypes are known, with each genotype including two to four vari­ants. Six serotypes are recognized that correspond well to genotypes(¹4⁰).

PD affects many parrot species originating from several continents. The disease is seen mainly in Amazon parrots (Amazona spp., African grey parrots (Psittacus erithacus), macaws (Ara spp.) and cockatoos (Cacatua spp.). South American conures (Aratinga spp. and Pyrrhura spp.) are less frequently affected, but they often survive following infection and develop a carrier state associated with faecal virus excretion, which is important for lateral spread of virus.

In recent decades, free-living, sustainable populations of some parrot species, mainly parakeets of the Genus Psit- tacula spp., that have escaped from private collections have established in several Northern European and Mediterra­nean countries. Birds of this genus are not endemic in Europe but are susceptible to PD virus. So far, cases of PD in these free-living parrot populations have not been pub­lished, but PD does occur in captive psittacines in Europe. Natural transmission to endemic avian species in Europe has not been recorded.

Psittacine birds acquire the infection by oropharyngeal uptake of virus from contaminated food and water but also by coprophagia. Initial virus multiplication occurs in the upper respiratory and digestive tracts, followed by viraemia and spread of the virus to almost all the internal organs. In chronic cases, death follows as a result of emaciation, dehydration and dysfunction of multiple organs.

Postmortem findings in acute cases consist of good body condition (owing to the short duration of illness) and enteritis, enlarged liver and spleen with focal necrosis, enlarged ureters filled with urates. No prominent gross lesions are detectable in peracute disease forms.

Clinical signs of PD develop after an incubation period of 1 week and consist initially of lethargy, anorexia, ruffled feathers, closed eyelids and occasionally respiratory signs. During further progression of the disease greenish-yellow liquid droppings with larger amounts of urates are seen. Rarely, CNS disorders develop. The clinical course of the disease before death is a matter of days.

The diagnosis is based on virus isolation in cell cultures or by PCR. Virus differentiation is done by geno- and serotyping. Recovered birds have antibodies that can be differentiated into serotypes by neutralization tests. In his­topathology, INIB can aid in the diagnosis.

All psittacine herpesviruses are sensitive to chemical disinfectants and radiation by ultraviolet light. Improved hygiene is recommended to reduce the risks of spread within bird collections. Only PD-negative birds should attend exhibitions. As psittacine herpesviruses consist of several geno- and serotypes, autogenous vaccines are very effective to prevent spread and further losses in affected bird collections. These vaccines — specific for each bird collection — are produced from cell culture- grown virus that is purified and inactivated by formalin and supple­mented with potent adjuvants. Vaccinated birds develop neutralizing antibodies.

All non-psittacine birds, mammals and humans are resistant to infection by psittacine herpesviruses. Legal restrictions do not exist.

REFERENCES

1. McGeoch, D.J., Dolan, A. & Ralph, A. C. Toward a comprehensive phylogeny for mammalian and avian herpesviruses. Journal of Virol­ogy. 2000;74:10401-6.

2. McGeoch, D.J., Cook, S., Dolan, A., Jamieson, F.E. & Telford, E.A.R. Molecular phylogeny and evolutionary timescale for the family of mammalian herpesviruses. Journal of Molecular Biology. 1995;247: 443-58.

3. ISI Web of Knowledge. Available online at: http://science. thomsonreuters.com. [accessed 29 February 2012].

4. Taller, D., Bilek, A., Revilla-Fernandez, S. et al. Diagnosis of Aujesz- ky's disease in a dog in Austria. Wiener Tierarztliche Monatsschrift. 2006;93:62-7.

5. Leschnik, M.W., Benetka, V., Url, A. et al. Virale enzephalitiden beim hund in Osterreich: diagnostische und epidemiologische aspecte. Wiener Tierarztliche Monatsschrift. 2008;95:190-9.

6. Cay, A.B. & Letellier, C. Isolation of Aujeszkys disease virus from two hunting dogs in Belgium after hunting wild boars. Vlaams Dierg Tijdschr. 2009;78:194-5.

7. Muller, T., Klupp, B.G., Freuling, C. et al. Characterization of pseu­dorabies virus of wild boar origin from Europe. Epidemiology and Infection. 2010;138:1590-600.

8. Wright, J.C. &Thawley, D.G. Role of the raccoon in the transmission of pseudorabies: a field and laboratory investigation. American Journal of Veterinary Research. 1980;41:581-3.

9. Vicente, J., Ruiz-Fons, F., Vidal, D. et al. Serosurvey of Aujeszky^,s disease virus infection in European wild boar in Spain. Veterinary Record. 2005;156:408-12.

