inclusion body rhinitis
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IdentityTop of page
Preferred Scientific Name
- inclusion body rhinitis
International Common Names
- English: cytomegalic inclusion body disease in swine; cytomegalic inclusion disease; inclusion body rhinitis of pigs; inclusion body rhinitis of swine; inclusion-body rhinitis of swine; infectious bovine pustular vulvovaginitis, ipv, herpesvirus 1; infectious bovine rhinotracheitis, ibr, herpesvirus 1; infectious bovine rhinotracheitis, ibr, in goats; infectious pustular vulvovaginitis; mucopurulant vaginitis; porcine cytomegalic inclusion disease; porcine cytomegalovirus infection; porcine cytomegalovirus, suid herpesvirus-2 in swine; vaginitis and vulvitis
Pathogen/sTop of page suid herpesvirus 2
OverviewTop of page
Done (1955) first characterized a particular type of the historically common disease of rhinitis in pigs, calling it 'inclusion body rhinitis' (IBR). The transmissibility of the disease was first demonstrated by Bakos et al. (1960). Subsequent reports of the disease indicated it had a global distribution. Histolologically, IBR was characterized by the presence of large, basophilic intranuclear inclusion bodies in the mucosal epithelial cells of porcine turbinate tissue. The evidence of Cameron-Stephen (1961) and Corner et al. (1964), indicated that the agent of IBR was also responsible for a range of other clinical signs including lower respiratory tract disease and reproductive failure. Clinical signs of IBR were largely restricted to piglets up to 3 weeks of age. In gestating pigs the disease was manifested by the production of mummified, stillborn or weak, stunted piglets. Affected piglets displayed a range of respiratory signs including coughing, snuffling and sneezing in addition to catarrhal rhinitis, anorexia, lethargy, paresis, stunting and sudden death. In some cases a more disseminated form of the disease was seen in piglets and very occasionally in older pigs; histology carried out post-mortem revealed inclusion bodies in multiple organ systems (Hartley and Done, 1963; Corner et al., 1964).
Observations of field cases showed similarities between IBR and the clinical signs of salivary gland virus infection of infant children, puppies and guinea pigs (Done, 1955; Hartley and Done, 1963). The viral aetiology of inclusion body rhinitis, first suggested by Done (1955) was supported by Duncan et al. (1965), who compared the electron microscopic appearance in tissues collected from cases of IBR with that of other cytomegaloviruses. Further confirmation came from the work of L’Ecuyer and Corner (1966) and Booth et al. (1967). The 'cytomegaly' was caused by swelling of the mitochondria and dilatation of the endoplasmic reticulum, Golgi apparatus and nucleus due to the presence of large numbers of virus particles. This feature was common to the murine, guinea pig and human cytomegaloviruses (Valicek et al., 1969).
In common with all herpesviruses, PCMV has been shown to establish a reactivatible latency in selected cells. Although not proven, long term isolation of PCMV from lung macrophages indicates that latency may reside in a lymphoid site (Edington et al., 1976a; Narita et al., 1985).
The virus was first propagated in vitro in pig lung cell cultures (L’Ecuyer and Corner, 1966). Cultivation of PCMV in vitro was improved by the use of pig lung macrophages (Watt et al., 1973) and a cell line derived from porcine fallopian tube cells (Bouillant et al., 1975). These developments facilitated the development of diagnostic assays for cytolological examination (Watt et al., 1973) and for antigen detection (direct fluorescent antibody test; FAT) and specific antibody detection (indirect fluorescent antibody test; IFAT) by Plowright et al. (1976). Significant elements of the pathogenesis of PCMV were determined by the observation that the virus can cross the placenta in gestating pigs (Rac, 1961; L’Ecuyer et al., 1972; Edington et al., 1977).
