- Host Animals
- Hosts/Species Affected
- Systems Affected
- Distribution Table
- List of Symptoms/Signs
- Disease Course
- Impact: Economic
- Zoonoses and Food Safety
- Disease Treatment
- Prevention and Control
- Links to Websites
- Distribution Maps
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PicturesTop of page
IdentityTop of page
Preferred Scientific Name
International Common Names
- English: cowdriosis; heartwater, cowdria ruminantium, cowdriosis, in ruminants- exotic; heartwater, ehrlichia ruminantium, in ruminants- exotic
- French: cowdriose
Local Common Names
- Guadeloupe: Mal cadik
- Mali: tiéoudé
- Nigeria: kaboa
- South Africa: dronkgalsiekte; hartwater; nintas
OverviewTop of page
Heartwater (or cowdriosis) is a tickborne disease of sheep, goats, cattle and some wild ruminants caused by the rickettsia, Ehrlichia ruminantium (previously Cowdria ruminantium). It is a small pleomorphic organism (0.2-2.7 µm) and colonies named morula containing varying numbers are found in the cytoplasm of endothelial cells (Cowdry, 1926; Pienaar, 1970).
According to Neitz (1968), the first record of heartwater was probably made in South Africa by the Voortrekker pioneer, Louis Trichard in 1838. In an entry in his diary on the 9th March 1838 he mentions a fatal disease, ‘nintas’, amongst his sheep, approximately 3 weeks after a massive tick infestation. However, it was only in 1876, almost 50 years later, that the first official report describing heartwater as a generally known disease along the coast and borders of King William’s Town, was presented before the Cattle and Sheep Disease Commission in Grahamstown, South Africa. According to Webb (1877), the disease was introduced into the eastern Cape region at about the same time that William Bowker found a bont tick (Amblyomma hebraeum) on a cow which was imported from northern KwaZulu-Natal (then Zululand) in approximately 1837. Due to confusion with other prevalent conditions of unknown aetiologies at that time, it is difficult to follow the introduction and spread of the disease in Africa. It is, for instance, still not clear whether heartwater is a disease indigenous to the African continent or whether it was imported at some stage, possibly from Madagascar. However, current knowledge suggests that it is a disease of the African mainland. Despite the fact that certain Amblyomma species occur on the Asian and American continents, there is no evidence that the disease exists there.
The first major breakthrough in understanding the disease came when Dixon (1898) and Edington (1898) proved that it could be produced experimentally by inoculating blood from diseased to susceptible animals. Although the causative organism could not be demonstrated in the blood or tissue of diseased animals at the time, it was believed that heartwater was caused by a microorganism (Hutcheon, 1900), possibly a virus (Spreull, 1904). At about the same time Lounsbury (1900) confirmed the long-standing suspicion that the bont tick (A. hebraeum) was the vector in South Africa. However, it was not until 1925 that Cowdry (Cowdry, 1925a; Cowdry, 1925b) successfully demonstrated the organism in tissue of infected animals and ticks. Cowdry named the organism Rickettsia ruminantium but this was later changed to Cowdria ruminantium (Moshkovski, 1947) and finally to E. ruminantium (Dumler et al., 2001).
Traditional rickettsial taxonomy assigned Cowdria ruminantium as the sole member of the genus Cowdria in the tribe Ehrlichieae. This was one of three tribes within the family Rickettsiaceae in the order Rickettsiales which initially encompassed all intracellular bacteria but from which the Chlamydiae were later removed (Moulders, 1984). The obligate intracellular nature of E. ruminantium, coupled with morphological features suggestive of a Chlamydia-like life cycle led to confusion as to its position in the ehrlichial hierarchy (Uilenberg, 1983). In 2001, Dumler et al. defined after 16S ribosomal DNA and groESL gene comparisons that all members of the tribes Ehrlichieae and Wolbachieae had to be transferred to the family Anaplasmataceae and that the family Rickettsiaceae had to be eliminated. In Anaplasmataceae, four genera are present Anaplasma, Ehrlichia, Wolbachia and Neorickettsia. The genus Ehrlichia now includes E. ruminantium (formerly Cowdria ruminantium) but no longer Anaplasma bovis, phagocytophila, and HGE (belonging now to genus Anaplasma) and the genus neorickettsia (including Ehrlichia sennetsu and Ehrlichia risticii). In this reorganization, E. ruminantium is closer to E. chaffeensis, ewingii, muris, ovis and canis.
