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paratuberculosis

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paratuberculosis

Summary

  • Last modified
  • 16 October 2018
  • Datasheet Type(s)
  • Animal Disease
  • Preferred Scientific Name
  • paratuberculosis
  • 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 pa...

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Pictures

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PictureTitleCaptionCopyright
Cattle with end-stage clinical Johne's disease, showing signs of severe emaciation.
TitleSymptoms
CaptionCattle with end-stage clinical Johne's disease, showing signs of severe emaciation.
CopyrightJudith R. Stabel/USDA-ARS-National Animal Disease Center
Cattle with end-stage clinical Johne's disease, showing signs of severe emaciation.
SymptomsCattle with end-stage clinical Johne's disease, showing signs of severe emaciation.Judith R. Stabel/USDA-ARS-National Animal Disease Center
Cow with clinical Johne's disease, exhibiting severe watery diarrhoea.
TitleSymptoms
CaptionCow with clinical Johne's disease, exhibiting severe watery diarrhoea.
CopyrightJudith R. Stabel/USDA-ARS-National Animal Disease Center
Cow with clinical Johne's disease, exhibiting severe watery diarrhoea.
SymptomsCow with clinical Johne's disease, exhibiting severe watery diarrhoea.Judith R. Stabel/USDA-ARS-National Animal Disease Center
Cow with clinical Johne's disease exhibiting submandibular oedema.
TitleSymptoms
CaptionCow with clinical Johne's disease exhibiting submandibular oedema.
CopyrightJudith R. Stabel/USDA-ARS-National Animal Disease Center
Cow with clinical Johne's disease exhibiting submandibular oedema.
SymptomsCow with clinical Johne's disease exhibiting submandibular oedema.Judith R. Stabel/USDA-ARS-National Animal Disease Center
Gross pathology of section of ileum from cow with clinical disease, showing corrugation and thickening.
TitlePathology
CaptionGross pathology of section of ileum from cow with clinical disease, showing corrugation and thickening.
CopyrightJudith R. Stabel/USDA-ARS-National Animal Disease Center
Gross pathology of section of ileum from cow with clinical disease, showing corrugation and thickening.
PathologyGross pathology of section of ileum from cow with clinical disease, showing corrugation and thickening.Judith R. Stabel/USDA-ARS-National Animal Disease Center

Identity

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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

Overview

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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.

Hosts/Species Affected

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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).

Distribution

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

Distribution Table

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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.

Continent/Country/RegionDistributionLast ReportedOriginFirst ReportedInvasiveReferenceNotes

Asia

AfghanistanNo information availableOIE, 2009
ArmeniaDisease not reportedOIE, 2009
AzerbaijanDisease not reportedOIE, 2009
BahrainDisease never reportedOIE, 2009
BangladeshPresentOIE, 2009
BhutanDisease not reported2004OIE, 2009
Brunei DarussalamDisease not reportedOIE Handistatus, 2005
CambodiaNo information availableOIE, 2009
ChinaNo information availableNULLXie et al., 1982; OIE, 2009
-Hong KongNo information availableOIE, 2009
Georgia (Republic of)Disease never reportedOIE Handistatus, 2005
IndiaRestricted distributionNULLDatta, 1934; Juneja et al., 1991; OIE, 2009
-AssamPresentPande, 1940
-HaryanaPresentBali and Singh, 1980
-Indian PunjabPresentSharma et al., 1985
-KeralaPresentPaily et al., 1979
-MaharashtraPresentSrivastava and More, 1987
IndonesiaDisease not reportedOIE, 2009
IranPresentNULLTalatchian, 1965; Muhammed and Ivoghli, 1983; OIE, 2009
IraqDisease never reportedOIE, 2009
IsraelPresentNULLOIE, 2009
JapanPresentNULLNemoto, 1961; OIE, 2009
JordanNo information availableNULLOIE, 2009
KazakhstanDisease not reportedOIE, 2009
Korea, DPRDisease not reportedOIE Handistatus, 2005
Korea, Republic ofPresentNULLOIE, 2009
KuwaitDisease not reportedOIE, 2009
KyrgyzstanDisease not reportedOIE, 2009
LaosDisease not reportedOIE, 2009
LebanonDisease not reported1999OIE, 2009
MalaysiaPresentNULLOIE, 2009
-Peninsular MalaysiaSerological evidence and/or isolation of the agentOIE Handistatus, 2005
-SabahReported present or known to be presentOIE Handistatus, 2005
-SarawakSerological evidence and/or isolation of the agentOIE Handistatus, 2005
MongoliaAbsent, reported but not confirmedNULLOIE, 2009
MyanmarNo information availableNULLOIE, 2009
NepalAbsent, reported but not confirmedNULLJoshi et al., 1974; OIE, 2009
OmanNo information availableNULLOIE, 2009
PakistanDisease not reported1992Khan and Huq, 1962; OIE, 2009
PhilippinesNo information availableNULLNovilla, 1972; OIE, 2009
QatarPresentOIE, 2009
Saudi ArabiaAbsent, reported but not confirmedNULLOIE, 2009
SingaporeDisease never reportedOIE, 2009
Sri LankaDisease never reportedOIE, 2009
SyriaDisease not reportedNULLBaradi et al., 1972; Lu et al., 1987; OIE, 2009
TaiwanReported present or known to be presentOIE Handistatus, 2005
TajikistanDisease never reportedOIE, 2009
ThailandPresentNULLOIE, 2009
TurkeyNo information availableNULLAlibasoglu et al., 1973; Vural and Atala, 1988; OIE, 2009
TurkmenistanDisease not reportedOIE Handistatus, 2005
United Arab EmiratesNo information availableNULLOIE, 2009
UzbekistanDisease not reportedOIE Handistatus, 2005
VietnamAbsent, reported but not confirmedOIE, 2009
YemenNo information availableOIE, 2009