Despite having a high herd prevalence and a global distribution, PCMV-associated overt clinical disease is not common. This became more obvious when it was shown that the agent of IBR was not a common denominator in the aetiology of the widespread disease of atrophic rhinitis (AR). There was however, some evidence that mucosal damage caused by IBR predisposed to colonization by the bacterial pathogens implicated as the principal causative agents of atrophic rhinitis (Mitchell and Corner, 1958; Corner et al., 1964; Marquardt, 1979; Rondhuis et al., 1980; Rademacherova, 1981). The low rate of diagnosis of PCMV is also affected by the pig industry and regulatory bodies effectively accepting to 'live with the disease' and by the absence of rapid, robust and sensitive assays for virus and antibody detection.
More recently the profile of PCMV has been raised by the potential for the use of pig organs in xenotransplantation. The ubiquitous and persistent nature of PCMV infection, coupled with its ability to cross the placenta raised concerns over the potential transmission of this virus via pig organs transplanted to highly immunosuppressed human recipients. This has led to increased efforts to develop improved diagnostic techniques and antiviral therapies aimed at the generation and maintenance of PCMV-free donor herds.
The cytomegaloviruses are thought to have evolved from a common herpesvirus progenitor approximately 100 million years ago. Thereafter the evidence indicates that the evolution of herpesviruses has followed that of the hosts in a process of co-speciation (McGeoch et al., 1995). Based on cultural and disease characteristics, the presence of basophilic, intranuclear inclusion bodies and electron micropscopic studies, the virus was classified as a member of the subfamily Betaherpesvirinae (Roizman, 1982).
Relatively little is known about the molecular biology of PCMV, however, the recent renewed interest in the virus, fuelled by xenotransplantation issues has led to the elucidation of the nucleic acid and putative protein sequences of two genes in a 7.2 kb segment of the unique long part of the genome (Widen et al., 2000).
In summary, PCMV appears to be an extremely well adapted virus which has infected most of the world’s domestic pig population by efficient vertical and horizontal transmission and which causes relatively little overt clinical disease.
Host AnimalsTop of page
Hosts/Species AffectedTop of page
The pig is the definitive and only known host of PCMV. A characteristic of beta-herpesviruses is host specificity and this is further reflected in the failure to cultivate the virus on anything other than cells of porcine origin. Although all age groups may be infected with PCMV, overt clinical signs are almost entirely restricted to piglets infected in utero or neonatally.
Systems AffectedTop of page blood and circulatory system diseases of pigs
digestive diseases of pigs
mammary gland diseases of pigs
multisystemic diseases of pigs
nervous system diseases of pigs
reproductive diseases of pigs
respiratory diseases of pigs
DistributionTop of page
Inclusion body rhinitis of swine was first reported as a disease entity in England in 1955 (Done, 1955). Within eight years of this report the disease was reported in eleven other countries in Europe, North America and the Antipodes. A difficult virus to cultivate in vitro, porcine cytomegalovirus (PCMV) has been isolated in relatively few countries but it is thought to exist at a high prevalence in most if not all national domestic pig herds. In many countries in which it has been isolated it has been described as ubiquitous in distribution and to date, no country has eradicated or demonstrated freedom from the virus. The situation in feral swine and wild boar is unknown.
Distribution TableTop of page
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.Last updated: 10 Jan 2020
PathologyTop of page
In common with postmortem findings, the preponderance of histopathological lesions in epithelial tissues compared to reticulo-endothelial tissue lesions has been presented by Edington et al. (1976b) as a discrimination between the localized or disseminated infection of older animals and the generalized infection of the foetus or neonate.
In older pigs, no macroscopic lesions are found apart from a mild to severe rhinitis. In foetal or neonatal pigs, postmortem findings show mild to severe petechiation involving the heart, kidneys, intestine, lungs, lymph nodes and meninges. Petechial lesions are usually most severe in the kidney, resulting in the appearance ranging from speckling to solid black or purple. Pulmonary congestion, consolidation and oedema, with hydrothorax and a 5-fold enlargement of bronchial and mediastinal lymph nodes are often seen. The ventral areas of the apical, cardiac and diagphragmatic lobes of the lung often show dark or purple discolouration. There is a mild catarrhal rhinitis with a mucus nasal exudate and congestion of nasal mucosa. In addition to pulmonary and occasional peritoneal oedema, the jaw, neck and the tarsal joints are a common site of oedema in acutely infected piglets. Anaemia is a common finding with packed cell volumes recorded at between 10% and 12 % and haemaglobin levels of 4-6 gramme % (Corner et al., 1964; L’Ecuyer et al. 1972; Edington et al., 1976a, b; Orr et al., 1988).