Economic impact and prevalence
Heartwater is one of the main tickborne diseases together with theileriosis and trypanosomosis in tropical countries. For the Southern Africa Development Community (Angola, Botswana, Malawi, Mozambique, South Africa, Swaziland, Tanzania and Zimbabwe) the losses are estimated around 47.6 millions of dollars per year. Important losses are due to mortality, diminution of productivity in farming systems and cost of treatment (use of antibiotics and acaricides). It is a major, and in some instances, the most important obstacle against introducing high producing animals into Africa with the aim of upgrading or replacing local stock (Uilenberg, 1982a). It is a major disease problem when local animals are, usually for the sake of grazing, moved from heartwater-free to heartwater-infected areas (Neitz, 1967). It remains a problem and a threat in endemic areas especially amongst small stock (Thomas and Mansvelt, 1957). The effect of dipping and environmental changes influences endemic stability, which is often difficult or impossible to manipulate (Bezuidenhout and Bigalke, 1987).
The development of molecular diagnostic tools allows a better estimation of the prevalence of heartwater thanks to detection both in organs from suspected dead ruminants and in ticks. In Burkina Faso, the E. ruminantium prevalence in ticks by pCS20 nested PCR has been evaluated from 3 to 10% depending on the year of tick samplings (Dr Hassane Adakal, personal communication, Adakal et al., 2010b). Moreover, a study evaluating the efficiency of the inactivated vaccine in field conditions in Burkina Faso allowed identifying the impact of heartwater on susceptible ruminants. In this study, two successive trial assays on susceptible imported sahelian sheep demonstrated that 51% and 53% of unvaccinated sheep died from heartwater (Adakal et al., 2010a). In the Gambia, the seroprevalence rate per site in small ruminants varied from 6.9% and 100% (5 regions) (Faburay et al., 2005). The percentage of E. ruminantium infected Amblyomma ticks collected on 15 different sites, varied strongly from 1.6 to 15.1% depending on the site of sampling (Faburay et al., 2007a). These results showed a gradient risk of increasing heartwater from the East to the West of the Gambia. In Nigeria a study done in 2011 on 7 sites in the south of Nigeria shows a 9.6% of E. ruminantium tick prevalence (Personal communications, Dr Maxwell Opara). In the Caribbean region, only Guadeloupe and Antigua are infected with heartwater. In Guadeloupe, the E. ruminantium tick prevalence is higher (i.e. 19.1% in Marie Galante with 73.8% of herds infested) compared to Antigua 5.8% of E.ruminantium infected ticks with only 2.2% of herds infested (Vachiéry et al., 2008b). These islands still represent a reservoir for ticks and heartwater in the Caribbean. It is a threat to areas such as the American mainland due to migratory birds potentially carrying infected ticks from the Caribbean area where the disease is present. Moreover, potential vectors are present but do not harbour the disease (Uilenberg, 1982b; Uilenberg et al., 1984). It is also a threat to countries where the vectors may be introduced and become established (Wilson and Richard, 1984; Barré et al., 1987).
It will therefore probably remain a disease of major importance until an effective and safe vaccine becomes available.
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. For further information on this disease from OIE, see the website: www.oie.int
Host AnimalsTop of page
|Animal name||Context||Life stage||System|
|Aepyceros melampus||Wild host|
|Ammotragus lervia (aoudad)||Domesticated host||Sheep & Goats: All Stages|
|Antidorcas marsupialis||Wild host|
|Axis axis (Indian spotted deer)||Experimental settings, Wild host|
|Bos indicus (zebu)||Domesticated host||Cattle & Buffaloes: All Stages|
|Bos taurus (cattle)||Domesticated host||Cattle & Buffaloes: All Stages|
|Boselaphus tragocamelus||Wild host|
|Bubalus bubalis (Asian water buffalo)||Wild host||Cattle & Buffaloes: All Stages|
|Capra hircus (goats)||Domesticated host||Sheep & Goats: All Stages|
|Cervus dama||Experimental settings, Wild host|
|Cervus timorensis||Experimental settings, Wild host|
|Connochaetes gnou||Wild host|
|Connochaetes taurinus||Wild host|
|Damaliscus albifrons||Wild host|
|Diceros bicornis||Wild host|
|Geochelone pardalis||Experimental settings|
|Giraffa camelopardalis||Wild host|
|Hemitragus jemlahicus||Wild host|
|Kobus ellipsiprymnus||Wild host|
|Lepus saxatilis||Experimental settings|
|Loxodonta africana||Experimental settings|
|Mastomys coucha||Wild host|
|Odocoileus virginianus||Experimental settings, Wild host|
|Ovis aries (sheep)||Domesticated host||Sheep & Goats: All Stages|
|Ovis orientalis||Domesticated host||Sheep & Goats: All Stages|
|Rhabdomys pumilio||Experimental settings, Wild host|
|Syncerus caffer||Wild host||Cattle & Buffaloes: All Stages|