10. Acevedo, P, Vicente, J., Hofle, U. et al. Estimation of European wild boar relative abundance and aggregation: a novel method in epide­miological risk assessment. Epidemiology and Infection. 2007;135:519- 27.

11. Romero, C.H., Meade, PN., Shultz, J.E. et al. Venereal transmission of pseudorabies viruses indigenous to feral swine. Journal of Wildlife Diseases. 2001;37:289-96.

12. Schulze, C., Hlinak, A., Wohlsein, P. et al. Spontaneous Aujeszkys disease (pseudorabies) in European wild boars (Sus scrofa) in the federal state of Brandenburg, Germany. Berliner undMunchener Tier- arztliche WochenschriJi. 2010;123:359-64.

13. Gortazar, C., Vicente, J., Fierro, Y. et al. Natural Aujeszkys disease in a Spanish wild boar population. Annals of the New York Academy of Sciences. 2002;969:210-2.

14. Muller, T.F., Teuffert, J., Zellmer, R. et al. Experimental infection of European wild boars and domestic pigs with pseudorabies viruses with differing virulence. American Journal of Veterinary Research. 2001;62:252-8.

15. Marcaccini, A., Lopez Pena, M., Quiroga, M.I. et al. Pseudorabies virus infection in mink: a host-specific pathogenesis. Veterinary Immu­nology Immunopathology. 2008;124:264-73.

16. Kimman, T.G. & van Oirschot, J.T. Pathology of Aujeszkys disease in mink. Veterinary Pathology. 1986;23:303-9.

17. Muller, T., Teuffert, J., Staubach, C. et al. Long-term studies on maternal immunity for Aujeszkys disease and classical swine fever in wild boar piglets. Journal of Veterinary Medicine B. 2005;52:432- 6.

18. Ruiz-Fons, F., Vidal, D., Hofle, U. et al. Aujeszkys disease virus infec­tion patterns in European wild boar. Veterinary Microbiology. 2007;120:241-50.

19. Ruiz-Fons, F., Rodriguez, O., Mateu, E. et al. Antibody response of wild boar (Sus scroJa) piglets vaccinated against Aujeszkys disease virus. Veterinary Record. 2008;162:484-5.

20. Anusz, Z., Szweda, W, Popko, J. et al. Is Aujeszkys disease a zoonosis? Przeglad Epidemiologiczny. 1992;46:181-6.

21. Heuschele, W.P. & Reid, H. W. Malignant catarrhal fever. In: Wil­liams, E.S. & Baker, I.K., (eds). Infectious Diseases of Wild Mammals, 3rd edn. Ames, Iowa: Iowa State University Press; 2001; pp. 157­64.

22. Reid, H.W, Buxton, D., Pow, I. & Finlayson, J. Isolation and char­acterisation of lymphoblastoid cells from cattle and deer affected with ‘sheep-associated’ malignant catarrhal fever. Research in Veterinary Science. 1989;47:90-6.

23. Russell, G.C., Stewart, J.P & Haig, D.M. Malignant catarrhal fever: a review. Veterinary Journal. 2009;179:423-35.

24. Milne, I., Wright, F., Rowe, G., Marshal, D.F., Husmeier, D. & McGuire. G. TOPALi: Software for automatic identification of recombinant sequences within DNA multiple alignments. Bioinfor­matics. 2004;20:1806-7.

25. Plowright, W, Ferris, R.D. & Scott, G.R. Blue wildebeest and the aetiological agent of bovine catarrhal fever. Nature. 1960;188: 1167-9.

26. Vikoren, T., Li, H., Lillehaug, A., Jonassen, C.M., Bockerman, I. & Handeland, K. Malignant catarrhal fever in free-ranging cervids asso­ciated with OvHV-2 and CpHV-2 DNA. Journal of Wildlife Diseases. 2006;42:797-807.

27. Neimanis, A.S., Hill, J.E., Jardine, C.M. & Bollinger, T.K. Sheep- associated malignant catarrhal fever in free-ranging moose (Alces alces} in Saskatchewan, Canada. Journal of Wildlife Diseases. 2009;45: 213-7.

28. Schultheiss, PC., Van Campen, H., Spraker, T.R., Bishop, C., Wolfe, L. & Podell, B. Malignant catarrhal fever associated with ovine herpervirus-2 in free-ranging mule deer in Colorado. Journal of Wild­life Diseasei. 2007;43:533-7.

29. Loken, T., Aleksandersen, M., Reid, H.W & Pow, I. Malignant catarrhal fever caused by ovine herpesvirus-2 in pigs in Norway. Veterinary Record. 1998;143:464-7.