The most consistent histopathological finding is the presence of intranuclear cytomegalic inclusions in the glandular tissue of the nasal turbinate mucosa, acinar and duct cells of the lachrymal, Harderian and salivary glands and tubular epithelial cells and medullary interstitial tissue in the kidney. In addition inflammatory lesions in the absence of inclusions may be observed in the brain, where a focal microgliosis lung and occasionally in other tissues. In salivary glands there may be evidence of acinar necrosis and fusion of cells to form syncitia with hyaline or vacuolated eosinophilic cytoplasm (Corner et al., 1964; Kelly, 1967; Edington et al., 1976b).
In an experimental reproduction of the generalized form of the disease Edington et al. (1976b) observed inclusion bodies in multiple tissues and organ systems including lung, small intestine, liver, kidney, gonad, tonsil, bronchial lymph node, mesenteric lymph node, spleen, bone marrow, CNS, mandibular lymph node, adrenal gland and thyroid. In the lungs the interlobular septa and many of the alveoli in the lung were filled with a fluid exudate in which macrophages were common and lymphocytes rare. These areas contained many alveoli which were collapsed or were filled with intact and lysed erthrocytes. In the kidney the inclusions were present in medullary capillaries but were most common in the cortex in peripheral areas of differentiating renal tissue and in the glomerular capillary endothelium. In the spleen, large numbers of inclusion bodies were seen in free-lying mononuclear cells. In the liver the involvement of hepatic cells was shown by focal necrosis and haemorrhage, leading to disruption of hepatic lobules.
In the brain, inclusions in capillary endothelium were most commonly seen in the choroid plexus or in vessels lying outside the cerebral cortex. In a number of piglets a multifocal encephalitis was observed consisting of astrocytes and microglial cells in association with perivascular cuffing. No inclusion bodies were seen in these lesions.
In some pigs with generalized infection, evidence of of haematopoiesis was indicated in liver and spleen and by or the presence of megakaryocyte giant cells and nucleated erthrocytes (Corner et al., 1964; Ohlinger, 1989).
Ultrastructural changes in infected cells have been observed by electron microscope (EM) studies. The cytomegalic state of the cell was a result of the dilation of the endoplasmic reticulum and Golgi apparatus and a swelling of the mitochondria. Large aggregates of virus particles were observed within cytoplasmic vesicles (Duncan et al., 1965; Valicek et al., 1969; Narita et al., 1987). Further EM observations are described in the section 'Disease Course'.
DiagnosisTop of page Clinical Diagnosis
The clinical diagnosis of PCMV infection is based on the observation of mummified, stillborn or weak litters and/or the signs of rhinitis and other lower respiratory tract disease amongst suckling and neonatal pigs. In affected piglets a commonly observed sign is the presence of black discolouration around the eyes caused by conjunctival exudate. None of these signs is sufficiently specific for a diagnosis to be made in the absence of laboratory confirmation.
There are no pathognomonic macroscopic external lesions associated with PCMV infection. Lesions observed postmortem are indicated under 'Pathology'.
Agents causing porcine reproductive failure and respiratory tract disease should be considered. The differential diagnosis should therefore include virus infections caused by Aujeszky’s disease (AD; Pseudorabies), porcine respiratory and reproductive syndrome (Yoon et al., 1996), and the so-called SMEDI viruses diseases including porcine parvovirus and the abortigenic enteroviruses. More recently, the post weaning multi-systemic wasting syndrome (PMWS) has now to be considered in the differential diagnosis of porcine reproductive failure.
The agents of porcine atrophic rhinitis (Bordetella bronchiseptica and Pasturella spp.) and adenovirus disease should be considered in cases of rhinitis, the former particularly in cases where snout abnormality is observed.