Africa

AlgeriaDisease not reportedOIE, 2009
AngolaNo information availableOIE, 2009
BeninNo information availableOIE, 2009
BotswanaDisease not reportedOIE, 2009
Burkina FasoNo information availableOIE, 2009
BurundiNo information availableOIE Handistatus, 2005
CameroonNo information availableOIE Handistatus, 2005
Cape VerdeNo information availableMichel and Bastianello, 2000; OIE Handistatus, 2005
Central African RepublicDisease not reportedOIE Handistatus, 2005
ChadNo information availableOIE, 2009
CongoNo information availableNULLDeom and Moretelmans, 1955; OIE, 2009
Congo Democratic RepublicNo information availableOIE Handistatus, 2005
Côte d'IvoireDisease not reportedOIE Handistatus, 2005
DjiboutiDisease not reportedOIE, 2009
EgyptDisease never reportedOIE, 2009
EritreaPresentNULLOIE, 2009
EthiopiaNo information availableNULLOIE, 2009
GabonNo information availableOIE, 2009
GambiaNo information availableOIE, 2009
GhanaNo information availableOIE, 2009
GuineaNo information availableOIE, 2009
Guinea-BissauNo information availableOIE, 2009
KenyaAbsent, reported but not confirmedNULLPaling et al., 1988; OIE, 2009
LesothoDisease not reportedOIE, 2009
LibyaDisease not reportedOIE Handistatus, 2005
MadagascarDisease never reportedOIE, 2009
MalawiNo information availableOIE, 2009
MaliNo information availableOIE, 2009
MauritiusDisease not reportedOIE, 2009
MoroccoNo information availableNULLBenazzi et al., 1996; OIE, 2009
MozambiqueDisease not reportedOIE, 2009
NamibiaDisease not reportedOIE, 2009
NigeriaNo information availableNULLJohnson et al., 1962; OIE, 2009
RéunionNo information availableOIE Handistatus, 2005
RwandaDisease never reportedOIE, 2009
Sao Tome and PrincipeDisease not reportedOIE Handistatus, 2005
SenegalNo information availableNULLKonte, 1988; OIE, 2009
SeychellesDisease not reportedOIE Handistatus, 2005
SomaliaNo information availableOIE Handistatus, 2005
South AfricaPresentNULLOIE, 2009
SudanDisease not reported200405Fawi and Obeid, 1964; OIE, 2009
SwazilandNo information availableOIE, 2009
TanzaniaDisease not reportedOIE, 2009
TogoNo information availableOIE, 2009
TunisiaDisease not reported2002Gallo et al., 1989; OIE, 2009
UgandaAbsent, reported but not confirmedOIE, 2009
ZambiaNo information availableNULLPandey et al., 1987; OIE, 2009
ZimbabweDisease not reported1996Pandey et al., 1987; OIE, 2009

