paratuberculosis

Pictures

Picture	Title	Caption	Copyright
Title Symptoms Caption Cattle with end-stage clinical Johne's disease, showing signs of severe emaciation. Copyright Judith R. Stabel/USDA-ARS-National Animal Disease Center	Symptoms	Cattle with end-stage clinical Johne's disease, showing signs of severe emaciation.	Judith R. Stabel/USDA-ARS-National Animal Disease Center
Title Symptoms Caption Cow with clinical Johne's disease, exhibiting severe watery diarrhoea. Copyright Judith R. Stabel/USDA-ARS-National Animal Disease Center	Symptoms	Cow with clinical Johne's disease, exhibiting severe watery diarrhoea.	Judith R. Stabel/USDA-ARS-National Animal Disease Center
Title Symptoms Caption Cow with clinical Johne's disease exhibiting submandibular oedema. Copyright Judith R. Stabel/USDA-ARS-National Animal Disease Center	Symptoms	Cow with clinical Johne's disease exhibiting submandibular oedema.	Judith R. Stabel/USDA-ARS-National Animal Disease Center
Title Pathology Caption Gross pathology of section of ileum from cow with clinical disease, showing corrugation and thickening. Copyright Judith R. Stabel/USDA-ARS-National Animal Disease Center	Pathology	Gross pathology of section of ileum from cow with clinical disease, showing corrugation and thickening.	Judith R. Stabel/USDA-ARS-National Animal Disease Center

Identity

Preferred Scientific Name

• paratuberculosis

International Common Names

• English: Johne's disease; johne's disease, mycobacterium avium subsp paratuberculosis in cattle; johne's disease, mycobacterium avium subsp paratuberculosis in goats; johne's disease, mycobacterium avium subsp paratuberculosis in sheep

English acronym

• PTB

Pathogen/s

Overview

Johne's disease or paratuberculosis (PTB) caused by Mycobacterium avium subspecies paratuberculosis, initially infects the ileum causing a granulomatous enteritis. It then gradually spreads to other parts of the intestine and regional lymph nodes. Johne's disease is a list B disease of the OIE. Incidence of the disease should be reported once yearly.

The disorder known as paratuberculosis or Johne's disease was first described in 1895 by Johne and Frothingham, who identified organisms in the granulomatous lesions in the intestines of affected cattle. These microorganisms stained acid-fast, indicating some type of mycobacterial organism. The bacteria were cultured from cattle in 1910, classified as a mycobacterium and named Mycobacterium chronicae pseudotuberculosae bovis Johne (Twort, 1910; Twort and Ingram, 1912). The organism was later named Mycobacteriumparatuberculosis and the disease was referred to as Johne's disease or paratuberculosis (Bergey et al., 1923). In 1990, the organism was reclassified as M. avium subsp. paratuberculosis due to the close genetic relationship between M. paratuberculosis and M.avium (Thorel et al., 1990).

Paratuberculosis is widely distributed both nationally and internationally in domestic ruminants such as cattle, sheep and goats, as well as in wildlife such as deer, antelope and bison. The prevalence of the disease in the USA is difficult to ascertain because few comprehensive studies have been conducted. In 1996, the National Animal Health Monitoring System conducted a survey of dairy farms in the USA using serological analysis and estimated that 20-40% of these herds were infected with M. avium subsp. paratuberculosis at some level (Wells and Wagner, 2000). European countries such as Belgium, France, Spain, The Netherlands and the UK, together with Canada and some countries in South America, have reported similar prevalence rates for paratuberculosis. Approximately 7% of dairy herds were positive for Johne's disease in a recent survey in Austria (Gasteiner et al., 1999). High prevalence estimates of 55%, 47% and 60% were reported for The Netherlands, Denmark and New Zealand, respectively (Brett, 1998; Muskens et al., 2000; Nielsen et al., 2000). The accuracy of prevalence estimates from these studies is limited by the sensitivity of the diagnostic tests used, accurate recognition and reporting of the disease, and the number of animals sampled. It is estimated that annual losses in the USA from paratuberculosis in cattle herds may exceed US $220 million (Ott et al., 1999). This figure is extrapolated from estimated prevalence values together with computation of financial losses due to culling or death of clinically infected cows, and reduced reproductive efficiency, feed efficiency and decreased milk production in subclinically infected animals. The significance of subclinical infection on economic losses to the producer are detailed in a recent review (Johnson-Ifearulundu and Kaneene, 1997) with a 15-16% reduction in milk production accounting for the major portion of net monetary loss (Abbas et al., 1983; Benedictus et al., 1987). M. paratuberculosis-infected cows beyond the second lactation have demonstrated losses of 1300-2800 lb (590-1270 kg) of milk per lactation (Wilson et al., 1993).

Prevalence rates suggest that paratuberculosis can also be a significant economic factor in small ruminant populations. Studies have reported that 2-5% of sheep flocks in Spain and up to 9% of goat herds in Norway were infected with M. avium subsp. paratuberculosis (Aduriz et al., 1994; Saxegaard and Fodstad, 1985). In addition, Johne's disease has had a major impact on the sheep industry in Australia and New Zealand. Paratuberculosis has also been reported in farmed and free-ranging deer populations in Europe, New Zealand, and North America (Temple et al., 1979; Chiodini and Van Kruiningen, 1983; Power et al., 1993; Fawcett et al., 1995).

This disease is on the list of diseases notifiable to the World Organisation for Animal Health (OIE). The distribution section contains data from OIE's WAHID database on disease occurrence. Please see the AHPC library for further information on this disease from OIE, including the International Animal Health Code and the Manual of Standards for Diagnostic Tests and Vaccines. Also see the website: www.oie.int.

Host Animals

Hosts/Species Affected

Paratuberculosis is widely distributed in many species of domesticated and non-domesticated ruminants throughout the world. It is recognized that herd prevalence may vary widely for many of these species, depending upon a variety of factors such as environment, farm management and control procedures, and host animal genetics. Environmental factors such as soil iron content and soil pH have been implicated in the incidence of paratuberculosis in dairy cattle herds in Michigan (Johnson-Ifearulundu and Kaneene, 1997). Some differences in disease susceptibility have been reported in cattle breeds. A higher incidence of paratuberculosis has been reported for Channel Island breeds in the UK and a higher incidence in Shorthorns in the UK, USA and Canada (Withers, 1959; Julian, 1975). More recently, a higher prevalence for paratuberculosis was noted for Holstein cattle compared to other dairy breeds in Austria (Gasteiner et al., 2000). In addition, a Danish study demonstrated the highest probability for infection in older cows (parity > 4) and the lowest probability in first parity, large-breed cows (Jakobsen et al., 2000). Paratuberculosis is reported most frequently in the Holstein breed in the USA, but this breed comprises a major portion of the dairy cattle population.

Strain differences have been demonstrated between M. avium subsp. paratuberculosis isolates from cattle, sheep and wild ruminants (Pavlik et al., 1995). Restriction fragment length polymorphism analysis has demonstrated distinct differences in genetic patterns between sheep and cattle strains of M. avium subsp. paratuberculosis (Whipple et al., 1990).

Systems Affected

Distribution

For current information on disease incidence, see OIE's WAHID Interface.

Distribution Table

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

Pathology

Gross pathology of affected animals is primarily associated with the lower small intestine and regional lymph nodes. Paratuberculosis is characterized by a granulomatous inflammation involving, mainly the ileo-caecal valve, ileum, and associated lymph nodes (Chiodini et al., 1984). This inflammation makes the intestinal wall appear thickened and corrugated, associated lymph nodes become enlarged and oedematous. However, in advanced stages of clinical disease, infection may become systemic and affect many tissues including the mammary gland, kidneys, liver, and male and female reproductive organs (Doyle, 1954; Kopecky et al., 1967; Larsen and Kopecky, 1970; Larsen et al., 1981; Hines et al., 1987). Atherosclerosis and calcification of the aorta and heart have been observed in cattle with end-stage clinical disease (Buergelt et al., 1978).

Tissue changes are accompanied by increased leakage of plasma proteins across the intestinal wall and malabsorption of amino acids and other nutrients from the intestine, with little or no evidence of mucosal necrosis. A decrease of calcium total protein and albumin has been observed in serum from cattle and sheep with clinical paratuberculosis (Jones and Kay, 1996). Accumulations of lymphocytes, macrophages, epithelioid cells, and other inflammatory cells are found in the lamina propria. Microscopic lesions consist primarily of macrophage infiltrate; they are multifocal in the early stages of disease but become more diffuse as the disease progresses to more advanced stages. Giant cells are a characteristic of Johne's disease and occur more frequently in advanced clinical disease.

In some clinical cases, inflammatory cells may be observed in the submucosa as a band of epithelioid cells and lymphocytes along the muscularis mucosa. In general, granulomas or diffuse cellular infiltrates containing numerous acid-fast bacilli are present in clinical cases. Villi become blunt and may fuse together. It should be noted that cases with severe lesions and few bacilli can be observed, mainly in sheep (Marin et al., 1992; Pérez et al., 1996; Clarke, 1997). Caseous necrosis has been observed in paratuberculous lesions most frequently in goats, and certain captive wild species such as deer (Stehman, 1996). In sheep and goats lymphangiectasia, is most often observed in clinical cases (Marin et al., 1992; Perez et al., 1996; Clarke, 1997). Moreover, in sheep and goats gross changes associated with a thickening of the ileum and jejunum are sometimes difficult to detect, and do not resemble those observed in cattle (Marin et al., 1992; Clarke, 1997).

Diagnosis

Host immunity

Diagnosis of paratuberculosis is difficult because of the fastidious growth pattern of the microorganism and because of the paradoxical pattern of the host animal immune response to infection. Efforts to control Johne's disease in cattle and other species has been limited by the lack of rapid, reliable diagnostic tests for identifying all M. avium subsp. paratuberculosis-infected animals. Moreover, the sensitivity of different diagnostic methods is influenced by the prevalence of the infection in the herd and by the stage of the disease, depending on the individual host response (Clarke, 1997; Marin et al., 1992; Sweeney et al., 1995; Pérez et al., 1997; Pérez et al., 1999).

During the early subclinical stages of infection, the microorganism elicits a cell-mediated response that can be characterized by strong delayed type IV hypersensitivity reactions, lymphocyte proliferative responses to mitogens and production of cytokines by stimulated T lymphocytes (Kormendy et al., 1990; Stabel, 1996; Perez et al., 1999). When the disease progresses from subclinical to clinical stages, the cell-mediated immune response wanes and a strong humoral response predominates. The presence of antibody to M. paratuberculosis does not protect the host against the disease; indeed, active cell-mediated immunity appears to be essential to keep the infection in check. During the final stages of disease, lack of antigen-specific cell-mediated immune responses or complete anergy may result, allowing for rapid dissemination of the infection throughout the host (Bendixen, 1978).

Clinical diagnosis

The most characteristic sign of paratuberculosis is the occurrence of profuse, watery diarrhoea in infected animals. The diarrhoea does not respond to treatment and may be chronic or appear intermittently over more protracted periods. In the early stages of clinical disease, the animal may continue to eat well but the diarrhoea causes a rapid weight loss. Because of the malabsorption of nutrients, animals begin to show signs of unthriftiness such as rough hair coat and dry skin. At the end stage of clinical disease, severe emaciation and submandibular oedema or ‘bottle-jaw’ may develop due to the leaking of serum proteins across capillary walls. In sheep and goats, diarrhoea is not a characteristic sign but some softening of the stool is observed; however, weight loss and unthriftiness do occur. Fever has been reported in some cases but is not a consistent sign of infection with M. avium subsp. paratuberculosis.

Laboratory diagnosis

Bacterial culture is the most definitive method of diagnosis because it can detect animals that are subclinically infected and also animals in advanced stages of disease. Use of faecal samples in mycobacteriologic culture is routine to confirm diagnosis of M.avium subsp. paratuberdulosis. Subclinically infected animals shed low numbers of M. avium subsp. paratuberculosis in their faeces and usually shed intermittently over time. During the clinical phase of infection, faecal shedding of the microorganism is quite high and may exceed 10⁸ cfu per gram of faeces (Chiodini et al., 1984). Primary culture of the bacilli is very time-consuming, requiring up to 12 weeks of incubation, and is also labour intensive. Contamination by other fungal and bacterial microorganisms is often a problem when M. avium subsp. paratuberculosis is being cultured from faecal specimens, so incorporation of a decontamination procedure before culture is standard protocol for diagnostic laboratories. Because infected animals may shed the organism intermittently in their faeces, use of faecal culture alone as a diagnostic tool may result in misrepresentation of infection within the herd; only about 50% of M. avium subsp. paratuberculosis infection is detected by faecal culture (Sanftleben, 1990). For surveillance purposes detection of M. avium subsp. paratuberculosis in environmental or pooled faecal samples is an efficient procedure (van Schaik et al., 2007; Aly et al., 2009).

Though several serologic tests have been developed for detecting antibodies in sera of cattle experimentally or naturally exposed to M. avium subsp. paratuberculosis, many of these tests have not been widely used under field conditions. Serological tests for diagnosis of paratuberculosis, such as agar gel immunodiffusion (AGID), enzyme-linked immunosorbent assay (ELISA) and complement fixation (CF), are relatively easy to perform but sensitivity of detection is only moderate (Colgrove et al., 1989). AGID is most often used as a rapid diagnostic method for confirmation of clinical paratuberculosis.