Betaherpesviruses by nature are slow growing, fastidious in terms of host and cell type and do not produce high titres of virus in vitro. Accordingly the first method of diagnosis was a microscopic examination for the characteristic intranuclear basophilic inclusion bodies in stained, wax embedded sections of nasal tissue. In the absence of in vitro culture systems, this diagnostic tool remained one of the most effective ways of diagnosing PCMV infection. A slight modification of this method was introduced by Done (1958) whereby the use of Giemsa or methylene blue stained nasal scrapings enabled a diagnosis in the live pig.
A more complete diagnosis required the isolation of PCMV from the tissues of infected pigs. Propagation of PCMV in a range of primary cell cultures was reported by L’Ecuyer and Corner (1966), Booth et al. (1967) and Watt et al. (1973). In these trials only pig lung macrophage (PLM) cultures were shown to be susceptible to passage with PCMV. This observation was used by Plowright et al. (1976) to develop methods for the detection, isolation and assay of PCMV, in particular the fluorescent antibody test (FAT). In this method virus was isolated by inoculation of tissue homogenates onto PLM cells grown on flying coverslips. PCVM was detected by use of a PCMV antiserum conjugated to a fluorescent dye. This technique took 3-5 days but was a substantial reduction on the 10-14 days normally required for the development of a detectable cytopathic effect in the PLMs. The method represents the gold standard for PCMV isolation from tissues and swab samples.
The PLMs were used for inoculation with nasal swabs, tissue homogenates or lung washings. The FAT on cryostat sections also enabled confirmation of the infecting virus from tissue samples. Recommended tissues for either virus isolation or FAT were nasal mucosa, lung (including PLMs) and kidney. It was noted that following storage of a carcass or tissues at 4°C, the virus was detectable by FAT for 24 hours after viral infectivity for tissue cultures had ceased (Edington, 1992).
Disadvantages of the use of PLMs for PCMV isolation were stated as: variation in quality and sensitivity, pre-existing viral infection, short lifespan (10-12 days), low toxicity threshold, low yield from piglets and the fact that piglets had to be killed for each supply of PLMs, in contrast to many other cell culture systems (Watt et al., 1973; Plowright et al., 1976). The difficulty in obtaining PCMV free piglets also reduces the facility of this method, but despite the demonstration of PCMV replication in swine testis, salivary gland and fallopian tube cells (L’Ecuyer and Corner, 1966; Booth et al., 1967; Bouillant et al., 1975) PLMs remain the most susceptible to PCMV infection. For virus isolation Valicek et al. (1977) showed that virus could be isolated and demonstrated within 1 or 2 days by direct culture of PLMs obtained by lavage from diseased pigs. This compared favourably with the several weeks required for the same result by the repeated passage of nasal mucosa suspensions on PLMs.
A method for EM examination of nasal mucosa following negative staining by the method of Brenner and Horner (1959) was described by Scott et al. (1973). This had equal sensitivity and specificity and yet was more rapid than examination of either stained, wax-embedded sections of nasal mucosa or the method using stained scrapings of nasal mucusa (Done, 1958). The value of the EM approach was further developed and confirmed by Valicek et al. (1977).
In the last few years the generation of DNA sequence data has allowed the development of polymerase chain reaction (PCR) techniques for the detection of PCMV DNA in pig tissues (Widen et al., 1999). By adoption of a nested PCR approach this technique has proven more sensitive than the existing gold standard of virus isolation/FAT on PLM cultures.
The PLM cultures were also adapted by Plowright et al. (1976) for the development of an indirect FAT (IFAT) for detection and titration of specific antibody by use of an anti-pig IgG serum (goat or rabbit) conjugated to a fluorescent dye. Assaf et al. (1982), using PCMV antigen produced by sonication of infected porcine fallopian tube (PFT) cells, developed an ELISA for PCMV antibody assay. In comparison with the IFAT the ELISA was equally as specific and slightly more sensitive. A similar comparative exercise was carried out by Tajima et al. (1993) using antigen derived from a hypotonic buffer and detergent lysate of PCMV-infected PFT cells. Included in this comparative exercise was a serum neutralization test (SNT) developed earlier by Kawamura et al. (1992). The ELISA was slightly more sensitive than the IFAT and both assays were markedly more sensitive than the SNT. Additional advantages of the ELISA included a machine-read result and a greater potential volume throughput. The ELISA was further refined to include a protocol using whole blood collected onto filter paper in place of serum. A good correlation of ELISA result was observed between the two sample matrices (Tajima et al., 1994).