North America

BermudaDisease not reportedOIE Handistatus, 2005
CanadaPresentNULLOIE, 2009
-OntarioPresentMcNab et al., 1991
-Prince Edward IslandPresentvan Leeuwen et al., 2001
GreenlandDisease never reportedOIE, 2009
MexicoPresentNULLRamirez et al., 1979; Salman et al., 1990; OIE, 2009
USAPresentOIE, 2009
-ArizonaPresentKopecky and, 1973
-ArkansasPresentMerkal et al., 1987
-CaliforniaPresentMerkal et al., 1987
-ColoradoPresentMerkal et al., 1987
-ConnecticutPresentKopecky and, 1973
-FloridaPresentMerkal et al., 1987
-GeorgiaPresentMerkal et al., 1987
-IllinoisPresentMerkal et al., 1987
-IndianaPresentKopecky and, 1973
-IowaPresentKopecky and, 1973
-KansasPresentMerkal et al., 1987
-KentuckyPresentMerkal et al., 1987
-LouisianaPresentKopecky and, 1973
-MainePresentMerkal et al., 1987
-MarylandPresentKopecky and, 1973
-MassachusettsPresentKopecky and, 1973
-MichiganPresentMerkal et al., 1987
-MinnesotaPresentMerkal et al., 1987
-MississippiPresentKopecky and, 1973
-MissouriPresentMerkal et al., 1987
-NebraskaPresentMerkal et al., 1987
-NevadaPresentKopecky and, 1973
-New JerseyPresentKopecky and, 1973
-New MexicoPresentKopecky and, 1973
-New YorkPresentMerkal et al., 1987
-North CarolinaPresentKopecky and, 1973
-North DakotaPresentKopecky and, 1973
-OhioPresentMerkal et al., 1987
-OklahomaPresentMerkal et al., 1987
-OregonPresentMerkal et al., 1987
-PennsylvaniaPresentMerkal et al., 1987
-South CarolinaPresentKopecky and, 1973
-South DakotaPresentKopecky and, 1973
-TennesseePresentMerkal et al., 1987
-TexasPresentMerkal et al., 1987
-UtahPresentKopecky and, 1973
-VirginiaPresentMerkal et al., 1987
-WashingtonPresentMerkal et al., 1987
-West VirginiaPresentKopecky and, 1973
-WisconsinPresentMerkal et al., 1987

Central America and Caribbean

BarbadosSerological evidence and/or isolation of the agentOIE Handistatus, 2005
BelizeDisease not reportedOIE, 2009
British Virgin IslandsDisease never reportedOIE Handistatus, 2005
Cayman IslandsDisease not reportedOIE Handistatus, 2005
Costa RicaPresentNULLDolz et al., 1999; OIE, 2009
CubaPresentNULLOIE, 2009
CuraçaoDisease not reportedOIE Handistatus, 2005
DominicaDisease not reportedOIE Handistatus, 2005
Dominican RepublicDisease never reportedOIE, 2009
El SalvadorNo information availableOIE, 2009
GuadeloupeNo information availableOIE, 2009
GuatemalaDisease never reportedOIE, 2009
HaitiDisease never reportedOIE, 2009
HondurasNo information availableOIE, 2009
JamaicaDisease not reportedOIE, 2009
MartiniqueAbsent, reported but not confirmedOIE, 2009
NicaraguaNo information availableOIE, 2009
PanamaNo information availableOIE, 2009
Saint Kitts and NevisDisease never reportedOIE Handistatus, 2005
Saint Vincent and the GrenadinesDisease never reportedOIE Handistatus, 2005
Trinidad and TobagoReported present or known to be presentOIE Handistatus, 2005

South America

ArgentinaPresentNULLUbach, 1941; OIE, 2009
BoliviaAbsent, reported but not confirmedOIE, 2009
BrazilDisease not reported2003Santos and da Silva, 1956; OIE, 2009
-Santa CatarinaPresentPortugal et al., 1979
-Sao PauloPresentCastro and Nemoto, 1972
ChilePresentNULLZamora et al., 1975; OIE, 2009
-Easter IslandPresentBoulanger et al., 1968
ColombiaPresentNULLMogollón et al., 1983; OIE, 2009
EcuadorDisease not reported2006OIE, 2009
Falkland IslandsLast reported1993OIE Handistatus, 2005
French GuianaDisease not reported2003OIE, 2009
GuyanaDisease not reportedOIE Handistatus, 2005
ParaguayDisease not reportedOIE Handistatus, 2005
PeruRestricted distributionNULLRamos Saco, 1953; OIE, 2009
UruguayPresentNULLErrico and Bermúdez, 1983; OIE, 2009
VenezuelaNo information availableNULLIlukevich et al., 1979; OIE, 2009