30. Buxton, D., Reid, H.W, Finlayson, J. & Pow, I. Pathogenesis of sheep-associated malignant catarrhal fever in rabbits. Research in Vet­erinary Science. 1984;36:205-11.

31. Baxter, S.I.F., Wujono, P, Pow, I. & Reid, H.W. Identification of ovine herpesvirus-2 infection in sheep. Archives of Virology. 1997;142:823-31.

32. Li, H., Taus, N.S., Jones, C., Murphy, B., Evermann, J.F. & Crawford, T.B. A devastating outbreak of malignant catarrhal fever in a bison feedlot. Journal of Veterinary Diagnostic Investigation. 2006;18:119-23.

33. Herring, A.J., Reid, H.W, Inglis, N. & Pow, I. Immunoblotting analysis of the reaction of wildebeest, sheep and cattle sera with the structural antigens of Alcelaphine herpesvirus-1 (malignant catarrhal fever virus). Veterinary Microbiology. 1989;19:205-15.

34. Das Neves, C.G. Cervid Herpesvirus 2 Infection in Reindeer in Norway. Oslo: Norwegian School of Veterinary Science; 2009.

35. Thiry, J., Muylkens, B. & Thiry, E. Infectious bovine rhinotracheitis and the epidemiological role of the other ruminant species. Hungarian Veterinary Journai 2008;130:116-23.

36. Keel, M.K., Patterson, J.G., Noon, T.H., Bradley, G.A. & Collins, J.K. Caprine herpesvirus- 2 in association with naturally occurring malignant catarrhal fever in captive sika deer ( Cervus nippon). Journal of Veterinary Diagnostic Investigation. 2003;15:179-83.

37. Thiry, E., Vercouter, M., Dubuisson, J. et al. Serological survey of herpesvirus infections in wild ruminants of France and Belgium. Journal of Wildlife Diseases. 1988;24:268-73.

38. Tempesta, M., Camero, M., Sciorsci, R.L. et al. Experimental infec­tion of goats at different stages of pregnancy with caprine herpesvirus 1. Comparative Immunology, Microbiology and Infectious Diseasei. 2004;27:25-32.

39. Mettler, F., Engels, M., Wild, P. & Bivetti, A. Herpesvirus-infektion bei Zieklein in der Schweiz. Schweizer Archiv fur Tierheilkunde. 1979;121:655-62.

40. Grewal, A.S. & Wells, R. Vulvovaginitis of goats due to a herpesvirus. Austrailian Veterinary Journal. 1986;63:79-82.

41. Buddle, B.M., Pfeffer, A., Cole, D.J., Pulford, H.D. & Ralston, M.J. A caprine pneumonia outbreak associated with caprine herpesvirus and Pasteurella haemolytica respiratory infections. New Zealand Veteri­nary Journal. 1990;38:28-31.

42. Engels, M., Palatini, M., Metzler, A.E., Probst, U., Kihm, U. & Ackermann, M. Interactions of bovine and caprine herpesviruses with the natural and the foreign hosts. Veterinary Microbiology. 1992;33: 69-78.

43. Tryland, M., Das Neves, C.G., Sunde, M. & Mork, T. Cervid her­pesvirus 2, the primary agent in an outbreak of infectious keratocon­junctivitis in semi domesticated reindeer. Jo urnal of Clinical Microbiology. 2009;47:3707-13.

44. Das Neves, C.G., Rimstad, E., Tryland, M. et al. Cervid herpesvirus 2 causes respiratory and fetal infections in semidomesticated reindeer. Journal of Clinical Microbiology. 2009;47:1309-13.

45. Inglis, D.M., Bowie, J.M., Allan, M.J. & Nettleton, P.F. Ocular disease in red deer calves associated with a herpesvirus- infection. Veterinary Record. 1983;113:182-3.

46. Thiry, J., Widen, F., Gregoire, F., Linden, A., Belak, S. & Thiry, E. Isolation and characterisation of a ruminant alphaherpesvirus closely related to bovine herpesvirus 1 in a free-ranging red deer. BMC Vet­erinary Research. 2007;3:26.

47. Ek-Kommonen, C., Pelkonen, S. & Nettleton, P.F. Isolation of a herpesvirus serologically related to bovine herpesvirus 1 from a reindeer (Rangifer tarandus). Acta Veterinaria Scandinavica. 1986;27: 299-301.

48. Das Neves, C.G., Roth, S., Rimstad, E., Thiry, E. & Tryland, M. Cervid herpesvirus 2 infection in reindeer: a review. Veterinary Micro­biology. 2010;143:70-80.