Serological surveys are valuable to establish the range and prevalence of a disease. However, in a situation where a herd has a high seroprevalence the diagnostic value of serology is less clear. In these cases it is necessary to look at antibody levels in a cohort of animals or preferably to use paired acute and convalescent phase samples to detect a rise in antibody titre. A standard rule is that a fourfold rise in titre between the acute phase sample and the convalescent phase sample collected two to three weeks later, is an indication of active infection or reactivation.
Immune competence in the pig foetus begins at 50-60 days gestation (Bourne et al., 1974; Bachmann et al., 1975). Whether this produces an increased ability to resist infection with PCMV is not supported by the observation that most transplacental transmission occurs in mid to late pregnancy (L’Ecuyer et al., 1972; Edington et al., 1977).
In the naïve pig antibody is detectable by IFAT three weeks post infection and persists for the life of the pig. In piglets born of immune sows, maternal antibody decays by 5-6 weeks of age. A rise in antibody level is then observed, consistent with infection by circulating virus following mixing of litters at weaning (Plowright et al., 1976). In pigs infected congenitally or neonatally in a closed herd system the antibody level may remain low through the fattening period (Edington, 1989).
List of Symptoms/SignsTop of page
|Cardiovascular Signs / Tachycardia, rapid pulse, high heart rate||Sign|
|Digestive Signs / Anorexia, loss or decreased appetite, not nursing, off feed||Pigs:All Stages||Diagnosis|
|Digestive Signs / Diarrhoea||Pigs:Piglet||Sign|
|Digestive Signs / Excessive salivation, frothing at the mouth, ptyalism||Sign|
|Digestive Signs / Oral mucosal ulcers, vesicles, plaques, pustules, erosions, tears||Sign|
|Digestive Signs / Tongue ulcers, vesicles, erosions, sores, blisters, cuts, tears||Sign|
|Digestive Signs / Vomiting or regurgitation, emesis||Pigs:Piglet||Sign|
|General Signs / Discomfort, restlessness in birds||Pigs:Piglet||Sign|
|General Signs / Fever, pyrexia, hyperthermia||Sign|
|General Signs / Fever, pyrexia, hyperthermia||Sign|
|General Signs / Forelimb swelling, mass in fore leg joint and / or non-joint area||Pigs:Piglet||Sign|
|General Signs / Forelimb weakness, paresis, paralysis front leg||Pigs:Piglet||Sign|
|General Signs / Generalized weakness, paresis, paralysis||Pigs:Piglet||Sign|
|General Signs / Head, face, ears, jaw, nose, nasal, swelling, mass||Pigs:Piglet||Sign|
|General Signs / Hindlimb swelling, mass in hind leg joint and / or non-joint area||Pigs:Piglet||Sign|
|General Signs / Hypothermia, low temperature||Sign|
|General Signs / Lack of growth or weight gain, retarded, stunted growth||Pigs:Piglet||Diagnosis|
|General Signs / Neck swelling, mass cervical region||Pigs:Piglet||Sign|
|General Signs / Pale mucous membranes or skin, anemia||Pigs:Piglet||Sign|
|General Signs / Paraparesis, weakness, paralysis both hind limbs||Pigs:Piglet||Sign|
|General Signs / Petechiae or ecchymoses, bruises, ecchymosis||Sign|
|General Signs / Reluctant to move, refusal to move||Pigs:Piglet||Sign|
|General Signs / Sudden death, found dead||Pigs:Piglet||Diagnosis|
|General Signs / Swelling mass, vulva, clitoris||Sign|
|General Signs / Tetraparesis, weakness, paralysis all four limbs||Pigs:Piglet||Sign|
|General Signs / Thoracic swelling, mass, thorax, chest, ribs, sternum||Pigs:Piglet||Sign|
|General Signs / Trembling, shivering, fasciculations, chilling||Sign|
|General Signs / Underweight, poor condition, thin, emaciated, unthriftiness, ill thrift||Pigs:Piglet||Sign|
|General Signs / Weakness of one hindlimb, paresis paralysis rear leg||Pigs:Piglet||Sign|
|General Signs / Weight loss||Pigs:All Stages||Sign|
|Nervous Signs / Dullness, depression, lethargy, depressed, lethargic, listless||Sign|