Most widely used is the ELISA, sometimes in conjunction with other diagnostic methods such as faecal culture or direct faeces PCR. Reported sensitivity values for ELISA are 15-57% for subclinically infected cattle shedding low numbers of organisms in their faeces; the average is 88% for clinically infected cattle shedding high numbers of organisms (Collins and Sockett, 1993; Sweeney et al., 1995). More recently, a multi-university study also demonstrated that several ELISA tests (three on serum and one on milk) had sensitivities ranging from 27.8-28.9% (Collins et al., 2005). Milk ELISA testing is an attractive low cost method for diagnosing paratuberculosis in large scale monitoring systems (van Weering et al., 2007). The CF test is most frequently used to test cattle for import and export purposes, yet has a lower specificity than the AGID or ELISA tests. A study of sera from cattle in herds with a previous diagnosis of paratuberculosis provided additional evidence about the sensitivity of ELISA in cattle with clinical disease (Thoen and Moore, 1989). However, a high percentage of subclinically affected cattle, sheep and goats, in which M. avium subsp. paratuberculosis was isolated from tissue specimens at necropsy failed to have detectable mycobacterial antibodies in sera. Therefore, although serologic tests are useful in detecting cattle with clinical paratuberculosis, the application of ELISA for identifying cattle in the early stages of infection or shedding small numbers of organisms in the faeces is of limited value (Clarke, 1997; Sweeney et al., 1995; Perez et al., 1997). The AGID is more efficient in small ruminants than in cattle; the sensitivity is related with the stage of the infection, (high in clinical or preclinical cases and very low in the early stages of the infection) (Marin et al., 1992; Shulaw et al., 1993; Perez et al., 1997).

Intradermal injection of PPD prepared from the culture filtrate of M. avium subsp. paratuberculosis has been used to detect cattle with paratuberculosis. Although some cattle that respond to such an injection have been suggested to be only sensitized and not infected, recent findings indicate the organism is isolated in a high percentage of skin test positive animals. This test requires a veterinarian to inject the animal with johnin and read the change in skin thickness 24-48 h after injection. Although the test is inexpensive and rapid, it does require two visits by the veterinarian and has a low sensitivity of detection compared to other techniques. The specificity of the test may be improved markedly by the use of purified antigens.

In vitro lymphocyte blastogenic assays, which are considered an in vitro correlate of delayed-type hypersensitivity, have been developed for use in the diagnosis of paratuberculosis and seem to be valuable in detecting some cattle with subclinical disease. The measure of interferon-gamma (IFN-g ) has been evaluated as diagnostic tool for paratuberculosis (Billman-Jacobe et al., 1992). IFN-g is a protein that is released by activated T cells after stimulation with mycobacterial antigens (Wood et al., 1990). This test is proving to be efficacious in identifying animals in the early subclinical stages of disease. The IFN-γ assay should be used to test the level of exposure to paratuberculosis bacteria the animals have experienced, and thereby assist in maintaining rational in-herd management procedures and in the establishment of paratuberculosis status of a given herd. Combining the ELISA and IFN-g tests to screen herds infected with M. avium subsp. paratuberculosis resulted in the identification of 70-90% of infected animals based upon faecal culture, indicating that using a combination of cellular and antibody assays may improve the detection of M.avium subsp. paratuberculosis-infected cattle.

Specific gene probes and polymerase chain reaction (PCR) have been used for rapid detection of M. avium subsp. paratuberculosis in clinical specimens (Giessen et al., 1992; Collins et al., 1993; Wentnik et al., 1994; Theon and Haagsma, 1996). These techniques provide for reporting of results to the clinician in 4 to 5 days, and are valuable when purchasing replacement cattle, in which obtaining results in a short time is necessary. Because false-positive reactions have been observed in reference laboratories conducting PCR analysis, it is necessary to include suitable controls to validate results (Cousins et al., 1999). False-negative reactions also may be observed; the sensitivity of PCR test is not as high as faecal culturing (Sockett et al, 1992). The use of PCR on blood samples for the detection of M. avium subsp. paratuberculosis in macrophages is possible however only in a limited number of cases of advanced paratuberculosis (Giessen et al., 1992; Zhong et al., 2009). The most commonly used gene sequence for PCR detection of M. avium subsp. paratuberculosis is the insertion element, IS900 (Vary et al., 1990). Nested PCR has increased the sensitivity of this technique several fold over regular PCR, detecting 50 organisms per gram of faeces versus 104 organisms per gram of faeces (Collins et al., 1993). A limitation to the use of PCR is the expense of the method, at two to three times the price of faecal culturing. In the US, the cost of PCR is comparable to culture (US $15-25). Recent advances have focussed on more efficient techniques to isolate nucleic acids from faecal samples while limiting extraction of PCR inhibitors, thus increasing sensitivity of these techniques for practical purposes (Okwumabua et al., 2010).

Restriction fragment length polymorphism analysis, PGFE, MIRU-VNTR and SSR typing have provided for the differentiation of M. avium subsp. paratuberculosis isolates from animals, and are valuable in epidemiological investigations (Collins et al., 1990; Pavlik et al., 1995; Sevilla et al., 2008; Pradhan et al., 2011; van Hulzen et al., 2011). However, the cost of these molecular techniques limits their use on a herd basis.

For practical purposes the present day challenge for practitioners is to select the appropriate test for the intended purpose. The intended purposes range from testing in control programs in M. avium subsp. paratuberculosis infected high prevalence herds, with the absorbed ELISA as the preferred test,to surveillance and confirming clinical diagnoses. If the primary aim is surveillance then environmental or pooled faecal samples should be used (van Schaik et al., 2007; Aly et al., 2009). For eradication strategies the pooled faecal culture or pooled faecal PCR are the test of choice. In case of confirmation of a clinical diagnoses in unsuspect herds necropsy, faecal culture or faecal PCR should be chosen. In case of confirmation of a clinical diagnoses in herds with endemic paratuberculosis, ELISA, faecal culture or faecal PCR can be used. (Collins, 2011).

List of Symptoms/Signs

Disease Course

The clinical signs that occur, following a prolonged incubation period of one to ten years, include gradual weight loss despite a normal appetite. Other than the loose consistency the faeces appear normal. As the disease progresses, the affected animals become increasingly lethargic and emaciated. Cachexia and ‘waterhose’ diarrhoea characterize the terminal stages of the disease in cattle, but diarrhoea is not a common feature in sheep and goats (Chiodini et al., 1984; Marin et al., 1992). The main clinical signs of the disease are progressive weight loss and a decrease in milk production; therefore, most farmers cull the animals before severe clinical signs are present and before the diagnosis of PTB is made. Consequently, the number of clinical cases of PTB may be underestimated (Chiodini et al., 1984; Marin et al., 1992).

Transmission of M. avium subsp. paratuberculosis in cattle herds varies considerably, depending on the management practices followed. In some herds, the disease may persist for years before clinical signs appear. The most common route of exposure of calves to M. avium subsp. paratuberculosis is by ingestion of contaminated faeces on the surface of the dam’s mammary gland during suckling (Chiodini and Davis, 1993; Thoen and Haagsma, 1996). Young calves, seem to be more susceptible to the infection than older or adult cows (Hagan, 1938; Payne and Rankin, 1961; Larsen et al., 1975). Faeces contaminate pastures so feed and water are a source of infection. M. avium subsp. paratuberculosis can resist destruction for several months in the natural environment, making prevention and control difficult (Jorgensen, 1977). Moreover, colostrum, placenta and uterus as well as the foetus may be infected adding to the difficulty in the control of paratuberculosis (McQueen and Russell, 1979; Seitz et al., 1989; Rohde and Shulaw, 1990; Sweeney et al., 1992a,b; Streeter et al., 1995). The primary route of infection is through ingestion of faecal material, milk or colostrum containing M. avium subsp. paratuberculosis microorganisms (Chiodini et al., 1984) which, once ingested, survive and replicate within macrophages of the small intestine and in regional lymph nodes. The organisms have been isolated from semen and reproductive organs of bulls, but the importance of these findings in transmission is unclear. The incubation period, which is the interval between infection and the observation of clinical disease (diarrhoea and weight loss), usually varies from 1 to 5 years or more. Cattle become infected with M. avium subsp. paratuberculosis as calves but often do not develop clinical signs until 2-5 years old.

It is most important to emphasize that clinical disease may be observed in less than 30% of the M. avium subsp. paratuberculosis-infected cattle in a herd. Some animals remain subclinically infected throughout their lifetime and never show clincial signs of disease. Clinical disease is most often observed in cattle 3 to 6 years old. While in sheep and goats, most of the cases are present in animals 2 to 4 years of age or mainly during the first and second lactation (Clarke, 1997). Stress is an important factor that may contribute to the onset of clinical disease; the stress may be related to parturition and/or increased milk production. In beef bulls, stress may be associated with the breeding season.

When clinical signs are evident, the animal usually sheds M. avium subsp. paratuberculosis in the faeces; however, some animals shed intermittently and micro-organisms may not be found on the examination of a single faecal sample. Mycobacterium avium subsp. paratuberculosis has been isolated from the milk of cows with clinical paratuberculosis and from milk samples and supra-mammary lymph node specimens in cows with subclinical infection (Taylor et al., 1981; Sweeney et al., 1992a; Streeter et al., 1995).

After an incubation period of several years, extensive granulomatous inflammation occurs in the terminal small intestine, leading to malabsorption of nutrients and protein-losing enteropathy. During this period the animal may suffer from chronic watery diarrhoea, rapid weight loss, diffuse oedema and rough hair coat. The chronic diarrhoea fails to respond to antibiotic treatment and animals continue to lose weight, despite adequate food intake. Clinical disease may persist for 6 months or more; diarrhoea may be intermittent and cattle sometimes seem to recover for a few weeks. In terminal stages, diarrhoea results in emaciation and death. However, in small ruminants diarrhoea can be absent in a high percentage of clinical cases (Stamp and Watt, 1954; Clarke, 1997).

A decrease in total serum protein, albumin, triglycerides and cholesterol concomitant with increased creatine kinase and aldolase may occur in the latter stages of disease due to muscle damage (Patterson et al., 1965, 1967, 1968). Some animals will progress rapidly to an end-stage of recumbency and death, whereas others may go into remission for a period of time, although generally these animals will succumb to clinical signs at a later date. Clinical signs of paratuberculosis in sheep, goats and deer are limited to weight loss, unthriftiness and slight softening of their stools. Poor condition of wool is associated with end-stage disease in sheep (Cranwell, 1993). Intestinal thickening is variable in these species with some enlargement of lymph nodes.

Pathogenesis

M. avium subsp. paratuberculosis penetrates the intestinal epithelial layers through the follicle-associated epithelium or M cells (Momotani et al., 1988). Phagocytes engulf the organisms and are usually unable to degrade them and they remain viable and protected from humoral factors. The first small and limited granulomatous lesion is detected in the ileo-caecal and jejunal Peyers patches, and can persist as latent infection for long periods of time, as has been observed in experimental infections in sheep and goats as well as in natural infections (Pérez et al., 1996). If the infection progresses, lesions spread to mucosa affecting different parts of the small intestine and associated lymph nodes (Juste et al., 1994; Pérez et al., 1996).

During early infection, a cell-mediated immunity (CMI) is predominant, but, as the lesions and disease progresses a humoral response appears due to the presence of large numbers of bacilli released by dying macrophages. The first response is a cell-mediated type Th1 with cytokines such as IL-2 and gamma IFN, as reported in other mycobacterial infections. The progression of the disease is associated with the T-gamma delta cell which inhibits CD4 and helper lymphocytes (Chiodini and Davis, 1993). Th2-like cytokines (IL-4 and IL-10) are predominant in clinical cases with multibacillary and non-lymphocitic lesions (Wards et al., 1995; Little et al., 1996; Burrells et al., 1998; Navarro et al., 1998).

In natural cases a similar situation has been reported with cellular immune response associated with latent-focal subclinical lesions and humoral immunity with high antibody response in diffuse-severe and multibacillary cases (Bendixen, 1978; Clarke, 1997; Pérez et al., 1997; Pérez et al., 1999). However, in some clinical cases with severe disease animals can show a strong CMI response (skin hypersensitivity and low antibody response) with predominant presence of lymphocytes in the lesion and few bacilli (Clarke, 1997; Pérez et al., 1997; Pérez et al., 1999). This is more common in sheep. Therefore, in M. avium subsp. paratuberculosis. infections it has been proposed that an immunopathological spectrum develops, which depends on the host response (Pérez et al., 1999). A clear relationship with pathology, number of bacilli, and the cellular or humoral immune mediated response has been established in sheep (Clarke, 1997; Pérez et al., 1997; Pérez et al., 1999).

Epidemiology

Paratuberculosis is observed in cattle and small ruminants worldwide (Chiodini et al., 1984). Prevalence of the disease, on the basis of mycobacteriologic examination of tissue specimens collected from adult cattle at slaughter or on histopathological and serologic studies (i.e., ELISA), varies considerably. In the United States, the overall prevalence was reported to be 1.6% (Merkal et al., 1987); however, in the north-eastern section of the country, the reported prevalence ranged from 1.9 to 18.7%. A study conducted in 1996 by the National Animal Health Monitoring Service estimated that 21.6% of the dairy herds in the US are infected with M. avium subsp. paratuberculosis. These data may, however, markedly underestimate the true incidence of infection in animal populations. In Europe, several surveys made in slaughterhouses suggested a prevalence from 9.8% in Denmark to 11% and 17% in England (Jorgensen, 1972). In sheep in different countries and using various methods (serology, histopathology and culture) the prevalence was found to be higher than 10% in individual animals and 40% in flocks (Juste et al., 1991). In some populations, in particular those of goats, the prevalence has been reported to be 90% or more.

Cattle usually become infected with M. avium subsp. paratuberculosis as calves, but often do not develop clinical signs until 2-5 years of age (Larsen et al., 1975). Calves may become infected in utero or as neonates through ingestion of faecal material, milk or colostrum containing M. avium subsp. paratuberculosis microorganisms. Once ingested, it is believed that the M cells of the Peyer's patches serve as the portals of entry for M. avium subsp. paratuberculosis into the lymphatic system, as they do for other enteric intracellular pathogens such as Salmonella (Momotani et al., 1988). Live M. avium subsp. paratuberculosis will then traverse the M cell by transcytosis and be expelled on the basolateral side of the cell to be scavenged by the macrophages or dendritic cells. M. avium subsp. paratuberculosis may survive and replicate within the macrophages of the intestine and regional lymph nodes of the host animal, leading to a protracted subclinical phase of infection. Cattle shed minimal amounts of M. avium subsp. paratuberculosis in their faeces in the subclinical phase of infection, yet, over time, this shedding can lead to significant contamination of the environment and an insidious spread of infection throughout the herd. After an incubation period of several years, the immune system of the host animal may become compromised and the animal is no longer able to contain the infection. At this point, extensive granulomatous inflammation occurs in the terminal small intestine due to an influx of macrophages. Thickening of the intestinal epithelium occurs, resulting in malabsorption of nutrients and protein-losing enteropathy. The terminal stage of disease is characterized by chronic diarrhoea, rapid weight loss, diffuse oedema and inappetence.