49. Vanderplasschen, A., Bublot, M., Pastoret, PP &Thiry, E. Restriction maps of the DNA of cervid herpesvirus 1 and cervid herpesvirus 2, two viruses related to bovine herpesvirus 1. Archives of Virology. 1993;128:379-88.

50. Ek-Kommonen, C., Veijalainen, P, Rantala, M. & Neuvonen, E. Neutralizing antibodies to bovine herpesvirus 1 in reindeer. Acta Veterinaria Scandinavica. 1982;23:565-9.

51. Das Neves, C.G., Thiry, J., Skjerve, E. et al. Alphaherpesvirus infec­tions in semidomesticated reindeer: a cross-sectional serological study. Veterinary Microbiology. 2009;139:262-9.

52. The Danish Veterinary and Food Administration. Report on the Animal Health Situation in Greenland 1999. The Danish Veterinary and Food Administration; 1999; Contract No.: 15-3-2009. Available online at: http://www.fodevarestyrelsen.dk/fdir/Pub/2000640/ rapport1.htm [accessed 15 March 2012].

53. Lillehaug, A., Vikoren, T., Larsen, I.L., Akerstedt, J., Tharaldsen, J. & Handeland, K. Antibodies to ruminant alpha-herpesviruses and pestiviruses in Norwegian cervids. Journal of Wildlife Diseases. 2003;39:779-86.

54. Stuen, S., Krogsrud, J., Hyllseth, B. & Tyler, N.J.C. Serosurvey of three virus infections in reindeer in northern Norway and Svalbard. Rangifer. 1993;13:215-9.

55. Reid, H.W., Nettleton, PF., Pow, I. & Sinclair, J.A. Experimental infection of red deer (Cervus elaphus) and cattle with a herpesvirus isolated from red deer. Veterinary Record. 1986;118:156-8.

56. Das Neves, C.G., Mork, T., Godfroid, J. et al. Experimental infection of reindeer with cervid herpesvirus 2. Clinical and Vaccine Immunol­ogy. 2009;16:1758-65.

57. Das Neves, C.G., Mork, T., Thiry, J. et al. Cervid herpesvirus 2 experimentally reactivated in reindeer can produce generalized viremia and abortion. Virus Research. 2009;145:321-8.

58. Tryland, M., Das Neves, C.G., Sunde, M. & Mork, T Cervid her­pesvirus 2, the primary agent in an outbreak of infectious keratocon­junctivitis in semidomesticated reindeer. Journal of Clinical Microbiology. 2009;47:3707-13.

59. Nettleton, PF., Ek-Kommonen, C., Tanskanen, R., Reid, H.W., Sin­clair, J.A. & Herring, J.A. Studies on the epidemiology and patho­genesis oh alphaherpesvirus from red deer (Cervus elaphus) and reindeer (Rangifer tarandus). In: Reid HW, (ed.). The Management and Health of Farmed Deer. London: Kluwer Academic Publishers; 1988. pp. 143-8.

60. Osterhaus, A.D.M.E., Yang, H., Spijkers, H.E.M., Groen, J., Teppema, J.S. & van Steenis, G. The isolation and partial characteri­zation of a highly pathogenic herpesvirus from the harbour seal (Phoca vitulina). Archives of Virology. 1985;86:239-51.

61. Goldstein, T., Mazet, J.A., Lowenstine, L.J. et al. Tissue distribution of phocine herpesvirus-1 (PhHV-1) in infected harbour seals (Phoca vitulina) from the central Californian coast and a comparison of diagnostic methods. Journal of Comparative Pathology. 2005;133:175- 83.

62. Borst, G.H.A., Walvoort, H.C., Reijnders, PJ.H., van der Kamp, J.S. & Osterhaus, A.D.M.E. An outbreak of a herpesvirus infection in harbor seals (Phoca vitulina) Journal of Wildlife Diseases. 1986;22:1- 6.

63. Gulland, F.M., Lowenstine, L.J., Lapointe, J.M., Spraker, T & King, D.P Herpesvirus infection in stranded Pacific harbor seals of coastal California. Journal of Wildlife Diseases. 1997;33:450-8.

64. Blanchard, T.W., Santiago, N.T., Lipscomb, T.P., Garber, R.L., McFee, WE. & Knowles, S. Two novel alphaherpesviruses associated with fatal disseminated infections in Atlantic bottlenose dolphins. Journal of Wildlife Diseases. 2001;37:297-305.

65. Arbelo, M., Sierra, E., Esperon, F. et al. Herpesvirus infection with severe lymphoid necrosis affecting a beaked whale stranded in the Canary Islands. Diseases of Aquatic Organisms. 2010;89:261^.