|Nervous Signs / Dullness, depression, lethargy, depressed, lethargic, listless||Sign|
|Nervous Signs / Head tilt||Pigs:Piglet||Sign|
|Ophthalmology Signs / Blepharospasm||Sign|
|Ophthalmology Signs / Chemosis, conjunctival, scleral edema, swelling||Sign|
|Ophthalmology Signs / Conjunctival, scleral, injection, abnormal vasculature||Sign|
|Ophthalmology Signs / Conjunctival, scleral, papules||Sign|
|Ophthalmology Signs / Conjunctival, scleral, redness||Pigs:Piglet||Sign|
|Ophthalmology Signs / Corneal edema, opacity||Sign|
|Ophthalmology Signs / Corneal neovascularization, pannus||Sign|
|Ophthalmology Signs / Lacrimation, tearing, serous ocular discharge, watery eyes||Pigs:Piglet||Diagnosis|
|Ophthalmology Signs / Photophobia||Sign|
|Ophthalmology Signs / Purulent discharge from eye||Sign|
|Pain / Discomfort Signs / Pain, penis||Sign|
|Pain / Discomfort Signs / Pain, vulva, vagina||Sign|
|Reproductive Signs / Abortion or weak newborns, stillbirth||Pigs:Sow||Diagnosis|
|Reproductive Signs / Agalactia, decreased, absent milk production||Pigs:Sow||Sign|
|Reproductive Signs / Female infertility, repeat breeder||Sign|
|Reproductive Signs / Mucous discharge, vulvar, vaginal||Sign|
|Reproductive Signs / Mummy, mummified fetus||Pigs:Sow||Diagnosis|
|Reproductive Signs / Papule, pustule, vesicle, ulcer penis or prepuce||Sign|
|Reproductive Signs / Purulent discharge, vulvar, vaginal||Sign|
|Reproductive Signs / Vaginal or cervical ulcers, vesicles, erosions, tears, papules, pustules||Sign|
|Reproductive Signs / Vulval ulcers, vesicles, erosions, tears, cuts, pustules, papules||Sign|
|Respiratory Signs / Abnormal breath odor, foul odor mouth||Sign|
|Respiratory Signs / Abnormal breathing sounds of the upper airway, airflow obstruction, stertor, snoring||Sign|
|Respiratory Signs / Abnormal lung or pleural sounds, rales, crackles, wheezes, friction rubs||Sign|
|Respiratory Signs / Coughing, coughs||Pigs:Piglet||Sign|
|Respiratory Signs / Dyspnea, difficult, open mouth breathing, grunt, gasping||Pigs:Piglet||Diagnosis|
|Respiratory Signs / Epistaxis, nosebleed, nasal haemorrhage, bleeding||Sign|
|Respiratory Signs / Increased respiratory rate, polypnea, tachypnea, hyperpnea||Pigs:Piglet||Sign|
|Respiratory Signs / Mucoid nasal discharge, serous, watery||Pigs:Piglet||Diagnosis|
|Respiratory Signs / Nasal mucosal ulcers, vesicles, erosions, cuts, tears, papules, pustules||Sign|
|Respiratory Signs / Purulent nasal discharge||Pigs:Piglet||Diagnosis|
|Respiratory Signs / Sneezing, sneeze||Pigs:Piglet||Diagnosis|
|Skin / Integumentary Signs / Skin crusts, scabs||Sign|
|Skin / Integumentary Signs / Skin edema||Sign|
|Urinary Signs / Dysuria, difficult urination, stranguria||Sign|
|Urinary Signs / Increased frequency of urination, pollakiuria||Sign|
Disease CourseTop of page
The primary sites of virus entry and replication are the mucosal surfaces of the nasal mucous glands or the lachrymal or Harderian gland (Edington, 1992). At 14-21 days post infection (dpi), infective virus may be detected in the blood. The viraemia, most commonly transient to 2-3 days in duration in most pigs, was detected from 5-19 dpi in gnotobiotic pigs infected at one day of age, indicating an age-dependent susceptibility (Goodwin and Whittlestone, 1967; Edington et al., 1976a). Virus is shed for 1 to 3 weeks predominantly from the nasal mucosa during the acute phase of infection; titres of virus were shown to exceed 106.9 TCID50/g of nasal mucosa and 102.5 TCID50/g of kidney tissue (Edington et al., 1976b). Excretion has been demonstrated for up to 10 weeks in a commercial pig herd (Plowright et al., 1976). Narita et al. (1987) showed that the release of virus from nasal mucosa was a two-stage process beginning with the desquamation of the nasal tubuloalveolar gland cells followed by destruction of the plasma membrane and release of the cytoplasmic contents.