Transplacental infection of foetuses with M. avium subsp. paratuberculosis occurs most frequently in foetuses from infected cows in the clinical stage of disease. A study examining infection of foetuses from cows with clinical signs of paratuberculosis found that 26.4% of foetuses had tissues that were culture-positive (Seitz et al., 1989). A lower level of foetal infection was noted in a more recent study evaluating foetal infection in cows that were faecal-culture positive for M. avium subsp. paratuberculosis but asymptomatic for clinical signs (Sweeney et al., 1992a). Foetal tissues were positive for M. avium subsp. paratuberculosis in only 8.6% of infected cows and these cows were shedding high numbers of the bacterium in their faeces. Results from these studies suggest that foetal infection is highly correlated with the level of faecal shedding of M. avium subsp. paratuberculosis and most commonly associated with cows demonstrating clinical signs of disease.

Faecal contamination of the environment poses the most likely threat for calfhood infection. In the clinical stage of disease, cows may shed up to 10⁸ organisms per gram of faeces (Chiodini et al., 1984). Contamination of the maternity pen is a primary source for calves to gain contact with the bacterium. However, contamination of water sources, feed troughs and pasture may also contribute to transmission of infection. Early studies established that M. avium subsp. paratuberculosis is capable of surviving in water, soil and manure for periods up to 1 year. A recent study, however, indicates that the organism does not survive in high numbers during composting and liquid manure storage methods (Grewal et al., 2006). M. avium subsp. paratuberculosis is also shed in the colostrum and milk of infected dams, albeit in low numbers. Although shedding of the bacterium in colostrum has not been adequately quantified it is highly correlated with heavy shedding in the faeces of infected dams (Streeter et al., 1995). Shedding of M. avium subsp. paratuberculosis in the milk of infected dams is also associated with the degree of faecal shedding by the dam, and M. avium subsp. paratuberculosis has been found in viable numbers at low concentrations (5-8 cfu/50 ml of milk) (Sweeney et al., 1992b).

Viable M. avium subsp. paratuberculosis organisms have also been isolated from the reproductive organs of infected females and the reproductive organs and semen of infected bulls, which may contribute to disease transmission (Philpott, 1993). Although it is unclear whether embryo transfer contributes to the incidence of disease, viable M. avium subsp. paratuberculosis organisms have been cultured from uterine washings of clinically infected dams (Rohde and Shulaw, 1990). Experimental evidence also indicates that M. avium subsp. paratuberculosis may attach to bovine ova, suggesting a potential source of uterine infection of the recipient cow (Rohde et al., 1990).

Impact: Economic

Economic losses caused by paratuberculosis are attributable to decreased value of breeding stock, lower milk production, reduced salvage or slaughter value of animals with advanced clinical disease, increased cull rates of high-producing animals, decreased feed conversion rates, and reduced fertility (Benedictus et al., 1984; Merkal et al., 1987; Thoen and Moore, 1989). The significance of subclinical paratuberculosis on economic losses to the producer are detailed in a recent review; a reduction in milk production of 15-16% accounts for the major portion of net monetary loss (Johnson-Ifearulundu and Kaneene, 1997). Cows that are infected with M. avium subsp. paratuberculosis beyond second lactation have demonstrated losses of 1300-2800 lb (590-1270 kg) of milk per lactation (Wilson et al., 1993).

Although the estimated losses from paratuberculosis vary, in commercial dairy herds in which M. avium subsp. paratuberculosis infection persists, losses are considerable (Benedictus et al., 1984). Decreases in milk production have been estimated between 4% to 18% in subclinical infected animals and about 20% or higher in clinical cows (Buergelt and Duncan, 1978; Benedictus et al., 1984; Goodger et al., 1996; Spangler et al., 1992). A recent survey of dairy herds in the USA suggested an annual loss of more than US $200 per cow for positive herds with at least 10% of their cull cows showing clinical signs of Johne's disease (Ott et al., 1999). Most of this cost could be attributed to a significant reduction in milk production per cow (1500 lb; 680 kg). The national average cost across all herds in the USA was US $22 per cow with an economic loss of US $220 million per year to the dairy industry. Other studies have reported costs per cow ranging from US $145 to US $1094 per cow with Johne's disease. Reduced production of meat, wool and milk has been reported for sheep with paratuberculosis, together with increased mortality rates (Aduriz et al., 1994; Chaitaweesub et al., 1999). An average loss of $29 per sheep was reported for flocks with Johne's disease in Australia, but some highly infected flocks demonstrated losses of $64 per sheep.

Zoonoses and Food Safety

It is not clear whether M. avium subsp. paratuberculosis is a human pathogen and what potential danger it may present to consumers exposed to dairy or meat products from infected animals. Several species of mycobacteria, including M. fortuitum, M. avium-intracellulare, M. cheloni and M. kansasaii, have been found in intestinal biopsy tissue from Crohn's disease patients (Chiodini, 1989). M. avium subsp. paratuberculosis has also been identified by primary isolation from intestinal tissue and by PCR analysis of DNA specific for this organism. Because the clinical symptoms of Crohn's disease closely mimic those found in animals with paratuberculosis in the late stages of disease, it has been proposed by a number of investigators that M. avium subsp. paratuberculosis may be a causative agent of this disorder in humans (Sanderson et al., 1992; Mishina et al., 1996).

The potential relationship between Crohn's disease and M. avium subsp. paratuberculosis has become an issue for the dairy industry since the publication of a report in 1994 by a group in the UK that suggested that viable M. avium subsp. paratuberculosis organisms were present in pasteurized milk purchased from retail markets (Millar et al., 1996). Studies evaluating the optimal time and temperature for heat inactivation of M. avium subsp. paratuberculosis have followed (Grant et al., 1996; Hope et al., 1996; Stabel et al., 1997; Sung and Collins, 1998; Keswani and Frank, 1998). In 2005, a USA study reported viable M. avium subsp. paratuberculosis was found in pasteurized milk purchased from retail stores (Ellingson et al, 2005).

Further concerns regarding exposure of humans to M. avium subsp. paratuberculosis have been raised by observations of viable M. avium subsp. paratuberculosis in cheese (Spahr and Schafroth, 2001; Williams and Withers, 2010); meat (Mutharia et al., 2010; Whittington et al., 2010), dust (Eisenberg et al., 2010) and water (reviewed in Gill et al., 2011).

Despite the enduring controversy on the causal etiological role of M. avium subsp. paratuberculosis in the development of Crohn’s disease in humans (BANR, 2003b) the concerns have been instrumental in establishment of national control programs in multiple countries including, Australia, Denmark, the Netherlands and the United States

Disease Treatment

Antimicrobial therapy for treatment of animals infected with M. avium subsp. paratuberculosis is not recommended unless the animal has a high genetic value. There is no cure for paratuberculosis, so drug therapy is used only to prolong the life of an infected animal and reduce clinical signs of disease. Standard antituberculosis drugs such as clofazimine, isoniazid, rifabutin, rifampin, and streptomycin have been tested both in vitro and in vivo for effects on M. avium subsp. paratuberculosis (St. Jean, 1996). Although many in vivo studies have demonstrated an improvement in clinical signs of paratuberculosis after treatment, the organism may be still be shed in the faeces. Combination therapy with two or more of the antimycobacterial agents has proven to be more effective than one compound (Rankin, 1955; Slocombe, 1982; Das et al., 1992). Treatment times are protracted and treatment may be necessary for the remainder of the animal's life. This is a significant disadvantage because the cost of therapy can range from US $1.00 to US $219.00 per animal per day, depending upon the compound used (St. Jean, 1996).

Prevention and Control

Farm level control

The economic impact of paratuberculosis in cattle herds can be greatly minimized when suitable control programmes are followed (Benedictus et al., 1984; Thoen and Moore, 1989). Control programs should control three major critical points in disease transmission. They should prevent exposure of susceptible animals, in particular young stock, to the infectious agent. They should enable the identification and stimulate the eradication of infected animals from the herd. Finally they should prevent entry of infected animals into the herd.

These programmes involve application of recognized tests and new molecular techniques to identify and remove cattle that have subclinical disease and shed M. avium subsp. paratuberculosis. Moreover, initiating changes in management is often necessary to prevent exposure of replacement heifers to the organism. Cattle to be introduced into the herd should be free of Johne's disease and herd replacements should originate from herds without a history of Johne's disease. When the status of the source herd is unknown, the purchased replacements should be quarantined for 3-4 months and retested for evidence of M. avium subsp. paratuberculosis infection. Thorough cleaning and disinfecting of pens is an essential part of a control program (Rossiter, 1996; Eisenberg et al., 2011a; Garry, 2011).

Within the herd, neonates are the most susceptible group of animals to infection with M. avium subsp. paratuberculosis; therefore, one recommendation is to remove the calf from the dam immediately after birth (Whitlock et al., 1994). Because M. avium subsp. paratuberculosis is shed into the colostrum and milk of clinically infected cows, the stockperson should ensure that calves are fed uncontaminated colostrum (from test-negative animals) and milk replacer to prevent infection. The maternity pen is also an excellent site for neonatal exposure to contaminated faecal matter in bedding, feed and on the dam's udder. In addition M. avium subsp. paratuberculosis can be transmitted from infected to susceptible individuals via bioaerosols such as dust (Eisenberg et al., 2010; Eisenberg et al., 2011b). Separate housing of calves in separate buildings is called for to prevent bioaerosol transmission.

Segregation of infected animals from uninfected animals is a good idea at any age, and the manure of each group should be disposed of separately. A common flaw of many dairy operations is the use of the same skid loader for feeding and manure disposal (Goodger et al., 1996). Cross-contamination of feed is a major contributor to the spread of paratuberculosis, and faeces are the major source of the causative organism. Use of improperly treated manure solids to fertilize pastures on the farm is another source of M. avium subsp. paratuberculosis infection because the organism can survive in the soil for up to 1 year. Furthermore, stagnant water sources are excellent reservoirs for numerous bacteria, and M. avium subsp. paratuberculosis has been found to survive in pond water for 270 days (Lovell et al., 1944). Therefore, it is advisable to prevent access to stagnant water. Preferably, a clean water source should be provided for each age group of animals.

Control of Johne's disease should include procedures to identify and remove adult cattle shedding M. avium subsp. paratuberculosis in faeces (Moyle, 1975).Microbial culture of faecal samples or faecal PCR should be used. Although standard faecal cultures can have prolonged incubation periods, these can be timed so that results are available for making management decisions. There are also liquid culture techniques available in some laboratories that provide test results in about half the time of a standard faecal culture. Cattle that shed organisms intermittently or in low numbers may not be identified on bacteriologic examination. The problem is further complicated because cattle in early stages of infection usually do not shed bacteria in their faeces, which is particularly important when a culture of faecal samples is used in attempts to eliminate paratuberculosis from a herd or to select replacement cattle free of disease. Therefore, recommendations are to test animals annually and make purchases based on herd level testing, not individual animal test results. This is especially true when purchasing young stock, in which infection is less likely to be identified by current testing methods. Due to strain differences, care must be taken in culturing samples from small ruminant species. Culture techniques using environmental (Raizman et al 2004) and pooled faecal samples (Wells et al., 2002; Whittington et al., 2000) provide more economical means of detecting the disease and estimating prevalence in surveillance strategies.

Vaccines

Vaccination of calves, using a whole bacterin vaccine at less than 35 days of age is practised in several countries (Thoen and Moore, 1989; Körmendy, 1994), however banned in others due to interference with tuberculosis control programs. In one study, the prevalence of paratuberculosis, as assessed by faecal microscopic tests, decreased from 48 to 1.4% in vaccinated cattle, whereas in non-vaccinated cattle, prevalence was maintained at about 30% (Körmendy, 1994). Others studies in vaccinated young cattle reported a decrease in number of shedder animals and the level of excretion (Jorgensen, 1984; Whipple et al., 1992). There is consensus that in vaccinated herds clinical cases markedly decrease or disappear and is therefore economically attractive despite the fact that this vaccination procedure does not prevent transmission of infection (Benedictus et al., 1988; van Schaik et al., 1996; Kalis et al., 2001).

Vaccine is used in several countries for the control of paratuberculosis in sheep and goats. Vaccination performed in these species, either in young or adult animals, has been useful to eliminate clinical cases and to reduce infection (Sigurdsson, 1960; Crowther et al., 1976; Pérez et al., 1995; Saxegaard and Fodstad, 1985; Corpa et al., 2000). In experimental studies, it has been shown that vaccination modifies the pathology of infection to regressive forms (Juste et al., 1994). A vaccine trial in sheep (Reddacliff et al, 2006), demonstrated that sheep vaccinated against Ovine Johne’s disease (OJD) had 90% reductions in mortality, number of animals shedding and amount of organisms shed.

In dairy cattle a significant economical benefit was reported in the herd using vaccine (Benedictus et al., 1984; Spangler et al., 1991). Vaccination should be recommended as a means of reducing disease in herds in which the infection rate is greater than 5% based on faecal culture and/or management changes cannot be implemented. The products used for vaccination vary from killed to attenuated live organisms, with or without adjuvant. In most of the countries, the use of vaccine requires official approval from regulatory officials. Regulatory officials often require tuberculin testing of adult cattle prior to approving use of vaccine, to exclude the presence of M. bovis infection. Vaccination is not recommended in herds in which M. bovis persists. In herds in which animals have been vaccinated serologic tests are of no value in identifying M. avium subsp. paratuberculosis-infected animals (Momotani et al., 1988; Spangler et al., 1992). Cellular assays such as lymphocyte blastogenesis, gamma IFN test or skin tests also should not be used, because false-positive reactions may be observed for several months or more (Inglies and Weipers, 1963). Due to the influence vaccine has on the immune system, Johne’s diagnostics in vaccinated herds should use organism detection based tests such as culture or PCR.