Europe

AlbaniaNo information availableNULLVavako, 1942; OIE, 2009
AndorraCAB Abstracts data miningOIE Handistatus, 2005
AustriaPresentOIE, 2009
BelarusDisease never reportedOIE, 2009
BelgiumDisease not reported2004Boelaert et al., 2000; OIE, 2009
Bosnia-HercegovinaDisease not reportedBadnjevic et al., 1962; OIE Handistatus, 2005
BulgariaDisease not reported1996Kujumgiev, 1964; OIE, 2009
CroatiaPresentNULLOIE, 2009
CyprusPresentNULLCrowther et al., 1976; OIE, 2009
Czech RepublicPresentNULLPavlik et al., 2000; OIE, 2009
DenmarkPresentNULLNielsen et al., 2000; OIE, 2009
EstoniaPresentOIE, 2009
FinlandDisease not reportedNULLOIE, 2009
FranceNo information availableNULLGasse, 1961; Rihal, 1979; OIE, 2009
GermanyPresentNULLIturria, 1989; OIE, 2009
GreecePresentNULLOIE, 2009
HungaryRestricted distributionNULLOIE, 2009
IcelandPresentNULLGunnarsson, 1979; OIE, 2009
IrelandNo information availableNULLO'Brien et al., 1972; Grant et al., 2001; OIE, 2009
Isle of Man (UK)Reported present or known to be presentOIE Handistatus, 2005
ItalyNo information availableNULLNebbia et al., 2000; OIE, 2009
JerseyCAB Abstracts data miningOIE Handistatus, 2005
LatviaDisease not reported199910OIE, 2009
LiechtensteinDisease not reportedOIE, 2009
LithuaniaDisease not reportedNULLOIE, 2009
LuxembourgPresentNULLOIE, 2009
MacedoniaAbsent, reported but not confirmedOIE, 2009
MaltaDisease not reportedOIE, 2009
MoldovaDisease not reportedOIE Handistatus, 2005
MontenegroPresentOIE, 2009
NetherlandsPresentNULLKalis et al., 2000; OIE, 2009
NorwayPresentNULLSaxegaard and Fodstad, 1985; OIE, 2009
PolandPresentNULLRamisz, 1970; OIE, 2009
PortugalPresentNULLOIE, 2009
RomaniaPresentOIE, 2009
Russian FederationRestricted distributionOIE, 2009
-Russia (Europe)Present
SerbiaDisease never reportedOIE, 2009
SlovakiaPresentNULLHanzlíková and Vilímek, 1989; OIE, 2009
SloveniaDisease not reported200709OIE, 2009
SpainRestricted distributionNULLMainar and Vázquez, 1998; OIE, 2009
SwedenDisease not reported200507Hoflund and Viriden, 1956; OIE, 2009
SwitzerlandPresentNULLOIE, 2009
UKPresentNULLCetinkaya et al., 1998; OIE, 2009
-Northern IrelandReported present or known to be presentOIE Handistatus, 2005
UkraineDisease not reported2004Tsellarius, 1969; OIE, 2009
Yugoslavia (former)No information availableOIE Handistatus, 2005
Yugoslavia (Serbia and Montenegro)Disease never reportedBadnjevic and Forsek, 1968; OIE Handistatus, 2005

Oceania

AustraliaRestricted distributionNULLAllworth and Kennedy, 2000; Kennedy and Allworth, 2000; OIE, 2009
-New South WalesPresentEverett et al., 1989
-QueenslandPresentFraser, 1989
-South AustraliaPresentVandegraaff, 1989
-TasmaniaPresentPitt, 1989
-VictoriaPresentAlbiston and Talbot, 1936
FijiPresent
French PolynesiaDisease not reportedOIE, 2009
New CaledoniaPresentNULLOIE, 2009
New ZealandPresentNULLDavidson, 1970; OIE, 2009
SamoaDisease never reportedOIE Handistatus, 2005
VanuatuDisease never reportedOIE Handistatus, 2005
Wallis and Futuna IslandsNo information availableOIE Handistatus, 2005