In most pigs above 3 weeks of age PCMV may be disseminated by haematogenous spread with a predilection for replication in epithelial cells. In addition to the primary sites of replication, the virus replicates in the epithelial cells of renal tubules. Occasionally the virus may be recovered from the epididymis, from hepatocytes, duodenal epithelium and the mucous glands of the oesophagus (Edington et al., 1976b). In pigs infected congenitally or neonatally, generalized lesions are seen consequent to infection of lymphoid tissues, including sinusoidal and capillary endothelial cells, pulmonary macrophages and lymphocytes, spleen, liver, renal tubules and in other organs (Edington et al., 1976b; Narita et al., 1987).
Transplacental infection appears to occur most often in mid to late gestation (L’Ecuyer et al., 1972; Edington et al., 1977). In producing transplacental infection experimentally, Edington et al. (1977) showed that the majority of dead foetuses had died 4-6 weeks post-inoculation of the sow, irrespective of the time of gestation of the inoculation. The lack of infected foci in the placenta indicates that the virus probably traverses the placenta in infected reticulo-endothelial cells such as macrophages. Whether the litter is infected as a result of multiple transits or by passage from foetus to foetus following a single infection point is not clear, but up to 30% of foetuses may be infected in the last two trimesters of gestation.(Edington et al., 1977). Infection in utero during mid to late term commonly results in a generalised infection manifested by the production of mummified, stillborn or weak, runted, live-born piglets, a number of which may die suddenly (L’Ecuyer et al., 1972; Edington et al., 1977). Virus may be recovered from many organs including brain tissue (Edington et al., 1976b; Edington et al., 1988a).
Minimal infection of the alveolar cells in the early embryo was observed, indicating a difference in tropism from the predominantly endotheliotropic behaviour in the foetus and neonatal pig (Edington et al., 1977, 1988b).
A sub-acute infection may result in reduced appetite and respiratory problems persisting for up to one month. In acutely affected litters mortality in is usually around 10% but may be as high as 50% with morbidity close to 100% (Ohlinger, 1989).
In convalescent pigs the virus remains in a dormant or latent form in certain cells. Although not proven, the evidence from long term isolation, reactivation studies and comparison with other cytomegaloviruses indicates that the site(s) of virus latency are lymphoid cells. Alveolar macrophages and cells of the salivary gland are strongly suspected as the sites of PCMV latency. The distribution of virus and lesions following reactivation appears similar to that observed following experimental or natural infection (Edington et al., 1976a; Narita et al., 1985).
EpidemiologyTop of page
The available evidence suggests that PCMV has a worldwide distribution and a high herd prevalence in domestic pigs (Edington, 1989) to the extent that freedom from PCMV is not requested as a condition of international trade in pigs. The situation in feral pigs or wild boar is unknown.
Although all ages of pigs may be infected and excrete virus following contact with PCMV, the clinical signs of infection are inapparent in most pigs greater than 3 weeks of age.