Other recommendations for control

Other recommendations for control include (BANR, 2003a):

1. Prevent introduction of disease through purchased animals


2. Isolate and slaughtering clinically affected animals


3. Culling most recent offspring of clinical cases as soon as possible


4. Removing calves from dams immediately upon birth (before suckling)


5. Isolating calves in separate calf-rearing area


6. Harvesting colostrum from cows with cleaned and sanitized udders


7. Feeding colostrum from test-negative cows to calves then only feed milk replacer or pasteurized milk


8. Preventing the contamination of calf feedstuffs, water, or bedding by adult cow manure


9. Do not apply manure to land that will be harvested for forage within the same season

Diagnostic testing can be a useful tool in controlling the spread of Johne’s disease within and between farms. A number of factors including species, estimated herd/flock prevalence, farm goals, and cost against benefit of testing, all play into the decision of whether to test and which test to use on an individual operation. Conducting a herd risk assessment and making a herd management plan are valuable practices in helping to make these decisions.

Calving pens and facilities should be properly cleaned and disinfected periodically with a cresylic or substituted cresylic disinfectant that kills M. avium subsp. paratuberculosis. This is probably the most important management change that can be readily made on most premises with little expense to the owner, but is often ignored by many producers.

In most commercial dairy herds, control of Johne's disease can often be accomplished by changes in management. New molecular techniques provide for more rapid identification of cattle shedding M. avium subsp. paratuberculosis and subsequent removal of such cattle from the herd, but the potential for false-positive and false-negative results must be considered by the veterinarian and client. Eradication of paratuberculosis will depend on development of sensitive immunological procedures (e.g.: skin tests or gamma-IFN with species-specific antigens) for detecting all M. avium subsp. paratuberculosis-infected animals.

References

Abbas, B, Riemann HP, Hird DW, 1983. Diagnosis of Johne's disease (paratuberculosis) in northern California cattle and a note on its economic significance. Calif. Vet., 37(10):16

Aduriz A, Juste RA, Saez de Ocariz C, 1994. An epidemiologic study of sheep paratuberculosis in the Basque Country of Spain: serology and productive data. In: Proceedings of the Fourth Intl. Colloq. Paratuberculosis, Cambridge, UK, 1994. Intl. Assoc. Paratuberculosis, 19

Albiston HE, Talbot RJ, 1936. Johne's disease in Victoria. Aust. Vet. J., 12:125-138

Alibasoglu M, Erturk E, Yucel N, 1973. Pathological investigations on the first cases of caprine paratuberculosis (Johne's disease) diagnosed in Turkey. Vet. Fakult. Derg., 20(1):43-63

Allworth MB, Kennedy DJ, 2000. Progress in national control and assurance programs for ovine Johne's disease in Australia. Veterinary Microbiology, 77(3/4):415-422; 9 ref

Aly SS, Anderson RJ, Whitlock RH, Fyock TL, McAdams S, Adaska JM, Jiang J, Gardner IA, 2009. Reliability of environmental sampling to quantify Mycobacterium avium subspecies paratuberculosis on California free-stall dairies. Journal of Dairy Science, 92(8):3634-3642. http://jds.fass.org/cgi/content/abstract/92/8/3634

Andrews WH, 1926. Animal diseases in South Africa. Vet. Rec., 6:447-455

Badnjevic B, Forsek Z, 1968. [Bovine and ovine paratuberculosis in certain regions of Yugoslavia]. Bull. Off. Int. Epizoot., 70(1):701-706

Badnjevic B, Forsek Z, Kovacevic E, 1962. Johne-ova bolest (paratuberkulosa) kod goveda. Prva laboratorijska dijagnoza u Bosni I Hercegovini [Johne's disease (paratuberculosis) in cattle. First laboratory diagnosis in the PR Bosna and Hercegovina (Yugoslavia)]. Veterinaria Saraj, 11:491-496

Bali MK, Singh RP, 1980. Incidence of tuberculosis and paratuberculosis of dairy cattle in HAU (Haryana Agricultural University) dairy farm. J. Res. Haryana Agric. Univ., 10(1):157-158

BANR, 2003. Control principles and programs. In: Diagnosis and Control of Johne’s Disease. Committee on Diagnosis and Control of Johne’s Disease, Board on Agriculure and Natural Resources, Division on Earth and Life Studies. Washington, DC, USA: National Academies Press, 66-67

BANR, 2003. Johne’s disease and Crohn’s disease. In: Diagnosis and Control of Johne’s Disease. Committee on Diagnosis and Control of Johne’s Disease, Board on Agriculure and Natural Resources, Division on Earth and Life Studies. Washington, DC, USA: National Academies Press, 104-120

Baradi, F., Ilchmann, G., Janetschke, P., Khaled, A.-H. Al, Rizk, G., 1972. [Occurrence of paratuberculosis in Syria.]. (Zum Vorkommen der Paratuberkulose in Syrien). Beitrage zur Tropischen und Subtropischen Landwirtschaft und Tropenveterinarmedizin, 10(heft 1), 77-82.

Benazzi S, El-Hamidi M, Schliesser T, 1996. Paratuberculosis in sheep flocks in Morocco: a serological, microscopical and cultural survey. Journal of Veterinary Medicine. Series B, 43(4):213-219; 42 ref

Bendixen PH, 1978. Immunological reactions caused by infection with M. paratuberculosis. Nordisk Veterinary Medicine, 30:163-168

Benedictus G, Dijkhuizen AA, Stelwagen J, 1987. Economic losses due to paratuberculosis in dairy cattle. Veterinary Record, 121(7):142-146; 20 ref

Benedictus G, Dinkia ETB, Wentink GM, 1988. Preliminary result of vaccination against paratuberculosis in adult dairy cattle. In: Thorel MF, Merkal, RS, eds. Second International Colloquium on Paratuberculosis. Paris. (France): 136-140

Bergey DH et al, 1923. In: Bergey's manual of determinative bacteriology, Baltimore, USA: Williams & Wilkins Co., 374-375

Billman-Jacobe H, Carrigan M, Cockram F, Corner LA, Gill IJ, Hill JF, Jessep T, Milner AR, Wood PR, 1992. A comparison of the interferon gamma assay with the absorbed ELISA for the diagnosis of Johne's disease in cattle. Australian Veterinary Journal, 69(2):25-28; 22 ref

Boelaert, F., Walravens, K., Biront, P., Vermeersch, J. P., Berkvens, D., Godfroid, J., 2000. Prevalence of paratuberculosis (Johne's disease) in the Belgian cattle population. Veterinary Microbiology, 77(3/4), 269-281. doi: 10.1016/S0378-1135(00)00312-6

Boulanger P, Gray DP, Gibbs HC, Murphy DA, 1968. A serological survey of sera from domestic animals on Easter Island. Can. J. Comp. Med., 32(2):425-429

Brees DJ, Reimer SB, Cheville NF, Florance A, Thoen CO, 2000. Immunohistochemical detection of Mycobacterium paratuberculosis in formalin-fixed, paraffin-embedded bovine tissue sections. Journal of Veterinary Diagnostic Investigation, 12(1):60-63; 12 ref

Brennan PJ, Nikaido H, 1995. The envelope of mycobacteria. Ann. Rev. Biochem., 64:29-63

Brett E, 1998. Johne's disease: an economic evaluation of control options for the New Zealand Livestock Industries. Agriculture New Zealand, Wellington

Buergelt CD, Duncan JR, 1978. Age and milk production data of cattle called from a dairy herd with PTB. JAVMA, 173:478-480

Buergelt CD, Hall C, McEntee K, Duncan JR, 1978. Pathological evaluation of paratuberculosis in naturally infected cattle. Vet. Pathol., 15(2):196-207

Burrells C, Clarke CJ, Colston A, Kay JM, Porter J, Little D, Sharp JM, 1998. A study of immunological responses of sheep clinically-affected with paratuberculosis (Johne's disease). The relationship of blood, mesenteric lymph node and intestinal lymphocyte responses to gross and microscopic pathology. Veterinary Immunology and Immunopathology, 66(3/4):343-358; 43 ref

Castro AFP, Nemoto H, 1972. Occurrence of atypical mycobacteria in the lymph nodes of apparently healthy slaughtered cattle in Sao Paulo, Brazil. Rev. Microbiol., 3(2):75-78

Cetinkaya B, Erdogan HM, Morgan KL, 1998. Prevalence, incidence and geographical distribution of Johne's disease in cattle in England and the Welsh borders. Veterinary Record, 143(10):265-269; 31 ref

Chaitaweesub P et al., 1999. Shedding of organisms and subclinical effects on production in preclinical Merino sheep affected with ovine paratuberculosis. In: Proceedings of the Sixth Intl. Colloq. Paratuberculosis, Melbourne, Australia, 1999. Intl. Assoc. Paratuberculosis, Rehoboth, MA, 126-131

Chiodini RJ, 1989. Crohn's disease and the mycobacterioses: a review and comparison of two disease entities. Clin. Microbiol. Rev., 2(1):90-117

Chiodini RJ, Davis WC, 1992. The cellular immunology of bovine paratuberculosis: the predominant response is mediated by cytotoxic gamma/delta T lymphocytes which prevent CD4 activity. Microbial Pathogenesis, 13(6):447-463; 36 ref

Chiodini RJ, Kruiningen HJvan, Merkal RS, 1984. Ruminant paratuberculosis (Johne's disease): the current status and future prospects. Cornell Veterinarian, 74(3):218-262; 330 ref

Chiodini, R. J., Kruiningen, H. J. van, 1983. Eastern white-tailed deer as a reservoir of ruminant paratuberculosis. Journal of the American Veterinary Medical Association, 182(2), 168-169.

Clarke CJ, 1997. The pathology and pathogenesis of paratuberculosis in ruminants and other species. Journal of Comparative Pathology, 116(3):217-261; 15 pp. of ref

Colgrove GS, Thoen CO, Blackburn BO, Murphy CD, 1989. Paratuberculosis in cattle: a comparison of three serologic tests with results of fecal culture. Veterinary Microbiology, 19(2):183-187; 9 ref

Collins DM, Gabric DM, Lisle GEde, 1990. Identification of two groups of Mycobacterium paratuberculosis strains by restriction endonuclease analysis and DNA hybridization. Journal of Clinical Microbiology, 28(7):1591-1596; 28 ref

Collins DM, Stephens DM, Lisle GWde, 1993. Comparison of polymerase chain reaction tests and faecal culture for detecting Mycobacterium paratuberculosis in bovine faeces. Veterinary Microbiology, 36(3/4):289-299; 20 ref

Collins MT, 2011. Diagnosis of paratuberculosis. Veterinary Clinics of North America, Food Animal Practice, 27(3):581-591. http://www.sciencedirect.com/science/article/pii/S0749072011000557

Collins MT, Sockett DC, 1993. Accuracy and economics of the USDA-licensed enzyme-linked immunosorbent assay for bovine paratuberculosis. Journal of the American Veterinary Medical Association, 203(10):1456-1463; 54 ref

Collins MT, Wells SJ, Petrini KR, Collins JE, Schultz RD, Whitlock RH, 2005. Evaluation of five antibody detection tests for diagnosis of bovine paratuberculosis. Clinical and Diagnostic Laboratory Immunology, 12(6):685-692

Corpa JM, Pérez V, Sánchez MA, García Marín JF, 2000. Control of paratuberculosis (Johne's disease) in goats by vaccination of adult animals. Veterinary Record, 146(7):195-196; 18 ref

Cousins DV, Evans RJ, Francis BR, 1995. Use of BACTEC radiometric culture method and polymerase chain reaction for the rapid screening of faeces and tissues for Mycobacterium paratuberculosis. Australian Veterinary Journal, 72(12):458-462; 13 ref

Cousins DV, Whittington R et al. , 1999. Mycobacteria distinct from Mycobacterium avium subsp. paratuberculosis isolated from the faeces of ruminants possess IS900-like sequences detectable by IS900 polymerase chain reaction: implications for diagnosis. Molecular and Cell. Probes, 14:431-442

Cranwell MP, 1993. Control of Johne's disease in a flock of sheep by vaccination. Veterinary Record, 133(9):219-220; 21 ref

Crowther RW, Polydorou K et al. , 1976. Johne's disease in sheep in Cyprus. Veterinary Record. 98:463

Das SK, Sinha RP, Chauhan HVS, 1992. Chemotherapy of paratuberculosis in goats: streptomycin, rifampicin and levamisole versus streptomycin, rifampicin and dapsone. Indian Journal of Animal Sciences, 62(1):8-13; 13 ref

Datta SCA, 1934. The situation of tuberculosis and Johne's disease in India with suggestions for the best methods to adopt for their diagnosis and control. Indian Vet. J., 11:93-98

Davidson RM, 1970. Ovine Johne's disease in New Zealand. N Z Vet. J., 18(3):28-31

Deom J, Moretelmans J, 1955. Existence de la maladie de Johne au Congo belge. [The existence of Johne's disease in the Congo belge]. Bull. Off. Int. Epizoot., 43:778-785

Dolz G, Araya LN, Suárez J, Jiménez C, 1999. Prevalence of antibodies to bovine paratuberculosis detected by a LAM-ELISA in Costa Rica. Veterinary Record, 144(12):322-323; 11 ref

Doyle TM, 1954. Isolation of Johne's bacillus from the udders of clinically affected cows. Br. Vet. J., 110:215-218

Eisenberg S, Nielen M, Hoeboer J, Bouman M, Heederik D, Koets A, 2011. Mycobacterium avium subspecies paratuberculosis in bioaerosols after depopulation and cleaning of two cattle barns. Veterinary Record, 168(22):587. http://veterinaryrecord.bvapublications.com/archive/

Eisenberg SWF, Koets AP, Nielen M, Heederik D, Mortier R, Buck Jde, Orsel K, 2011. Intestinal infection following aerosol challenge of calves with Mycobacterium avium subspecies paratuberculosis. Veterinary Research, 42(117):(3 December 2011). http://www.veterinaryresearch.org/content/42/1/117

Eisenberg SWF, Nielen M, Santema W, Houwers DJ, Heederik D, Koets AP, 2010. Detection of spatial and temporal spread of Mycobacterium avium subsp. paratuberculosis in the environment of a cattle farm through bio-aerosols. Veterinary Microbiology, 143(2/4):284-292. http://www.sciencedirect.com/science/journal/03781135