66. Soto, S., Gonzalez, B., Willoughby, K. et al. Systemic herpesvirus and morbillivirus co-infection in a striped dolphin (Stenella coeruleoalba) Journal of Comparative Pathology 2012;146:269-73.

67. Kennedy, S., Lindstedt, I.J., McAliskey, M.M., McConnell, S.A. & McCullough S.J. Herpesviral encephalitis in a harbor porpoise (Phoc- oena phocoend). Journal of Zoo and Wildlife Medicine 1992;23:374-9.

68. Martineau, D., Lagace, A., Higgins, R., Armstrong, D. & Shugart, L.R. Pathology of stranded Beluga whales (Delphinapterus leucas) from the St. Lawrence Estuary, Quebec, Canada. Journal of Comparative Pathology. 1988;98:287-311.

69. Baker, J.R. Causes of mortality and parasites and incidental lesions in dolphins and whales from British waters. Veterinary Record. 1992;130: 569-72.

70. Baker, J.R. & A.R. Martin. Causes of mortality and parasites and incidental lesions in harbour porpoises (Phocoena phocoencι) from British waters. Veterinary Record. 1992;130:554-8.

71. Manire, C.A., Smolarek, KA., Romero, C.H., Kinsel, M.J., Clauss, TM. & Byrd, L. Proliferative dermatitis associated with a novel alphaherpesvirus in an Atlantic bottlenose dolphin (Tursiops trunca- tus). Journal of Zoo and Wildlife Medicine. 2006;37:174-81.

72. van Elk, C.E., van de Bildt, M.W, de Jong, A.A., Osterhaus, A.D. & Kuiken,T. Herpesvirus in bottlenose dolphins (Tursiops truncatus): cultivation, epidemiology, and associated pathology. Journal of Wild­life Diseases. 2009;45:895-906.

73. Smolarek Benson, K.A., Manire, C.A., Ewing, R.Y. et al. Identifica­tion of novel alpha- and gammaherpesviruses from cutaneous and mucosal lesions of dolphins and whales. Journal ofVirological Methods. 2006;136:261-6.

74. Saliki, J.T., Cooper, E.J., Rotstein, D.S. et al. A novel gammaherpes­virus associated with genital lesions in a Blainvilles beaked whale (Mesoplo don dens iro str is). Journal of Wildlife Diseases. 2006;42: 142-8.

75. Daoust, PY., Taylor, R.G. & Greenlaw, B.L. Herpesvirus in botryo- mycotic lesions from a harp seal (Phoca groenlandica) Veterinary Pathology. 1994;31:385-7.

76. Lebich, M., Harder, TC., Frey, H.R., Visser, I.K., Osterhaus, A.D. & Liess, B. Comparative immunological characterization of type­specific and conserved B- cell epitopes of pinniped, felid and canid herpesviruses. Archives of Virology. 1994;136:335^7.

77. Martina, B.E., Jensen, TH., van de Bildt, M.W, Harder, TC. & Osterhaus, A.D. Variations in the severity of phocid herpesvirus type 1 infections with age in grey seals and harbour seals. Veterinary Record. 2002;150:572-5.

78. Martina, B.E.E., Verjans, G.M., Harder,T.C. et al. Seal gammaher­pesviruses: identification, characterisation and epidemiology. Virus Research. 2003;94:25-31.

79. Esperon, F., Fernandez, A. & Sanchez-Vizcaino, J.M. Herpes simplex­like infection in a bottlenose dolphin stranded in the Canary Islands. Diseases of Aquatic Organisms. 2008;81:73-6.

80. Belliere, E.N., Esperon, F., Arbelo, M., Munoz, M.J., Fernandez, A. & Sanchez-Vizcaino, J.M. Presence of herpesvirus in striped dolphins stranded during the cetacean morbillivirus epizootic along the Mediter­ranean Spanish coast in 2007. Archives of Virology. 2010;155:1307-11.

81. Maness, H.T., Nollens, H.H., Jensen, E.D. et al. Phylogenetic analysis

of marine mammal herpesviruses. Veterinary Microbiology.

2011;149:23-9.

82. Miyoshi, K., Nishida, S., Sone, E. et al. Molecular identification of novel alpha- and gammaherpesviruses from cetaceans stranded on Japanese coasts. Zoological Science. 2011;28:126-33.

83. Goldstein, T., Mazet, J.A., Gulland, F.M. et al. The transmission of phocine herpesvirus-1 in rehabilitating and free-ranging Pacific harbor seals (Phoca vitulina) in California. Veterinary Microbiology. 2004;103:131-41.