Following experimental infection of gnotobiotic pigs, virus was recovered in nasal secretions for a maximum period of 32 days (Edington et al., 1976a). Although nasal secretions undoubtedly represent the major source of transmissable virus, PCMV has also been demonstrated in phayngeal and ocular secretions, in urine and in the cervical fluids of PCMV-naïve, pregnant sows (Plowright et al., 1976; Edington et al., 1977). The detection of PCMV in cervical fluids 30-35 days post-infection was considered more likely to be of foetal origin, based on the estimated time of foetal death in utero, than of maternal origin (Edington et al., 1977). PCMV has not been isolated from or detected in porcine semen, but the isolation of PCMV from the epididymis and testis (Booth et al., 1967; Shirai et al., 1985) indicates that semen may be a possible route of virus transmission. This view is countered to some extent by the failure to detect replication or shedding of virus from the male reproductive tract following experimental intranasal or intrapreputial inoculation of PCMV (Edington et al., 1988b). The impact of urinogenital excretion of PCMV on the epidemiology of the disease is unclear. Transplacental transmission of PCMV was suggested by Rac (1961) and demonstrated experimentally by L’Ecuyer et al. (1972) and Edington et al. (1977). However, incidence of transplacental transmission in seropositive sows may not be high (Marquardt, 1981); all 87 piglets delivered by hysterotomy from 6 seropositive sows were all shown to be free of PCMV and viral antibody (once the maternal antibody had waned).
In a study of the behaviour of PCMV in commercial pig herds, Plowright et al. (1976) showed that nasal excretion of virus amongst piglets was first detected at 3 weeks and highest between 5 and 8 weeks of age. This period was associated with the decline in maternal antibody and the post weaning mixing of the litters of young pigs. The major route of virus dissemination was considered to be via nasal secretions during sneezing. The source of the infecting virus may have been small numbers of congenitally infected piglets (L’Ecuyer et al., 1972; Edington et al., 1977) or other breeding stock, including newly introduced gilts, sows or boars. Although PCMV can be reactivated experimentally (Edington et al., 1976a), the frequency and impact of spontaneous or natural stress-associated reactivation of latent PCMV is unknown. Edington et al. (1976a) suggest it may act as the major source of PCMV epizootics through the contact of recently moved breeding stock with piglets or by the precipitation of transplacental infection of foetuses.
Impact: EconomicTop of page
The economic impact of the disease is generally considered to be modest and freedom from the virus is rarely demanded for either internal or international trade. The evidence for the prevalence of PCMV is based on a relatively small sample size, largely as a result of the absence of high volume antibody assays. In all cases where surveys have been done however, a very high prevalence has been observed (Kanitz and Woodruff, 1976; Plowright et al., 1976; Rademacherova, 1981; Assaf et al., 1982; Tajima et al., 1993).
Zoonoses and Food SafetyTop of page
Porcine cytomegalovirus displays the characteristics of the Betaherpesvirinae in that it has a highly restricted host range. The presence of the virus has been detected only in domestic pigs and it poses no zoonotic risk. The risks associated with transmission to humans following xenotransplantation of a PCMV infected pig organ are unknown.
Prevention and ControlTop of page
No vaccines have been developed for use against PCMV.
Natural outbreaks of PCMV-induced disease most often resolve without intervention (Cameron-Stephen, 1961; Corner et al., 1964), and in many cases no particular treatment is necessary. Husbandry practices that reduce stress levels, particularly surrounding the introduction of new stock, and reduction of litter mixing, may all reduce the severity and frequency of PCMV disease. The ubiquitous nature of PCMV predicates that epizootics of damaging, acute clinical disease are most likely in closed, high health status herds which have a high proportion of PCMV-naïve stock (Orr et al., 1988).
A study by Marquardt (1981) indicates that transplacental transmission is not always a common event. The 87 gnotobiotic piglets obtained for this study originated from ten sows. Six of these sows were tested for porcine cytomegalovirus antibodies by the immunofluorescence test, and all were positive at a titre of 1:160. However, the progeny were free from infection as demonstrated by negative lung macrophage cultures and insignificant porcine cytomegalovirus antibody titres once colostral antibody had decayed.
There are no national movement restrictions attaching to PCMV status and no national vaccination or control schemes.
ReferencesTop of page
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