Errico F, Bermúdez J, 1983. Isolation of Mycobacterium paratuberculosis from cattle in Uruguay. Veterinaria, Uruguay, 19(83):13-16; 20 ref

Everett RE, North RN, Ottaway SJ, Scott-Orr H, 1989. The status of Johne's disease in New South Wales. Johne's disease: current trends in research, diagnosis and management., 14-18; 6 ref

Falkinham JO, Parker B C et al. , 1979. Epidemiology and ecology of the Mycobacterium avium-M. intracellulare and M. scrofulaceum (MAIS) group in the United States. In: Viallier J, ed. Colloque Intern. Sur Les Mycobacteries Atypiques, Lyon (France). Volume 1: 2-13

Fawcett AR, Goddard PJ, McKelvey WAC, Buxton D, Reid HW, Greig A, Macdonald AJ, 1995. Johne's disease in a herd of farmed red deer. Veterinary Record, 136(7):165-169; 23 ref

Fawi MT, Obeid HM, 1964. A note on Johne's disease among cattle in the Sudan. Bull. Epizoot. Dis. Afr., 12:437-438

Fraser I, 1989. The status of Johne's disease in Queensland. Johne's disease: current trends in research, diagnosis and management., 33-35; 1 ref

Fridriksdottir, V., Gunnarsson, E., Sigurdarson, S., Gudmundsdottir, K. B., 2000. Paratuberculosis in Iceland: epidemiology and control measures, past and present. Veterinary Microbiology, 77(3/4), 263-267. doi: 10.1016/S0378-1135(00)00311-4

Gallo C, Vesco G, Campo F, Haddad N, Abdelmoula H, 1989. Disease survey among goats and dromedaries in southern Tunisia. Maghreb Vétérinaire, 4(17):15-17; 3 ref

García Marín JF, Pérez V, García de Jalón JA, Heras Mde las, Barberán M, Fernández de Luco D, Badiola JJ, 1994. Diagnosis of clinical cases of paratuberculosis in sheep and goats. Medicina Veterinaria, 11(9):491.502; 45 ref

Garry F, 2011. Control of paratuberculosis in dairy herds. Veterinary Clinics of North America, Food Animal Practice, 27(3):599-607. http://www.sciencedirect.com/science/article/pii/S074907201100048X

Gasse H, 1961. L'enterite paratuberculeuse en France [Paratuberculosis enteritis in France]. Bull. Off. Int. Epizoot., 55:1132-1139

Gasteiner J, Awad-Masalmeh M, Baumgartner W, 2000. Mycobacterium avium subsp. paratuberculosis infection in cattle in Austria, diagnosis with culture, PCR and ELISA. Vet. Microbiol., 77(3-4):339-349

Gasteiner J, Wenzl H, Fuchs K, Jark U, Baumgartner W, 1999. Serological cross-sectional study of paratuberculosis in cattle in Austria. Journal of Veterinary Medicine. Series B, 46(7):457-466; 35 ref

Giessen JWBvan der, Haring RM, Vauclare E, Eger A, Haagsma J, Zeijst BAMvan der, 1992. Evaluation of the abilities of three diagnostic tests based on the polymerase chain reaction to detect Mycobacterium paratuberculosis in cattle: application in a control program. Journal of Clinical Microbiology, 30(5):1216-1219; 19 ref

Gill CO, Saucier L, Meadus WJ, 2011. Mycobacterium avium subsp. paratuberculosis in dairy products, meat, and drinking water. Journal of Food Protection, 74(3):480-499. http://www.foodprotection.org

Goodger WJ, Collins MT, Nordlund KV, Eisele C, Pelletier J, Thomas CB, Sockett DC, 1996. Epidemiologic study of on-farm management practices associated with prevalence of Mycobacterium paratuberculosis infections in dairy cattle. Journal of the American Veterinary Medical Association, 208(11):1877-1881; 18 ref

Grant IR, Ball HJ, Neill SD, Rowe MT, 1996. Inactivation of Mycobacterium paratuberculosis in cows' milk at pasteurization temperatures. Applied and Environmental Microbiology, 62(2):631-636; 29 ref

Grant IR, O'Riordan LM, Ball HJ, Rowe MT, 2001. Incidence of Mycobacterium paratuberculosis in raw sheep and goats' milk in England, Wales and Northern Ireland. Veterinary Microbiology, 79(2):123-131; 17 ref

Grewal SK, Rajeev S, Sreevatsan S, Michel FC, 2006. Persistence of Mycobacterium avium subsp. paratuberculosis and other zoonotic pathogens during simulated composting, manure packing, and liquid storage of dairy manure. Applied and Environmental Microbiology, 72(1):565-574

Gunnarsson E, 1979. Isolation of Mycobacterium paratuberculosis from sheep and cattle in Iceland. Acta Vet. Scand., 20(2):191-199

Hagan WA, 1938. Age as a factor in susceptibility to Johne's disease. Cornell Vet. 28:34-40

Hanzlíková M, Vilímek L, 1989. Prevalence of paratuberculosis in cattle in Slovakia in the period 1982-1987. Veterinárství, 39(10):462-463; [1 pl.]

Hines SA, Buergelt CD, Wilson JH, Bliss EL, 1987. Disseminated Mycobacterium paratuberculosis infection in a cow. Journal of the American Veterinary Medical Association, 190(6):681-683; 13 ref

Hoflund S, Viriden P, 1956. Paratuberkulosen eller Johne's sjukdom. En oversikt over sjukdomens upptradande I olika lander och I vart land [Paratuberculosis or Johne's disease. A survey of the occurrence of the disease in various countries and in our country]. Jord Groda Djor, 12:223-231

Hope AF, Tulk PA, Condron RJ, 1996. Commercial pasteurization of Mycobacterium paratuberculosis in whole milk. In: Proceedings of the Fifth Intl. Colloq. Paratuberculosis, Madison, WI, 1996. Intl. Assoc. Paratuberculosis, Rehoboth, MA, 377-382

Hulzen KJEvan, Heuven HCM, Nielen M, Hoeboer J, Santema WJ, Koets AP, 2011. Different Mycobacterium avium subsp. paratuberculosis MIRU-VNTR patterns coexist within cattle herds. Veterinary Microbiology, 148(2/4):419-424. http://www.sciencedirect.com/science/journal/03781135

Ilukevich A, Bermudez EC, Estrada E, 1979. Infectious diseases of cattle of growing importance in Venezuela. Cien. Vet. Maracaiba, Venezuela, 6(1):47-63

Inglies JSS, Weipers M, 1963. The effect of Johne's vaccination on the efficiency of the single intradermal comparative tuberculin test. British Veterinary Journal, 119:426-429

Iturria P, 1989. The animal health situation in the German Federal Republic. GDS Informations Sanitaires, No.96:17-30

Jakobsen MB, Alban L, Nielsen SS, 2000. A cross-sectional study of paratuberculosis in 1155 Danish dairy cows. Prev. Vet. Med., 46(1):15-27

Johne HA, Frothingham J, 1895. Ein eigenthuemlicher fall von tuberculose beim rind. Dtsch. Z. Tiermed. Pathol., 21:438-454

Johnson RH, Smith VW, Macadam I, 1962. A note on Johne's disease in Nigeria. Bull. Epizoot. Dis. Afr., 10:507-510

Johnson-Ifearulundu YJ, Kaneene JB, 1997. Relationship between soil type and Mycobacterium paratuberculosis. Journal of the American Veterinary Medical Association, 210(12):1735-1740; 44 ref

Jones DG, Kay JM, 1996. Serum biochemistry and the diagnosis of Johne's disease (paratuberculosis) in sheep. Veterinary Record, 139(20):498-499; 15 ref

Jorgensen JB, 1972. Studies of the occurrence of paratuberculosis in cattle in Denmark. Nordisk Vet. Med., 24:297-308

Jorgensen JB, 1977. Survival of Mycobacterium paratuberculosis in slurry. Nordisk Vet. Med., 29:267-270

JOrgensen JB, 1984. The effect of vaccination on the excretion of Mycobacterium paratuberculosis. Paratuberculosis, diagnostic methods, their practical application and experience with vaccination, 131-136; 11 ref

Joshi DD, Shukla RR, Pyakural S, 1974. Studies on diseases of domestic animals and birds in Nepal. III. Preliminary observations on the prevalence of tuberculosis, paratuberculosis and brucellosis at Livestock Farm, Pokhara. Bull. Vet. Sci. Anim. Husb., Nepal, 3(1):1-9

Julian RJ, 1975. A short review and some observations on Johne's disease with recommendations for control. Can. Vet. J., 16(2):33-43

Juneja LJ, Sastri NSR, Yadav BL, 1991. A study of the incidence of diseases and culling pattern in the herd of pure bred (Jersey and Holstein Friesians) cows in semi-arid region. Indian Journal of Animal Production and Management, 7(4):191-195; 4 ref

Juste RA, Badiola JJ et al. , 1991. A survey of ovine paratuberculosis in Aragon (Spain) by different methods. The Paratuberculosis Newsletter. 3:3-4

Juste RA, García Marín JF, Peris B, Sáez de Ocáriz C, Badiola JJ, 1994. Experimental infection of vaccinated and non-vaccinated lambs with Mycobacterium paratuberculosis. Journal of Comparative Pathology, 110(2):185-194; 17 ref

Juste RA, Marco JC, Sáez de Ocáriz C, Adúriz JJ, 1991. Comparison of different media for the isolation of small ruminant strains of Mycobacterium paratuberculosis.. Veterinary Microbiology, 28(4):385-390; 11 ref

Kalis CH, Hesselink JW, Barkema HW, Collins MT, 2000. Culture of strategically pooled bovine fecal samples as a method to screen herds for paratuberculosis. J. Vet. Diagn. Invest., 12(6):547-551

Kalis CHJ, Hesselink JW, Barkema HW, Collins MT, 2001. Use of long-term vaccination with a killed vaccine to prevent fecal shedding of Mycobacterium avium subsp paratuberculosis in dairy herds. American Journal of Veterinary Research, 62(2):270-274

Kazda J, 1973. Die Bedeutun von Wasser fur die Vervrteirtung von potentiel pathogenen Mydobcterien.- I: Moglichkeiten fur eine Vermehrung von Mykobakterien. Zentralbl. Bakteriol. (Orig. B), 158:161-169

Kennedy DJ, Allworth MB, 2000. Progress in national control and assurance programs for bovine Johne's disease in Australia. Veterinary Microbiology, 77(3/4):443-451; 6 ref

Keswani J, Frank JF, 1998. Thermal inactivation of Mycobacterium paratuberculosis in milk. Journal of Food Protection, 61(8):974-978; 26 ref

Khan MZ, Huq MM, 1962. Johne's disease in Pakistan. Bull. Off. Int. Epizoot., 58:7-10

Konte M, 1988. Paratuberculosis. Diagnosis of the first case in Senegal in an imported cow. Revue d'élevage et de Médecine Vétérinaire des Pays Tropicaux, 41(2):147-148; 14 ref

Kopecky KE, 1973. Distribution of bovine paratuberculosis in the United States. J. Am. Vet. Med. Assoc., 162(9):787-788

Kopecky KE, Larsen AB, Merkal RS, 1967. Uterine infection in bovine paratuberculosis. Am. J. Vet. Res., 28(125):1043-1045

Kruimingen MJvan, Chiodini RJ et al. , 1986. Experimental disease in infant goats induced by Mycobacterium paratuberculosis isolated from a patient with Crohn's disease. Digestive Diseases and Sciences, 31:1351-1360

Kujumgiev I, 1964. La paratuberculose en Bulgarie [Paratuberculosis in Bulgaria]. Bull. Off. Int. Epizoot., 61:51-66

Körmendy B, 1992. Paratuberculosis vaccine in a large dairy herd. Acta Veterinaria Hungarica, 40(3):171-184; 24 ref

Körmendy B, 1994. The effect of vaccination on the prevalence of paratuberculosis in large dairy herds. Veterinary Microbiology, 41(1/2):117-125; 23 ref

Körmendy B, Tuboly S, Nagy G, Török J, Abonyi T, 1990. Diagnostic value of intravenously administered johnin purified protein derivative (PPD) in bovine paratuberculosis. Acta Veterinaria Hungarica, 38(1/2):43-53; 22 ref

Lambrecht RS, Collins MT, 1992. Mycobacterium paratuberculosis. Factors that influence mycobactin dependence. Diagn. Microbiol. Infect. Dis., 15(3):239-246

Larsen AB, Kopecky KE, 1970. Mycobacterium paratuberculosis in reproductive organs and semen of bulls. American Journal of Veterinary Research, 31:255-258

Larsen AB, Merkal RS, Cutlip RC, 1975. Age of cattle as related to resistance to infection with Mycobacterium paratuberculosis. American Journal of Veterinary Research, 36:255-257

Larsen, A. B., Stalheim, O. H. V., Hughes, D. E., Appell, L. H., Richards, W. D., Himes, E. M., 1981. Mycobacterium paratuberculosis in the semen and genital organs of a semen-donor bull. Journal of the American Veterinary Medical Association, 179(2), 169-171.