Pathology

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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

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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

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SignLife StagesType
Cardiovascular Signs / Tachycardia, rapid pulse, high heart rate Sign
Cardiovascular Signs / Tachycardia, rapid pulse, high heart rate Sign
Cardiovascular Signs / Tachycardia, rapid pulse, high heart rate Sign
Digestive Signs / Anorexia, loss or decreased appetite, not nursing, off feed Cattle & Buffaloes:Cow Diagnosis
Digestive Signs / Dark colour stools, faeces Sign
Digestive Signs / Diarrhoea Cattle & Buffaloes:Cow Diagnosis
Digestive Signs / Parasites passed per rectum, in stools, faeces Cattle & Buffaloes:All Stages Diagnosis
Digestive Signs / Polyphagia, excessive appetite Sign
General Signs / Dehydration Cattle & Buffaloes:Cow Sign
General Signs / Fever, pyrexia, hyperthermia Sign
General Signs / Fever, pyrexia, hyperthermia Sign
General Signs / Generalized weakness, paresis, paralysis Cattle & Buffaloes:All Stages Sign
General Signs / Head, face, ears, jaw, nose, nasal, swelling, mass Sign
General Signs / Head, face, ears, jaw, nose, nasal, swelling, mass Sign
General Signs / Head, face, ears, jaw, nose, nasal, swelling, mass Sign
General Signs / Inability to stand, downer, prostration Sign
General Signs / Lack of growth or weight gain, retarded, stunted growth Sign
General Signs / Pale mucous membranes or skin, anemia Sign
General Signs / Pale mucous membranes or skin, anemia Sign
General Signs / Pale mucous membranes or skin, anemia Sign
General Signs / Polydipsia, excessive fluid consumption, excessive thirst Sign
General Signs / Underweight, poor condition, thin, emaciated, unthriftiness, ill thrift Cattle & Buffaloes:Cow Diagnosis
General Signs / Weight loss Cattle & Buffaloes:Cow Diagnosis
Nervous Signs / Dullness, depression, lethargy, depressed, lethargic, listless Sign
Nervous Signs / Dullness, depression, lethargy, depressed, lethargic, listless Sign
Reproductive Signs / Abortion or weak newborns, stillbirth Sign
Reproductive Signs / Agalactia, decreased, absent milk production Sign
Reproductive Signs / Agalactia, decreased, absent milk production Sign
Reproductive Signs / Anestrus, absence of reproductive cycle, no visible estrus Cattle & Buffaloes:Heifer,Cattle & Buffaloes:Cow Sign
Reproductive Signs / Decreased in size, small ovary, ovaries Cattle & Buffaloes:Heifer,Cattle & Buffaloes:Cow Sign
Reproductive Signs / Female infertility, repeat breeder Sign
Reproductive Signs / Female infertility, repeat breeder Sign
Reproductive Signs / Female infertility, repeat breeder Sign
Reproductive Signs / Male infertility Cattle & Buffaloes:Bull Sign
Skin / Integumentary Signs / Alopecia, thinning, shedding, easily epilated, loss of, hair Cattle & Buffaloes:All Stages Sign
Skin / Integumentary Signs / Decreased hair pigment, general or focal, poliosis Cattle & Buffaloes:All Stages Sign
Skin / Integumentary Signs / Dryness of skin or hair Sign
Skin / Integumentary Signs / Rough hair coat, dull, standing on end Cattle & Buffaloes:Cow Sign
Skin / Integumentary Signs / Skin edema Sign
Skin / Integumentary Signs / Skin edema Sign
Skin / Integumentary Signs / Skin edema Sign

Disease Course

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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

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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

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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

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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

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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

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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.

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Links to Websites

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International Association for paratuberculosishttp://www.paratuberculosis.org
Johnes Disease Information Centerhttp://www.johnesdisease.org/Part of the USA National Johne's Education Initiative. The Johne’s Education Initiative is a response to needs identified in the Johne’s Strategic Plan. The focus is on producers and those who work with them. Basic funding is being provided by USDA-APHIS, but one of the needs is to demonstrate industry support for the Johne’s program. It is anticipated that this will provide opportunities to increase funding for education efforts associated with the project.
Johnes Information centrehttp://www.johnes.orgSet up by The University of Wisconsin- School of veterinary medicine.

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