|Tragelaphus oryx||Wild host|
|Tragelaphus spekii||Wild host|
|Tragelaphus strepsiceros||Wild host|
Hosts/Species AffectedTop of page
All the domestic representatives of the family Bovidae are susceptible to clinical disease. The susceptibility of the different breeds of domestic ruminants, however, varies, Bos indicus (zebu) breeds being generally more resistant than European breeds (Bonsma, 1981; Uilenberg, 1983). The resistance of the local indigenous zebu breeds in Africa is probably inherited as a result of natural selection. Asian buffalo, Bubalus bubalis, are also susceptible to heartwater. Although sheep are more susceptible to heartwater than cattle, there is also a variation between breeds and the Blackheaded Persian possesses a certain degree of natural resistance (Uilenberg, 1983). The most sensitive species to heartwater is the goat. Wild ruminants including Cervidae, Bovidea and Giraffidae are also infected by E. ruminantium
So far only the bleskbok, black wildebeest, helmeted guinea fowl, leopard tortoise and scrub hare are known to harbour E. ruminantium subclinically for any length of time and constitute a tick and pathogen reservoir. Of all the indigenous African wild ruminant species only the eland, blesbok, springbok and black wildebeest have been reported to develop clinical disease (Oberem and Bezuidenhout, 1987).
A knowledge of the susceptibility of wild ruminants to heartwater is important where farmers wish to re-introduce ruminant game species into heartwater endemic areas. Wild ruminants also play a role as sources of infection for ticks, particularly in those areas where stringent tick control in domestic animals is practised.
Laboratory mice are also susceptible to E. ruminantium, however, the pathogenicity of the different stocks of Ehrlichia to mice varies significantly (MacKenzie and McHardy, 1987). Ball3 and Welgevonden strains are pathogenic for mice. The multimammate mouse (Mastomys coucha) (MacKenzie and McHardy, 1987) and the striped mouse (Rhabdomys pumilio) (Hudson and Henderson, 1941) are also susceptible to infection, but as wild rodents do not act as host for the tick vector they are unlikely to play a role in the epidemiology of the disease.
Systems AffectedTop of page blood and circulatory system diseases of large ruminants
blood and circulatory system diseases of small ruminants
nervous system diseases of large ruminants
nervous system diseases of small ruminants
respiratory diseases of large ruminants
respiratory diseases of small ruminants
DistributionTop of page
Heartwater only occurs where its tick vectors, Amblyomma, are present. Countries where heartwater has been conclusively diagnosed are listed in the table. The improvement of molecular diagnosis allows confirmation of the presence of E. ruminantium in different countries.
According to Camus et al. (1996), after examining various reports and veterinary literature since 1930, heartwater does not occur in Guinea, Sierra Leone, Togo, Saudi Arabia, Yemen and Oman, even though there is at least one efficient vector present in these countries and it occurs in the neighbouring countries. A nervous condition and lesions very reminiscent to those of heartwater have been described in cattle in Cuba (Figueroa and Sutherland, 1968; Figueroa et al., 1970; Figueroa and Sutherland, 1972) and in French Guiana (Sapin, 1981) however until now, no report or confirmation of heartwater clinical case occurs in both countries. Although no African vectors have been found in these countries, a potential vector, Amblyomma cajennense, does occur in these countries (Camus et al., 1996). In the Caribbean regions, only Guadeloupe and Antigua are infected with heartwater whereas Amblyomma variegatum is present in several islands of the lesser Antilles at lower level of infestation.
Therefore all countries where known Amblyomma vectors are parasites of livestock, or where neighbouring countries are infected, are at risk from the disease. These include the countries listed above, most of the Caribbean islands and the American continent. Quite surprisingly, heartwater has never been observed in Asia from where most ruminants originated, and despite the fact that many Amblyomma spp. ticks occur there.
According to AU-IBAR (2011), cowdriosis is present in Africa south of the Sahara and the islands of the Comoros, Zanzibar, Madagascar, Sao Tomé, Réunion, and Mauritius. Many ruminants, including some antelope species, are susceptible. In 2011, fourteen (14) countries reported to AU-IBAR the occurrence of heartwater where a total of 810 outbreaks 9,546 cases and 977 deaths were recorded (See table below). Zimbabwe (364) reported the highest number of outbreaks, followed by Botswana (88), Zambia (74), and Swaziland (68). The corresponding number of cases were highest in Tanzania (6680), followed by Zimbabwe (826), Zambia (470) and Somalia (396).