Little D, Alzuherri MM, Clarke CJ, 1996. Phenotypic characterization of intestinal lymphocytes in ovine paratuberculosis by immunohistochemistry. Veterinary Immunology and Immunopathology, 55:175-187

Lovell R, Levi M, Francis J, 1944. Studies on the survival of Johne's bacillus. J. Comp. Pathol. Ther., 54:120-129

Lu YS, Tsai HJ, Kwang MJ, Lai SY, Lee YL, Lin DF, Lee C, 1987. Serological survey for Mycobacterium paratuberculosis infection in Taiwan. Journal of the Chinese Society of Veterinary Science, 13(1):11-15; 7 ref

Mainar-Jaime RC, Vázquez-Boland JA, 1998. Factors associated with seroprevalence to Mycobacterium paratuberculosis in small-ruminant farms in the Madrid region (Spain). Preventive Veterinary Medicine, 34(4):317-327; 26 ref

Marin JFG, Chavez G et al. , 1992. Prevalence of paratuberculosis in infected goats flocks and comparison of different methods of diagnosis. In: Chiodini RJ, Kreeger JM, eds. Proceedings of the Third International Colloquium on Paratuberculosis, Orlando (USA):157-163

McNab WB, Meek AH, Duncan JR, Martin SW, Dreumel AAvan, 1991. An epidemiological study of paratuberculosis in dairy cattle in Ontario: study design and prevalence estimates. Canadian Journal of Veterinary Research, 55(3):246-251; 30 ref

McQueen DS, Russell EG, 1979. Culture of Mycobacterium paratuberculosis from bovine fetuses. Australian Veterinary Journal, 55:203-204

Merkal RS, Larsen AB, Booth GD, 1975. Analysis of the effects of inapparent bovine paratuberculosis. Am J Res., 36:837-838

Merkal RS, Rhoades KR, Gallagher JE, Ritchie AE, 1973. Scanning electron microscopy of mycobacteria. Ann. Rev. Resp. Dis., 108:382-387

Merkal RS, Thurston JR, 1966. Comparison of Mycobacterium paratuberculosis and other mycobacteria, using standard cytochemical tests. Am. J. Vet. Res., 27(117):519-521

Merkal RS, Whipple DL, Sacks JM, Snyder GR, 1987. Prevalence of Mycobacterium paratuberculosis in ileocecal lymph nodes of cattle culled in the United States. Journal of the American Veterinary Medical Association, 190(6):676-680; 22 ref

Michel AL, Bastianello SS, 2000. Paratuberculosis in sheep: an emerging disease in South Africa. Vet. Microbiol., 77:(3-4):299-307

Millar D, Ford J, Sanderson J, Withey S, Tizard M, Doran T, Hermon-Taylor J, 1996. IS900 PCR to detect Mycobacterium paratuberculosis in retail supplies of whole pasteurized cows' milk in England and Wales. Applied and Environmental Microbiology, 62(9):3446-3452; 42 ref

Mishina D et al, 1996. On the etiology of Crohn disease. Proc. Natl. Acad. Sci., USA, 93(18):9816-9820

Mogollón GJD, Galvis ALde, Tovar AL, Murillo BN, Pena BNE, Mossos CN, 1983. Prevalence of ovine paratuberculosis in the Cundi-Boyaca tableland. Revista Instituto Colombiano Agropêcuario, 18:479-484; 21 ref

Momotani E, Whipple DL, Thiermann AB, Cheville NF, 1988. Role of M cells and macrophages in the entrance of Mycobacterium paratuberculosis into domes of ileal Peyer's patches in calves. Veterinary Pathology, 25(2):131-137; 20 ref

Moyle AI, 1975. Culture and cull procedure for control of paratuberculosis. JAVMA, 166:689-690

Muhammed SI, Ivoghli B, 1983. Diagnosis of Johne's disease in sheep by counter-immunoelectrophoresis. Trop. Anim. Health Prod., 15(1):53-57

Muskens, J., Barkema, H. W., Russchen, E., Maanen, K. van, Schukken, Y. H., Bakker, D., 2000. Prevalence and regional distribution of paratuberculosis in dairy herds in the Netherlands. Veterinary Microbiology, 77(3/4), 253-261. doi: 10.1016/S0378-1135(00)00310-2

Mutharia LM, Klassen MD, Fairles J, Barbut S, Gill CO, 2010. Mycobacterium avium subsp. paratuberculosis in muscle, lymphatic and organ tissues from cows with advanced Johne's disease. International Journal of Food Microbiology, 136(3):340-344. http://www.sciencedirect.com/science?_ob=ArticleURL&_udi=B6T7K-4XK455V-3&_user=10&_coverDate=01%2F01%2F2010&_rdoc=14&_fmt=high&_orig=browse&_srch=doc-info(%23toc%235061%232010%23998639996%231577539%23FLA%23display%23Volume)&_cdi=5061&_sort=d&_docanchor=&_ct=26&_acct=C000050221&_version=1&_urlVersion=0&_userid=10&md5=7153fa4214f1365cd6bed1e86ff6fed0

Navarro JA, Bernabé A, Gómez MA, Sánchez J, Gómez S, 1991. Mycobacterial antigen detection by immunohistochemistry in goat paratuberculosis. Journal of Veterinary Medicine. Series B, 38(3):231-237; 7 ref

Navarro JA, Ramis G, Seva J, Pallarés FJ, Sánchez J, 1998. Changes in lymphocyte subsets in the intestine and mesenteric lymph nodes in caprine paratuberculosis. Journal of Comparative Pathology, 118(2):109-121; 22 ref

Nebbia, P., Robino, P., Ferroglio, E., Rossi, L., Meneguz, G., Rosati, S., 2000. Paratuberculosis in red deer (Cervus elaphus hippelaphus) in the Western Alps. Veterinary Research Communications, 24(7), 435-443. doi: 10.1023/A:1006482520336

Nemoto H, 1961. Johne's disease in Japan. Bull. Off. Int. Epizoot., 55:1204-1213

Nielsen SS, Thamsborg SM, Houe H, Bitsch V, 2000. Bulk-tank milk ELISA antibodies for estimating the prevalence of paratuberculosis in Danish dairy herds. Prev. Vet. Med., 44(1-2):1-7

Novilla MN, 1972. Histopathologic evidence of bovine paratuberculosis (Johne's disease) in the Phillipines. Phil. J. Vet. Med., 11(1):19-23

O'Brien JJ, Baskerville A, McClelland TG, 1972. Johne's disease in sheep in Northern Ireland. Br. Vet. J., 128(7):359-365

Office International de Epizooties, 1999. [Reports to 31 December 1999]. Disease Information - Office International des Epizooties, 12(50):185-188

OIE Handistatus, 2002. World Animal Health Publication and Handistatus II (dataset for 2001). Paris, France: Office International des Epizooties

OIE Handistatus, 2003. World Animal Health Publication and Handistatus II (dataset for 2002). Paris, France: Office International des Epizooties

OIE Handistatus, 2004. World Animal Health Publication and Handistatus II (data set for 2003). Paris, France: Office International des Epizooties

OIE Handistatus, 2005. World Animal Health Publication and Handistatus II (data set for 2004). Paris, France: Office International des Epizooties

OIE, 2009. World Animal Health Information Database - Version: 1.4. World Animal Health Information Database. Paris, France: World Organisation for Animal Health. http://www.oie.int

Okwumabua O, Shull E, O'Connor M, Moua TV, Danz T, Strelow K, 2010. Comparison of three methods for extraction of Mycobacterium avium subspecies paratuberculosis DNA for polymerase chain reaction from broth-based culture systems. Journal of Veterinary Diagnostic Investigation, 22(1):67-69. http://jvdi.org/archive/

Ott SL, Wells SJ, Wagner BA, 1999. Herd-level economic losses associated with Johne's disease on US dairy operations. Preventive Veterinary Medicine, 40(3/4):179-192; 19 ref

Paily EP, Georgekutty PT, Venugopal K, 1979. Incidence of Johne's disease in cattle and buffaloes in livestock farms of the Kerala Agricultural University. Kerala J. Vet. Sci., 10(2):344-345

Paling RW, Waghela S, Macowan KJ, Heath BR, 1988. The occurrence of infectious diseases in mixed farming of domesticated wild herbivores and livestock in Kenya. II. Bacterial diseases. Journal of Wildlife Diseases, 24(2):308-316; 30 ref

Pande PG, 1940. Paratuberculosis (Johne's disease) of cattle in Assam. 1. It's incidence and epizootiology. Indian J. Vet. Sci., 10:40-62

Pandey GS, Musonda TL, Chizyuka HGB, Schneebeli M, 1987. Paratuberculosis (Johne's disease) in a herd of Friesian cattle in Zambia. Veterinary Record, 120(15):369; 6 ref

Patterson DSP, Allen Wm, Berrett S, 1965. Plasma enzyme changes in clinical Johne's disease. Vet. Rec., 77(44):1287-1289

Patterson DSP, Allen WM, Berrett S, 1968. Some biochemical aspects of Johne's disease in cattle. Res. Vet. Sci., 9(2):117-129

Patterson DSP, Allen WM, Lloyd MK, 1967. Clinical Johne's disease as a protein losing enteropathy. Vet. Rec., 81(17):435-437

Pavlík I, Bejcková L, Pavlas M, Rozsypalová Z, Kosková S, 1995. Characterization by restriction endonuclease analysis and DNA hybridization using IS900 of bovine, ovine, caprine and human dependent strains of Mycobacterium paratuberculosis isolated in various localities. Veterinary Microbiology, 45(4):311-318; 15 ref

Pavlik, I., Bartl, J., Dvorska, L., Svastova, P., Maine, R. du, Machackova, M., Ayele, W. Y., Horvathova, A., 2000. Epidemiology of paratuberculosis in wild ruminants studied by restriction fragment length polymorphism in the Czech Republic during the period 1995-1998. Veterinary Microbiology, 77(3/4), 231-251. doi: 10.1016/S0378-1135(00)00309-6

Payne JM, Rankin JD, 1961. A comparison of the pathogenesis of experimental Johne's disease in calves and cows. Research in Veterinary Science, 2:175-179

Pérez V, García Marín JF, Badiola JJ, 1996. Description and classification of different types of lesion associated with natural paratuberculosis infection in sheep. Journal of Comparative Pathology, 114(2):107-122; [26 ref]

Pérez V, García Marín JF, Bru R, Moreno B, Badiola JJ, 1995. Results of a paratuberculosis vaccination trial in adult sheep. Medicina Veterinaria, 12(3):196-201; 33 ref

Pérez V, Tellechea J, Badiola JJ, Gutiérrez M, García MJF, 1997. Relation between serologic response and pathologic findings in sheep with naturally acquired paratuberculosis. American Journal of Veterinary Research, 58(8):799-803; 32 ref

Pérez V, Tellechea J, Corpa JM, Gutiérrez M, García Marín JF, 1999. Relation between pathologic findings and cellular immune responses in sheep with naturally acquired paratuberculosis. American Journal of Veterinary Research, 60(1):123-127; 31 ref

Philpott M, 1993. The dangers of disease transmission by artificial insemination and embryo transfer. British Veterinary Journal, 149(4):339-369; 198 ref

Pitt DJ, 1989. The status of Johne's disease in Tasmania. Johne's disease: current trends in research, diagnosis and management., 28-32

Portugal, M. A. S. C., Pimentel, J. N., Saliba, A. M., Baldassi, L., Sandoval, E. F. D., 1979. Occurrence of paratuberculosis in cattle in Santa Catarina State, Brazil. (Ocorrencia de paratuberculose no Estado de Santa Catarina). Biologico, 45(1/2), 19-24.

Power SB, Haagsma J, Smyth DP, 1993. Paratuberculosis in farmed red deer (Cervus elaphus) in Ireland. Veterinary Record, 132(9):213-216; 19 ref

Pradhan AK, Mitchell RM, Kramer AJ, Zurakowski MJ, Fyock TL, Whitlock RH, Smith JM, Hovingh E, Kessel JASvan, Karns JS, Schukken YH, 2011. Molecular epidemiology of Mycobacterium avium subsp. paratuberculosis in a longitudinal study of three dairy herds. Journal of Clinical Microbiology, 49(3):893-901. http://jcm.asm.org/cgi/content/short/49/3/893

Raizman EA, Wells SJ, Godden SM, Bey RF, Oakes MJ, Bentley DC, Olsen KE, 2004. The distribution of Mycobacterium avium ssp. paratuberculosis in the environment surrounding Minnesota dairy farms. American Dairy Science Association, 87:2959-2966

Ramirez PC, Trigo TE, Suarez GF, Merkal R, 1979. Isolation and identification of Mycobacterium paratuberculosis in Mexico. Tec. Pecu. Mex., 36:74-76

Ramisz A, 1970. Przypadek choroby Johnego u bydla na terenie Wojewodztwa Krakowskiego [An outbreak of Johne's disease in Krakow province (Poland)]. Med. Weteryn., 26:203-206

Ramos Saco T, 1953. La paratuberculosis bovis (enfermedad de Johne o enteritis paratuberculosa), nueve enfermedad halada en el ganado lechero en el Peru. Nota preliminar [Bovine paratuberculosis (Johne's disease or paratuberculosis enteritis), a new disease of cattle in Peru. Preliminary note]. Vida. Agric. (Lima), 30:829-830

Rankin JD, 1955. An attempt to prevent the establishment of Mycobacterium johneii in calves by means of isoniazid alone or in combination with streptomycin. Vet. Rec., 67:1105-1107

Reddacliff L, Eppleston J, Windsor P, Whittington R, Jones S, 2006. Efficacy of a killed vaccine for the control of paratuberculosis in Australian sheep flocks. Vet Microbiol. Accessed 5 April 2006: [Epub ahead of print]

Rihal R, 1979. Chronic diarrhoe of cattle in the Normandy region. Bull. Group Techn. Vet., 3:23-26

Rohde RF, Shulaw WP, 1990. Isolation of Mycobacterium paratuberculosis from the uterine flush fluids of cows with clinical paratuberculosis. Journal of the American Veterinary Medical Association, 197(11):1482-1483; 6 ref

Rohde RF, Shulaw WP, Hueston WD, Bech-Nielsen S, Haibel GK, Hoffsis GF, 1990. Isolation of Mycobacterium paratuberculosis from washed bovine ova after in vitro exposure. American Journal of Veterinary Research, 51(5):708-710; 14 ref

Rossiter CA, 1996. Approach for designing appropriate Johne's disease control programs on the farm. In: Proceedings of the Bovine Practioner, Vol. 29. Stillwater, OK, USA: Frontier Printers, Inc., 33-45

Salman MD, Hernandez JA, Braun I, 1990. A seroepidemiological study of five bovine diseases in dairy farms of the coastal region of Baja California, Mexico. Preventive Veterinary Medicine, 9(2):143-153; 34 ref

Sanderson JD, Moss MT, Tizard ML, Hermon-Taylor J, 1992. Mycobacterium paratuberculosis DNA in Crohn's disease tissue. Gut, 33(7):890-896

Sanftleben P, 1990. Quest continues for fast, reliable test for bovine paratuberculosis. J. Am. Vet. Med. Assoc., 197(3):299-305