84. Greenwood, A.G., Harrison, R.J. & Whitting, H.W Functional and pathological aspects of the skin of marine mammals. In: Harrison RJ, (ed.). Functional Anatomy of Marine Mammals. London: Academic Press; 1974; pp. 73-111.

85. King, D.P, Lie, A.R., Goldstein, T. et al. Humoral immune responses to phocine herpesvirus-1 in Pacific harbor seals (Phoca vitulina rich- ardsii) during an outbreak of clinical disease. Veterinary Microbiology. 2001;80:1-8.

86. Barr, B., J.L. Dunn, M.D. Daniel & A. Banford. Herpes-like viral dermatitis in a Beluga Whale ( Delphinapterus leucas). Journal of Wild­life Diseases. 1989;25:608-11.

87. Martina, B.E., van de Bildt, M.W, Kuiken, T., van Amerongen, G. & Osterhaus, A.D. Immunogenicity and efficacy of recombinant subunit vaccines against phocid herpesvirus type 1. Vaccine. 2003;21:2433-40.

88. Stack, M.J., Higgins, R.J., Challoner, D.J. & Gregory, M.W Herpes­virus in the liver of a hedgehog (Erinaceus europaeus). Veterinary Record. 1990;127:620-1.

89. Widen, F., Gavier-Widen, D., Nikiila, T. & Morner, T. Fatal herpes­virus infection in a hedgehog (Erinaceus europaeus). Veterinary Record. 1996;139:237-8.

90. Labrut, S., Hoby, S., Kappeler, A., Ryser, M-P. & Robert, N. Fatal herpesvirus encephalitis in a hedgehog (Erinaceus europaeus). Euro­pean Association of Zoo and Wildlife Veterinarians (EAZWV) 6th Scientific Meeting; May 24-28, Budapest, Hungary; 2006.

91. Wibbelt, G., Kurth, A., Yasmum, N. et al. Discovery of herpesviruses in bats. Journal of General Virology. 2007;88:2651-5.

92. Molnar, V, Janoska M., Harrach, B. et al. Detection of a novel bat gammaherpesvirus in Hungary. Acta Veterinaria Hungarica. 2008;56:529-38.

93. King, D.P, Mutukwa, N., Lesellier, S., Cheeseman, C., Chambers, M.A. & Banks, M. Detection of Mustelid Herpesvirus-1 Infected European Badgers (Meles meles) in the British Isles. Journal of Wildlife Diseases. 2004;40:99-102.

94. Becker, S.D., Bennett, M., Stewart, J.P & Hurst, J.L. Serological survey of virus infection among wild house mice (Mus domesticus) in the UK. Laboratory Animals. 2007;41:229-38.

95. Blasdell, K., McCracken, C., Morris, A. et al. The wood mouse is a natural host for Murid herpes-virus 4. Journal of General Virology. 2003;84:111-3.

96. Leutenegger, C.M., Hofmann-Lehmann, R., Riols C. et al. Viral infections in free-living populations of the European wild cat. Journal of Wildlife Diseases. 1999;35:678-86.

97. Daniels, M.J., Golder, M.C., Jarrett, O. & MacDonald, D.W Feline viruses in wildcats from Scotland. Journal of Wildlife Diseases. 1999;35:121-4.

98. Davison, A.J., Eberle, R., Ehlers, B. et al. The order Herpesvirales. Archives of Virology. 2009;154:171-7.

99. Kaleta, E.F. Herpesviruses of free-living and pet birds. In: Dufour- Zavala, L., Swayne, D.E., Glisson, J.R. et al. (eds). A Laboratory Manual for the Isolation, Identification, and Characterization of Avian Pathogens, 5th edn. Athens, Ga: American Association of Avian Pathologists; 2008; pp. 110-5.

100. de Thoisy, B., Lavergne, A., Semelin, J.E. et al. Outbreaks of disease possibly due to a natural avian herpesvirus infection in a colony of young magnificent frigatebirds (Fregata magniftcens) in French Guiana. Journal of Wildlife Diseases. 2009;45:802-7.

101. Li, Y., Bing, H., Ma, X., W. et al. Molecular characterization of the genome of duck enteritis virus. Virology. 2009;391:151-61.

102. Kaleta, E.F., Kuczka, A., Kuhnhold, A. et al. Outbreak of duck plague (duck viral enteritis) in numerous species of captive ducks and geese in temporal conjunction with enforced biosecurity (in-house keeping) due to the threat of avian influenza A virus of the subtype H5N1. German Veterinary JournaS 2007;114:3-11.

103. Salguero, FJ., Sanches-Cordon, PJ., Nunez, A. & Gomez-Villandos, J.C. Histopathological and ultrastructural changes associated with her­pesvirus infection of waterfowl. Avian Pathology. 2002;31:133^0.