Countries reporting cowdriosis to AU-IBAR in 2011.
For current information on disease incidence, see OIE's WAHID Interface.
Distribution TableTop of page
The distribution in this summary table is based on all the information available. When several references are cited, they may give conflicting information on the status. Further details may be available for individual references in the Distribution Table Details section which can be selected by going to Generate Report.
|Continent/Country/Region||Distribution||Last Reported||Origin||First Reported||Invasive||Reference||Notes|
|Afghanistan||No information available||OIE, 2009|
|Armenia||Disease not reported||OIE, 2009|
|Azerbaijan||Disease not reported||OIE, 2009|
|Bahrain||Disease not reported||OIE, 2009|
|Bangladesh||Disease never reported||OIE, 2009|
|Bhutan||Disease never reported||OIE, 2009|
|Brunei Darussalam||Disease not reported||OIE Handistatus, 2005|
|Cambodia||No information available||OIE, 2009|
|China||Disease never reported||OIE, 2009|
|-Hong Kong||No information available||OIE, 2009|
|Georgia (Republic of)||Disease never reported||OIE Handistatus, 2005|
|India||Disease never reported||OIE, 2009|
|Indonesia||Disease not reported||OIE, 2009|
|Iran||No information available||OIE, 2009|
|Iraq||No information available||OIE, 2009|
|Israel||Disease never reported||OIE, 2009|
|Japan||No information available||OIE, 2009|
|Jordan||Disease never reported||OIE, 2009|
|Kazakhstan||Disease never reported||OIE, 2009|
|Korea, DPR||Disease not reported||OIE Handistatus, 2005|
|Korea, Republic of||Disease never reported||OIE, 2009|
|Kuwait||Disease not reported||OIE, 2009|
|Kyrgyzstan||Disease not reported||OIE, 2009|
|Laos||No information available||OIE, 2009|
|Lebanon||Disease never reported||OIE, 2009|
|Malaysia||Disease never reported||OIE, 2009|
|-Peninsular Malaysia||Disease not reported||OIE Handistatus, 2005|
|-Sabah||Disease never reported||OIE Handistatus, 2005|
|-Sarawak||Disease never reported||OIE Handistatus, 2005|
|Mongolia||No information available||OIE, 2009|
|Myanmar||No information available||OIE, 2009|
|Nepal||No information available||OIE, 2009|
|Oman||Disease never reported||OIE, 2009|
|Pakistan||Disease not reported||OIE, 2009|
|Philippines||Disease never reported||OIE, 2009|
|Qatar||No information available||OIE, 2009|
|Saudi Arabia||Disease not reported||OIE, 2009|
|Singapore||Disease never reported||OIE, 2009|
|Sri Lanka||Disease never reported||OIE, 2009|
|Syria||Disease not reported||OIE, 2009|
|Taiwan||Disease never reported||OIE Handistatus, 2005|
|Tajikistan||Disease never reported||OIE, 2009|
|Thailand||No information available||OIE, 2009|
|Turkey||No information available||OIE, 2009|
|Turkmenistan||Disease not reported||OIE Handistatus, 2005|
|United Arab Emirates||Disease never reported||OIE, 2009|
|Uzbekistan||Disease never reported||OIE Handistatus, 2005|
|Vietnam||Absent, reported but not confirmed||OIE, 2009|
|Yemen||No information available||OIE, 2009|
|Algeria||Disease not reported||OIE, 2009|
|Angola||Present||NULL||Conceiçao, 1949; OIE, 2009|
|Benin||2007||CCTA, 1963; OIE, 2012|
|Botswana||Restricted distribution||NULL||Roe, 1955; OIE, 2009|
|Burkina Faso||Present||NULL||Adakal et al., 2010a; Camus et al., 1996; OIE, 2009|
|Burundi||Reported present or known to be present||Rwanda-Urundi, 1957; OIE Handistatus, 2005; OIE, 2012|
|Cameroon||Reported present or known to be present||Chad, 1967; OIE Handistatus, 2005|
|Cape Verde||Disease never reported||OIE Handistatus, 2005|
|Central African Republic||No information available||Camus et al., 1996; OIE Handistatus, 2005|