Santos JA, da Silva NL, 1956. Sobre a la observacao de paratuberculose no Brazil [On the first observations of paratuberculosis in Brazil]. Bolm. Soc. Bras. Med. Vet., 24:5-14

Saxegaard F, Fodstad FH, 1985. Control of paratuberculosis (Johne's disease) in goats by vaccination. Veterinary Record, 116(16):439-441; 26 ref

Schaik Gvan, Kalis CHJ, Benedictus G, Dijkhuizen AA, Huirne RBM, 1996. Cost-benefit analysis of vaccination against paratuberculosis in dairy cattle. Veterinary Record, 139(25):624-627; 10 ref

Schaik Gvan, Pradenas F M, Mella N A, Kruze V J, 2007. Diagnostic validity and costs of pooled fecal samples and individual blood or fecal samples to determine the cow- and herd-status for Mycobacterium avium subsp. paratuberculosis. Preventive Veterinary Medicine, 82(1/2):159-165. http://www.sciencedirect.com/science/journal/01675877

Seitz SE, Heider LE, Hueston WD, Bech-Nielsen S, Rings DM, Spangler L, 1989. Bovine fetal infection with Mycobacterium paratuberculosis.. Journal of the American Veterinary Medical Association, 194(10):1423-1426; 21 ref

Sevilla I, Li LL, Amonsin A, Garrido JM, Geijo MV, Kapur V, Juste RA, 2008. Comparative analysis of Mycobacterium avium subsp. paratuberculosis isolates from cattle, sheep and goats by short sequence repeat and pulsed-field gel electrophoresis typing. BMC Microbiology, 8(204):(25 November 2008). http://www.biomedcentral.com/1471-2180/8/204

Sharma DR, Kwatra MS, Kumar A, 1985. Prevalence of Johne's disease in sheep breeding farms in Punjab. Journal of Research, Punjab Agricultural University, 22(3):579-580; 10 ref

Shulaw WP, Bech-Nielsen S, Rings DM, Woodruff TS, 1993. Serodiagnosis of paratuberculosis in sheep by use of agar gel immunodiffusion. American Journal of Veterinary Research, 54(1):13-19; 28 ref

Sigurdsson B, 1960. A killed vaccine against paratuberculosis (Johne's disease) in sheep. American Journal of Veterinary Research, 21:54-67

Slocombe RF, 1982. Combined streptomycin-isoniazid-rifampicin therapy in the treatment of Johne's disease in a goat. Can. Vet. J., 23(5):160-163

Sockett DC, Carr DJ, Collins MT, 1992. Evaluation of conventional and radiometric fecal culture and a commercial DNA probe for diagnosis of Mycobacterium paratuberculosis infection in cattle. Canadian Journal of Veterinary Research, 56(2):148-153; 32 ref

Spahr U, Schafroth K, 2001. Fate of Mycobacterium avium subsp paratuberculosis in Swiss hard and semihard cheese manufactured from raw milk. Applied and Environmental Microbiology, 67(9):4199-4205

Spangler E, Bech-Nielsen S, Heider LE, 1992. Diagnostic performance of two serologic tests and fecal culture for subclinical paratuberculosis, and associations with production. Preventive Veterinary Medicine, 13(3):185-195; 27 ref

Spangler E, Heider LE, Bech-Nielsen S, Dorn CR, 1991. Serologic enzyme-linked immunosorbent assay responses of calves vaccinated with a killed Mycobacterium paratuberculosis vaccine. American Journal of Veterinary Research, 52(8):1197-1200; 11 ref

Srivastava AK, More BK, 1987. Prevalence of Johne's disease in local, Angora and crossbred goats in Maharashtra. Livestock Adviser, 12(2):33-37; 19 ref

Stabel JR, 1996. Cytokine secretion by peripheral blood mononuclear cells from cows infected with Mycobacterium paratuberculosis. Am. J. Vet. Res., 61(7):754-760

Stabel JR, Steadham EM, Bolin CA, 1997. Heat inactivation of Mycobacterium paratuberculosis in raw milk: are current pasteurization conditions effective?. Applied and Environmental Microbiology, 63(12):4975-4977; 13 ref

Stamp JT, Watt JA, 1954. Johne's disease in sheep. Journal of Comparative Pathology, 64:26-40

Stehman SM, 1996. Paratuberculosis in small ruminants, deer, and South American camelids. Veterinary Clinics of North America, Food Animal Practice, 12(2):441-455; 56 ref

StJean G, 1996. Treatment of clinical paratuberculosis in cattle. Veterinary Clinics of North America, Food Animal Practice, 12(2):417-430; 51 ref

Streeter RN, Hoffsis GF, Bech-Nielsen S, Shulaw WP, Rings M, 1995. Isolation of Mycobacterium paratuberculosis from colostrum and milk of subclinically infected cows. American Journal of Veterinary Research, 56(10):1322-1324; 23 ref

Sung NackMoon, Collins MT, 1998. Thermal tolerance of Mycobacterium paratuberculosis. Applied and Environmental Microbiology, 64(3):999-1005; 34 ref

Sweeney RW, Whitlock RH, Buckley CL, Spencer PA, 1995. Evaluation of a commercial enzyme-linked immunosorbent assay for the diagnosis of paratuberculosis in dairy cattle. Journal of Veterinary Diagnostic Investigation, 7(4):488-493; 15 ref

Sweeney RW, Whitlock RH, Rosenberger AE, 1992. Mycobacterium paratuberculosis cultured from milk and supramammary lymph nodes of infected asymptomatic cows. Journal of Clinical Microbiology, 30(1):166-171; 21 ref

Sweeney RW, Whitlock RH, Rosenberger AE, 1992. Mycobacterium paratuberculosis isolated from fetuses of infected cows not manifesting signs of the disease. American Journal of Veterinary Research, 53(4):477-480; 18 ref

Talatchian M, 1965. First report of Johne's disease in Iran. Bull. Off. Int. Epizoot., 64:779-782

Taylor TK, Wilks CR, McQueen DS, 1981. Isolation of Mycobacterium paratuberculosis from milk of cows with Johne's disease. Veterinary Record, 109:532-533

Temple, R. M. S., Muscoplat, C. C., Thoen, C. O., Himes, E. M., Johnson, D. W., 1979. Observations on diagnostic tests for paratuberculosis in a deer herd. Journal of the American Veterinary Medical Association, 175(9), 914-915.

Thoen CO, 1990. Mycobacterium. In: Carter GR, Cole J, eds. Diagnostic procedures in veterinary bacteriology and mycology. Fifth edition. San Diego, California, USA: Academic Press Inc., 287-298

Thoen CO, Haagsma J, 1996. Molecular techniques in the diagnosis and control of paratuberculosis in cattle. Journal of the American Veterinary Medical Association, 209(4):734-737; 35 ref

Thoen CO, Moore LA, 1989. Control of Johne's disease in four commercial dairy herds in Iowa. Journal of Veterinary Diagnostic Investigation, 1(3):223-226; 20 ref

Thorel MF, Krichevsky M, Lévy-Frébault VV, 1990. Numerical taxonomy of mycobactin-dependent mycobacteria, emended description of Mycobacterium avium, description of Mycobacterium avium subsp. avium subsp. nov., Mycobacterium avium subsp. paratuberculosis subsp. nov., and Mycobacterium avium subsp. silvaticum subsp. nov. International Journal of Systematic Bacteriology, 40(3):254-260; 53 ref

Tsellarius IK, 1969. Vivcenija paratuberkul'ozu velikoj rogatoj chudobi na Ukrajni [Bovine paratuberculosis in the Ukraine]. Vertinariya (Kiev), 24:31-37

Twort F, 1910. A method for isolating and growing the lepra bacillus of man (preliminary note). Proc. R. Soc. Lond., Series B, 83:156-158

Twort FW, Ingram GLY, 1912. A method for isolating and cultivating Mycobacterium enteriditis chronicae pseudotuberculosis bovis, Johne, and some experiments on the preparation of a diagnostic vaccine for pseudotuberculosis enteritis of bovines. Proc. R. Soc. Lond., Series B, 84:517-542

Ubach FA, 1941. Observations on Johne's disease and coccidial enteritis in cattle, sheep and goats in the Republic of Argentina. Revta. Med. Vet. Buenos Aires, 23:3-45

Vandegraaff R, 1989. The status of Johne's disease in South Australia. Johne's disease: current trends in research, diagnosis and management., 19-24; 5 ref

VanLeeuwen, J. A., Keefe, G. P., Tremblay, R., Power, C., Wichtel, J. J., 2001. Seroprevalence of infection with Mycobacterium avium subspecies paratuberculosis, bovine leukemia virus, and bovine viral diarrhea virus in Maritime Canada dairy cattle. Canadian Veterinary Journal, 42(3), 193-198.

Vary PH et al, 1990. Use of highly specific DNA probes and the polymerase chain reaction to detect Mycobacterium paratuberculosis in Johne's disease. J. Clin. Microbiol., 28(5):933-937

Vavako D, 1942. L'enterite paratubercolare dei bovini in Albania [Bovine paratuberculosis enteritis in Albania]. Clinica. Vet. Milano, 65:291-293

Vural B, Atala N, 1988. Serological study on bovine paratuberculosis in Central Anatolia using the micro-complement fixation and tube complement fixation tests. Etlik Veteriner Mikrobiyoloji Enstitütü Dergisi, 6(3):87-97; 28 ref

Wards BJ, Collins DM, Lisle GWde, 1995. Detection of Mycobacterium bovis in tissues by polymerase chain reaction. Veterinary Microbiology, 43(2/3):227-240; 41 ref

Weering Hvan, Schaik Gvan, Meulen Avan der, Waal M, Franken P, Maanen Kvan, 2007. Diagnostic performance of the Pourquier ELISA for detection of antibodies against Mycobacterium avium subspecies paratuberculosis in individual milk and bulk milk samples of dairy herds. Veterinary Microbiology, 125(1/2):49-58. http://www.sciencedirect.com/science/journal/03781135

Wells SJ, Wagner BA, 2000. Herd-level risk factors for infection with Mycobacterium paratuberculosis in US dairies and association between familiarity of the herd manager with the disease or prior diagnosis of the disease in that herd and use of preventive measures. J. Am. Vet. Med. Assoc., 216(9):1450-1457

Wells SJ, Whitlock RH, Lindeman CJ, Fyock T, 2002. Evaluation of bacteriologic culture of pooled faecal samples for detection of Mycobacterium paratuberculosis. American Journal of Veterinary Research, 63(8):1207-1211; 17 ref

Wentink GH, Bongers JH, Zeeuwen AAPA, Jaartsveld FHJ, 1994. Incidence of paratuberculosis after vaccination against M. paratuberculosis in two infected dairy herds. Journal of Veterinary Medicine. Series B, 41(7/8):517-522; 12 ref

Whipple D, Kapke P, Vary C, 1990. Identification of restriction fragment length polymorphisms in DNA from Mycobacterium paratuberculosis.. Journal of Clinical Microbiology, 28(11):2561-2564; 19 ref

Whipple DL, Kapke PA, Andersen PR, 1992. Comparison of a commercial DNA probe test and three cultivation procedures for detection of Mycobacterium paratuberculosis in bovine faeces. J Vet Diagn Invest, 4:23-27

Whitlock RH, Buergelt C, 1996. Preclinical and clinical manifestations of paratuberculosis (including pathology). Veterinary Clinics of North America, Food Animal Practice, 12(2):345-356; 39 ref

Whitlock RH, van Buskirk M, Sweeney RW, 1994. Pennsylvannia Johne's disease control program (1973-1993): a review of the twenty year program. In: Proceedings of the Fourth Intl. Colloq. Paratuberculosis, Cambridge, UK, 1994. Intl. Assoc. Paratuberculosis, Rehoboth, MA, 47-53

Whittington RJ et al, 1998. Rapid detection of Mycobacterium paratuberculosis in clinical samples from ruminants and in spiked environmental samples by modified BACTEC 12B radiometric culture and direct confirmation by IS900 PCR. J. Clin. Microbiol., 36(3):701-707

Whittington RJ, Fell S, Walker D, McAllister S, Marsh I, Sergeant E, Taragel CA, Marshall DJ, Links IJ, 2000. Use of pooled fecal culture for sensitive and economic detection of Mycobacterium avium subsp. paratuberculosis infection in flocks of sheep. Journal of Clinical Microbiology, 38(7):2550-2556; 26 ref

Whittington RJ, Waldron A, Warne D, 2010. Thermal inactivation profiles of Mycobacterium avium subsp. paratuberculosis in lamb skeletal muscle homogenate fluid. International Journal of Food Microbiology, 137(1):32-39. http://www.sciencedirect.com/science?_ob=ArticleURL&_udi=B6T7K-4XH5MD0-1&_user=10&_coverDate=01%2F31%2F2010&_rdoc=5&_fmt=high&_orig=browse&_srch=doc-info(%23toc%235061%232010%23998629998%231577606%23FLA%23display%23Volume)&_cdi=5061&_sort=d&_docanchor=&_ct=18&_acct=C000050221&_version=1&_urlVersion=0&_userid=10&md5=c7655a47fba42d258405be2c1dc0485c

Williams AG, Withers SE, 2010. Microbiological characterisation of artisanal farmhouse cheeses manufactured in Scotland. International Journal of Dairy Technology, 63(3):356-369. http://www.blackwell-synergy.com/loi/idt

Wilson DJ, Rossiter C, Han HR, Sears PM, 1993. Association of Mycobacterium paratuberculosis infection with reduced mastitis, but with decreased milk production and increased cull rate in clinically normal dairy cows. American Journal of Veterinary Research, 54(11):1851-1857; 15 ref

Withers FW, 1959. Incidence of the disease. Vet. Rec., 71:1150-1153

Wood PR, Corner LA, Plackett P, 1990. Development of a simple, rapid in vitro cellular assay for bovine tuberculosis based on the production of interferon. Research in Veterinary Science, 49(1):46-49; 21 ref

Xie X, Duan XW, Zhou Q, Gui HQ, 1982. Isolation and cultivation of Mycobacterium johnei from cattle for the first time in China [In Chinese]. Chin. J. Vet. Med., 8(10):2-3

Zamora J, Kruze J, Schifferli C, 1975. Johne's disease of sheep: first case described in Chile. Arch. Med. Vet., 7(1):15-17

Zhong L, Fiore Ldi, Taylor D, Begg D, Silva Kde, Whittington RJ, 2009. Identification of differentially expressed genes in ileum, intestinal lymph node and peripheral blood mononuclear cells of sheep infected with Mycobacterium avium subsp. paratuberculosis using differential display polymerase chain reaction. Veterinary Immunology and Immunopathology, 131(3/4):177-189. http://www.sciencedirect.com/science/journal/01652427

Distribution References

Albiston HE, Talbot RJ, 1936. Johne's disease in Victoria. In: Aust. Vet. J. 12 125-138.