104. Goldberg, D.R., Yuill, T.M. & Burgess, E.C. Mortality from duck plague virus in immunosuppressed adult mallard ducks. Journal of Wildlife Diseases. 1990;26:299-306.

105. Plummer, PJ., Alefantis, T., Kaplan, S., O'Connell, P, Shawky, S. & Schat, K.A. Detection of duck enteritis virus by polymerase chain reaction. Avian Diseases. 1998;42:554-64.

106. Jansen, J. The interference phenomenon in the development of resist­ance against duck plague. Journal of Comparative Pathology and Thera­peutics. 1964;74:3-7.

107. Sandhu, T.S. & Metwally S.A. Duck virus enteritis (duck plague). In: Saif, Y.M., Fadly, A.M., Glisson, J.R., McDougald, L.R., Noland,

L. K. & Swayne, D.E. (eds). Diseases of Poultry, 12th edn. Ames, Iowa: Blackwell Publishing; 2008; pp. 384-93.

108. Kaleta, E.F. Vermehrung, Interferenz und Interferoninduktion aviarer Herpesvirusarten. Beitrag zur Schutzimpfung gegen die Mareksche Krankheit. Zentralblatt fur VeterinUrmedizin B. 1975;24:406-75.

109. Guy, J.S. & Garcia, M. Laryngotracheitis. In: Saif, Y.M., Fadly, A.

M., Glisson, J.R., McDougald, L.R., Nolan, L.K. & Swayne, D.E., (eds). Diseases of Poultry. 12th edn. Ames, Iowa: Iowa State University Press; 2008; pp. 137-52.

110. Tripathy, D.N. & Garcia, M. Infectious laryngotracheitis. In: Dufour- Zavala, L., Swayne, D.E., Glisson, J.R. et al. (eds). A Laboratory Manual for the Isolation, Identification, and Characterization of Avian Pathogens, 5th edn. Athens, Ga: American Association of Avian Pathologists; 2008; pp. 94-8.

111. Gailbreath, K.L. & Oaks, J.L. Herpesviral inclusion body disease in owls and falcons is caused by the pigeon herpesvirus ( Columbid her­pes-virus 1 ). Journal of Wildlife Diseases. 2008;44:427-33.

112. Ramis, A., Mao, N., Pumarola, M., Fondevila, D. & Ferrer, L. Herpesvirus hepatitis in two eagles in Spain. Avian Diseases. 1994;38:197-200.

113. Kaleta, E.F., Heffels, U., Neumann, U. & Mikami, T. Serological differentiation of 14 avian herpesviruses by plaque reduction tests in cell cultures. Proceedings of the Second International Symposium of Veterinary Laboratory Diagnosticians'; June 24-26; Bern, Switzerland. 1980; pp. 38-41.

114. Gunther, B.M.F., Klupp, B.G., Gravendyck, M., Lohr, J.E., Met- tenleiter, T.C. & Kaleta, E.F. Comparison of the genomes of 15 avian herpesvirus isolates by restrictioon endonuclease analysis. Avian Pathology. 1997;26:305-16.

115. Vindevogel, H. & Pastoret, PP Herpesvirus infections of pigeons and wild birds. In: McFerran, J.B. & McNulty, M.S. (eds). Virus Infections of Birds. Amsterdam: Elsevier Science Publishers B.V; 1993; pp. 91-106.

116. Kaleta, E.F. & Docherty, D.E. Avian herpesviruses. In: Thomas, N.J., Hunter, D.B. & Atkinson, C.T. (eds). Infectious Diseases ofWild Birds. Ames, Iowa: Blackwell Publishing; 2007; pp. 63-86.

117. Burtscher, H. & Sibalin, M. Herpesvirus strigis: host spectrum and distribution in infected owls. Journal of Wildlife Diseases. 1975;11: 164-9.

118. Kaleta, E.F. & Druner, K. Hepatosplenitis infectiosa strigum und andere Krankheiten der Greifvogel und Eulen, 11. Fortschritte der Veterinarmedizin. 1976; pp. 173-80.

119. Heidenreich, M. Greifvogel: Krankheiten, Haltung, Zucht. Berlin: Blackwell Wissenschafts-Verlag; 1995.

120. Heinrichs, M.A. Herpesvirus-induzierte Infektionen und Krankheiten bei nicht domestizierten Vogelarten - eine vergleichende Liter- aturstudie: Universitat Giessen; 1992.

121. Burtscher, H. & Url, A. Die virusbedingte Hepatosplenitis strigum. 3. Mitteilung: Fluoreszenz-serologische Untersuchungen am Brutei. Zentralblatt fur Veterinarmedizin. 1970;B17:765-77.