Allworth M B, Kennedy D J, 2000. Progress in national control and assurance programs for ovine Johne's disease in Australia. Veterinary Microbiology. 77 (3/4), 415-422. DOI:10.1016/S0378-1135(00)00326-6

Badnjevic B, Forsek Z, 1968. Bovine and ovine paratuberculosis in certain regions of Yugoslavia. In: Bull. Off. Int. Epizoot. 70 (1) 701-706.

Badnjevic B, Forsek Z, Kovacevic E, 1962. (Johne-ova bolest (paratuberkulosa) kod goveda. Prva laboratorijska dijagnoza u Bosni I Hercegovini [Johne's disease (paratuberculosis) in cattle. First laboratory diagnosis in the PR Bosna and Hercegovina (Yugoslavia)]). In: Veterinaria Saraj, 11 491-496.

Bali MK, Singh RP, 1980. Incidence of tuberculosis and paratuberculosis of dairy cattle in HAU (Haryana Agricultural University) dairy farm. In: J. Res. Haryana Agric. Univ. 10 (1) 157-158.

Baradi F, Ilchmann G, Janetschke P, Khaled A-H Al, Rizk G, 1972. [Occurrence of paratuberculosis in Syria.]. (Zum Vorkommen der Paratuberkulose in Syrien.). Beitrage zur Tropischen und Subtropischen Landwirtschaft und Tropenveterinarmedizin. 10 (heft 1), 77-82.

Boelaert F, Walravens K, Biront P, Vermeersch J P, Berkvens D, Godfroid J, 2000. Prevalence of paratuberculosis (Johne's disease) in the Belgian cattle population. Veterinary Microbiology. 77 (3/4), 269-281. DOI:10.1016/S0378-1135(00)00312-6

Boulanger P, Gray D P, Gibbs H C, Murphy D A, 1968. A serological survey of sera from domestic animals on Easter Island. Canadian Journal of Comparative Medicine. 425-429.

CABI, Undated. Compendium record. Wallingford, UK: CABI

CABI, Undated a. CABI Compendium: Status as determined by CABI editor. Wallingford, UK: CABI

Castro A F P de, Nemoto H, 1972. Occurrence of atypical mycobacteria in the lymph nodes of apparently healthy slaughtered cattle in Sao Paulo, Brazil. Revista de Microbiologia. 3 (No.2), 75-78.

Çetinkaya B, Erdogan H M, Morgan K L, 1998. Prevalence, incidence and geographical distribution of Johne's disease in cattle in England and the Welsh borders. Veterinary Record. 143 (10), 265-269.

Datta SCA, 1934. The situation of tuberculosis and Johne's disease in India with suggestions for the best methods to adopt for their diagnosis and control. In: Indian Vet. J. 11 93-98.

Davidson RM, 1970. Ovine Johne's disease in New Zealand. In: N Z Vet. J. 18 (3) 28-31.

Dolz G, Araya L N, Suárez J, Jiménez C, 1999. Prevalence of antibodies to bovine paratuberculosis detected by a LAM-ELISA in Costa Rica. Veterinary Record. 144 (12), 322-323.

Errico F, Bermúdez J, 1983. Isolation of Mycobacterium paratuberculosis from cattle in Uruguay. (Aislamiento de Micobacterium paratuberculosis (M. johnei) en bovinos en el Uruguay.). Veterinaria, Uruguay. 19 (83), 13-16.

Everett R E, North R N, Ottaway S J, Scott-Orr H, 1989. The status of Johne's disease in New South Wales. In: Johne's disease: current trends in research, diagnosis and management. [ed. by Milner AR, Wood PR]. East Melbourne, Vic. 3002, Australia: CSIRO Publications. 14-18.

Fawi M T, Obeid H M, 1964. A note on Johne's disease among cattle in the Sudan. Bulletin of epizootic diseases of Africa. 437-438.

Fraser I, 1989. The status of Johne's disease in Queensland. In: Johne's disease: current trends in research, diagnosis and management. [ed. by Milner AR, Wood PR]. East Melbourne, Vic. 3002 CSIRO Publications, Australia: 33-35.

Gallo C, Vesco G, Campo F, Haddad N, Abdelmoula H, 1989. Disease survey among goats and dromedaries in southern Tunisia. (Enquête zoosanitaire chez les chèvres et les dromadaires au sud de la Tunisie.). Maghreb Vétérinaire. 4 (17), 15-17.

Gunnarsson E, 1979. Isolation of Mycobacterium paratuberculosis from sheep and cattle in Iceland. Acta Veterinaria Scandinavica. 20 (2), 191-199.

Hanzlíková M, Vilímek L, 1989. Prevalence of paratuberculosis in cattle in Slovakia in the period 1982-1987. (Výskyt paratuberkulózy u hovädzieko dobytka v SSR v rokoch 1982-1987.). Veterinářství. 39 (10), 462-463.

Hoflund S, Viriden P, 1956. Paratuberculosis or Johne's disease. A survey of the occurrence of the disease in various countries and in our country. (Paratuberkulosen eller Johne's sjukdom. En oversikt over sjukdomens upptradande I olika lander och I vart land). In: Jord Groda Djor, 12 223-231.

Iturria P, 1989. The animal health situation in the German Federal Republic. (Á la rencontre de nos partenaires européens: l'Allemagne.). GDS Informations Sanitaires. 17-30.

Joshi DD, Shukla RR, Pyakural S, 1974. Studies on diseases of domestic animals and birds in Nepal. III. Preliminary observations on the prevalence of tuberculosis, paratuberculosis and brucellosis at Livestock Farm, Pokhara. In: Bull. Vet. Sci. Anim. Husb., Nepal, 3 (1) 1-9.

Juneja L J, Sastri N S R, Yadav B L, 1991. A study of the incidence of diseases and culling pattern in the herd of pure bred (Jersey and Holstein Friesians) cows in semi-arid region. Indian Journal of Animal Production and Management. 7 (4), 191-195.

Kalis C H J, Hesselink J W, Barkema H W, Collins M T, 2000. Culture of strategically pooled bovine fecal samples as a method to screen herds for paratuberculosis. Journal of Veterinary Diagnostic Investigation. 12 (6), 547-551.

Kennedy D J, Allworth M B, 2000. Progress in national control and assurance programs for bovine Johne's disease in Australia. Veterinary Microbiology. 77 (3/4), 443-451. DOI:10.1016/S0378-1135(00)00329-1

Khan MZ, Huq MM, 1962. Johne's disease in Pakistan. In: Bull. Off. Int. Epizoot. 58 7-10.

Kujumgiev I, 1964. Paratuberculosis in Bulgaria. (La paratuberculose en Bulgarie). In: Bull. Off. Int. Epizoot. 61 51-66.

Lu Y S, Tsai H J, Kwang M J, Lai S Y, Lee Y L, Lin D F, Lee C, 1987. Serological survey for Mycobacterium paratuberculosis infection in Taiwan. Journal of the Chinese Society of Veterinary Science. 13 (1), 11-15.

McNab W B, Meek A H, Duncan J R, Martin S W, Dreumel A A van, 1991. An epidemiological study of paratuberculosis in dairy cattle in Ontario: study design and prevalence estimates. Canadian Journal of Veterinary Research. 55 (3), 246-251.

Merkal R S, Whipple D L, Sacks J M, Snyder G R, 1987. Prevalence of Mycobacterium paratuberculosis in ileocecal lymph nodes of cattle culled in the United States. Journal of the American Veterinary Medical Association. 190 (6), 676-680.

Mogollón G J D, Galvis A L de, Tovar A L, Murillo B N, Peña B N E, Mossos C N, 1983. Prevalence of ovine paratuberculosis in the Cundi-Boyaca tableland. (Prevalencia de paratuberculosis ovina en el altiplano Cundi-Boyacense.). Revista Instituto Colombiano Agropêcuario. 479-484.

Muhammed SI, Ivoghli B, 1983. Diagnosis of Johne's disease in sheep by counter-immunoelectrophoresis. In: Trop. Anim. Health Prod. 15 (1) 53-57.

Nemoto H, 1961. Johne's disease in Japan. In: Bull. Off. Int. Epizoot. 55 1204-1213.

Nielsen S S, Thamsborg S M, Houe H, Bitsch V, 2000. Bulk-tank milk ELISA antibodies for estimating the prevalence of paratuberculosis in Danish dairy herds. Preventive Veterinary Medicine. 44 (1/2), 1-7. DOI:10.1016/S0167-5877(00)00098-2

OIE Handistatus, 2005. World Animal Health Publication and Handistatus II (dataset for 2004)., Paris, France: Office International des Epizooties.

OIE, 2009. World Animal Health Information Database - Version: 1.4., Paris, France: World Organisation for Animal Health. https://www.oie.int/

Paily EP, Georgekutty PT, Venugopal K, 1979. Incidence of Johne's disease in cattle and buffaloes in livestock farms of the Kerala Agricultural University. In: Kerala J. Vet. Sci. 10 (2) 344-345.

Paling R W, Waghela S, Macowan K J, Heath B R, 1988. The occurrence of infectious diseases in mixed farming of domesticated wild herbivores and livestock in Kenya. II. Bacterial diseases. Journal of Wildlife Diseases. 24 (2), 308-316.

PANDE P G, 1940. Paratuberculosis (Johne's Disease) of Cattle in Assam. I. Its Incidence and Epizootiology. Indian Journal of Veterinary Science. 40-62.

Pandey G S, Musonda T L, Chizyuka H G B, Schneebeli M, 1987. Paratuberculosis (Johne's disease) in a herd of Friesian cattle in Zambia. Veterinary Record. 120 (15), 369.

Pavlik I, Bartl J, Dvorska L, Svastova P, Maine R du, Machackova M, Ayele W Y, Horvathova A, 2000. Epidemiology of paratuberculosis in wild ruminants studied by restriction fragment length polymorphism in the Czech Republic during the period 1995-1998. Veterinary Microbiology. 77 (3/4), 231-251. DOI:10.1016/S0378-1135(00)00309-6

Pitt D J, 1989. The status of Johne's disease in Tasmania. In: Johne's disease: current trends in research, diagnosis and management. [ed. by Milner AR, Wood PR]. East Melbourne, Vic. 3002, Australia: CSIRO Publications. 28-32.

Portugal M A S C, Pimentel J N, Saliba A M, Baldassi L, Sandoval E F D, 1979. Occurrence of paratuberculosis in cattle in Santa Catarina State, Brazil. (Ocorrencia de paratuberculose no Estado de Santa Catarina.). Biologico. 45 (1/2), 19-24.

Ramirez PC, Trigo TE, Suarez GF, Merkal R, 1979. Isolation and identification of Mycobacterium paratuberculosis in Mexico. In: Tec. Pecu. Mex. 36 74-76.

Ramisz A, 1970. An outbreak of Johne's disease in Krakow province (Poland). (Przypadek choroby Johnego u bydla na terenie Wojewodztwa Krakowskiego). In: Med. Weteryn. 26 203-206.

Ramos Saco T, 1953. Bovine paratuberculosis (Johne's disease or paratuberculosis enteritis), a new disease of cattle in Peru. Preliminary note. (La paratuberculosis bovis (enfermedad de Johne o enteritis paratuberculosa), nueve enfermedad halada en el ganado lechero en el Peru. Nota preliminar). In: Vida. Agric. 30 Lima, 829-830.

Salman M D, Hernandez J A, Braun I, 1990. A seroepidemiological study of five bovine diseases in dairy farms of the coastal region of Baja California, Mexico. Preventive Veterinary Medicine. 9 (2), 143-153. DOI:10.1016/0167-5877(90)90032-D

Saxegaard F, Fodstad F H, 1985. Control of paratuberculosis (Johne's disease) in goats by vaccination. Veterinary Record. 116 (16), 439-441.

Sharma D R, Kwatra M S, Kumar A, 1985. Prevalence of Johne's disease in sheep breeding farms in Punjab. Journal of Research, Punjab Agricultural University. 22 (3), 579-580.

Srivastava A K, More B K, 1987. Prevalence of Johne's disease in local, Angora and crossbred goats in Maharashtra. Livestock Adviser. 12 (2), 33-37.

Talatchian M, 1965. First report of Johne's disease in Iran. In: Bull. Off. Int. Epizoot. 64 779-782.

Tsellarius IK, 1969. Bovine paratuberculosis in the Ukraine. (Vivcenija paratuberkul'ozu velikoj rogatoj chudobi na Ukrajni). In: Vertinariya (Kiev), 24 31-37.

Ubach FA, 1941. Observations on Johne's disease and coccidial enteritis in cattle, sheep and goats in the Republic of Argentina. In: Revta. Med. Vet. Buenos Aires, 23 3-45.

Vandegraaff R, 1989. The status of Johne's disease in South Australia. In: Johne's disease: current trends in research, diagnosis and management. [ed. by Milner AR, Wood PR]. East Melbourne, Vic. 3002, Australia: CSIRO Publications. 19-24.

Vavako D, 1942. Bovine paratuberculosis enteritis in Albania. (L'enterite paratubercolare dei bovini in Albania). In: Clinica. Vet. Milano, 65 291-293.

Xie X, Duan X W, Zhou Q, Gui H Q, 1982. Isolation and cultivation of Mycobacterium johnei from cattle for the first time [? in China]. Chinese Journal of Veterinary Medicine (Zhongguo Shouyi Zazhi). 8 (10), 2-3.

Zamora J, Kruze J, Schifferli C, 1975. Johne's disease of sheep: first case described in Chile. In: Arch. Med. Vet. 7 (1) 15-17.

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