122. Zsivanovits, P, Forbes, N.A., Zvonar, L.T. et al. Investigation into the seroprevalence of falcon herpesvirus antibodies in raptors in the UK using virus neutralization tests and different herpesvirus isolates. Avian Pathology. 2004;33:599-604.

123. Wernery, U., Wernery, R. & Kinne, J. Production of a falcon herpes­virus vaccine. Berliner und Munchener Tierarztliche Wochenschrift. 1999;112:339-44.

124. Forster, S., Chastel, C. & Kaleta, E.F. Crane hepatitis herpesviruses. Zentralblattfur Veterinarmedizin. 1989;B36:433^1.

125. Schuh, J.C.L., Sileo, L., Siegfried, L.M. & Yuill, T.M. Inclusion body disease of cranes: comparison of pathologic findings in cranes with acquired versus experimentally in duced disease. Journal of the Ameri­can Veterinary Medical Association. 1986;189:993-6.

126. Burtscher, H. & Grunberg, W. Herpesvirus-Hepatitis bei Kranichen (Aves: Gruidae). I. Pathomorphologische Befunde. Zentralblatt fur Veterin a rmedizin. 1979;B26:561-9.

127. Docherty, D.E. & Romaine, R.I. Inclusion body disease of cranes: a serological follow-up to the 1978 die-off. Avian Diseases. 1983;27:830- 5.

128. Letchworth, G.J., Fishel, J.R. & Hansen, W A monoclonal antibody to inclusion body disease of cranes virus enabling specific immuno­histochemistry and competitive ELISA. Avian Diseases. 1997; 41:808-16.

129. Blumenstein, V Isolierung und biologische Eigenschaften von sechs neuen Herpesviren aus verschiedenen Sperlingsvogeln (Passeriformes). University Giessen; 1993.

130. Schonbauer, M. & Kohler, H. Uber eine Virusinfektion bei Prachtfinken (Estrildidae). Kleintierpraxis. 1982;27:149-52.

131. von Rotz, A., Rubel, A., Mettler, F. & Hoop, R. Letal verlaufende Herpesvirus-infection bei Gouldsamadinen (Chloebia gouldinae [Gould]). Sch∙weizer Archiv fur Tierheilkunde. 1984;126:651-8.

132. Wellehan, J.F.X., Gagea, M., Smith, D.A., Taylor, WM., Berhane, Y. & Bienzle, D. Characterization of a herpesvirus assiciated with tra­cheitis in Gouldian finches (Erythruna [Chloebia] gouldiae). Journal of Clinical Microbiology. 2003;41:4054-7.

133. Paulman, A., Lichtensteiger, C.A. & Kohrt, L.J. Outbreak of herpes­viral conjunctivitis and respiratory disease in Gouldian finches. Vet­erinary Pathology. 2006;43:963-70.

134. Gravendyck, M. Isolierung und biologische Eigenschaften eines neuen Herpesvirus aus einem Dreifarbenglanzstar (Lamprotornis superbus Ruppel, 1845) sowie Versuche zur Differenzierung von Her- pesviren aus Passeriformes durch Restriktionsendo- nukleasen: Uni­versity Giessen; 1996.

135. Tomaszewski, E.K., Gravendyck, M., Kaleta, E.F. & Phalen, D.N. Genetic characterization of a herpesvirus isolate from a superb starling (Lamprotornis superbus) as a psittacid herpesvirus genotype 1. Avian Diseases. 2004;48:212^.

136. Kaleta, E.F. & Kummerfeld, N. Herpesviruses and Newcastle diease viruses in white storks (Ciconia ciconia). Avian Pathology. 1983;12: 347-52.

137. Gomez-Villamandos, J.C., Hervas, J., Salguero, FJ., Quevedo, M.A., Aguilar, J.M. & Mozos, E. Haemorrhagic enteritis associated with herpesvirus in storks. Avian Pathology. 1998;27:229-36.

138. Kaleta, E.F. & Kummerfeld, N. Persistent viraemia of a cell-associated herpesvirus in white storks (Ciconia ciconios). Avian Pathology. 1986;15:447-53.

139. Pacheco, G. & Bier, O. Epizootia em papagaios no Brasil e suas relacoes com a psittacose. Archivos do Instituto Biologico de Defesa Agricola e Animat. 1931;4:89-120.

140. Tomaszewski, E.K., Kaleta, E.F. & Phalen, D.N. Molecular phylogeny of the psittacid herpesviruses causing Pacheco's disease: correlation of genotype with phenotypic expression. Journal of Virology. 2003;77: 11260-7.