|Chad||No information available||NULL||Malbrant et al., 1939; OIE, 2009|
|Comoros||Present||Plessis et al., 1989; OIE, 2012|
|Congo||Absent, reported but not confirmed||NULL||Rousselot, 1957; OIE, 2009|
|Congo Democratic Republic||No information available||Saceghem R van, 1918; Vaerenbergh, 1960; OIE Handistatus, 2005|
|Côte d'Ivoire||Aillerie, 1932; OIE Handistatus, 2005; OIE, 2012|
|Djibouti||Disease not reported||2005||Office International des Épizooties, 1980; OIE, 2009|
|Egypt||Disease never reported||OIE, 2009|
|Ethiopia||Disease not reported||Tarantino, 1939; Roetti, 1940; OIE, 2012|
|Gabon||No information available||NULL||CCTA, 1962; OIE, 2009|
|Gambia||No information available||NULL||Faburay et al., 2007a; Camus et al., 1996; Faburay et al., 2005; OIE, 2009|
|Ghana||Present||Stewart, 1933; OIE, 2012|
|Guinea||No information available||OIE, 2009|
|Guinea-Bissau||No information available||Tendeiro, 1945; OIE, 2012|
|Kenya||Daubney, 1930; OIE, 2012|
|Lesotho||Disease not reported||OIE, 2009|
|Libya||Disease never reported||OIE Handistatus, 2005|
|Madagascar||Present||Poisson, 1927; Alexander, 1931; OIE, 2012|
|Malawi||No information available||NULL||Meza J de, 1938; OIE, 2009|
|Mali||No information available||NULL||Curasson and Delphy, 1928; OIE, 2009|
|Mauritania||No information available||CCTA, 1962|
|Mauritius||Disease not reported||NULL||Perreau et al., 1980; OIE, 2009|
|Morocco||No information available||OIE, 2009|
|Mozambique||Present||Valadao, 1969; OIE, 2012|
|Namibia||Disease not reported||2011||Shaw, 1990; OIE, 2012|
|Nigeria||Disease not reported||2004||Hall, 1931; Okoh et al., 1987; OIE, 2009|
|Réunion||Perreau et al., 1980; OIE Handistatus, 2005|
|Rwanda||Disease never reported||NULL||Rwanda-Urundi, 1957; OIE, 2009|
|Sao Tome and Principe||CAB Abstracts data mining||Uilenberg, 1982a; OIE Handistatus, 2005|
|Senegal||Disease not reported||200504||Senegal, 1967; OIE, 2009|
|Seychelles||Disease not reported||OIE Handistatus, 2005|
|Somalia||Present||Pellegrini, 1945; Evans, 1963; OIE, 2012|
|South Africa||Present||NULL||Cowdry, 1925b; Neitz, 1968; OIE, 2009|
|Sudan||Restricted distribution||Kennedy, 1931; Karrar, 1959; OIE, 2012|
|Swaziland||Present||Faulkner, 1948; OIE, 2012|
|Tanzania||Present||McCall, 1930; OIE, 2012|
|-Zanzibar||Present||Flach et al., 1990|
|Tunisia||Disease never reported||OIE, 2009|
|Uganda||No information available||NULL||Lewis, 1939; OIE, 2009|
|Zambia||Present||CCTA, 1966; OIE, 2012|
|Zimbabwe||Present||Sinclair, 1927; OIE, 2012|
|Bermuda||Disease not reported||OIE Handistatus, 2005|
|Canada||Disease never reported||OIE, 2009|
|Greenland||Disease never reported||OIE, 2009|
|Mexico||Disease not reported||OIE, 2009|
|USA||Disease never reported||OIE, 2009|
Central America and Caribbean
|Antigua and Barbuda||Disease not reported||Vachiéry et al., 2008b; Birnie et al., 1985|
|Barbados||Disease never reported||OIE Handistatus, 2005|
|Belize||Disease never reported||OIE, 2009|
|British Virgin Islands||Disease never reported||OIE Handistatus, 2005|
|Cayman Islands||Disease never reported||OIE Handistatus, 2005|
|Costa Rica||Disease never reported||OIE, 2009|
|Cuba||Disease never reported||OIE, 2009|
|Curaçao||Disease not reported||OIE Handistatus, 2005|
|Dominica||Disease not reported||OIE Handistatus, 2005|
|Dominican Republic||Disease never reported||OIE, 2009|
|El Salvador||Disease never reported||OIE, 2009|
|Guadeloupe||Present||NULL||Perreau et al., 1980; Molia et al., 2008; OIE, 2009|
|Guatemala||Disease never reported||OIE, 2009|
|Haiti||Disease never reported||OIE, 2009|
|Honduras||Disease never reported||OIE, 2009|
|Jamaica||Disease never reported||OIE, 2009|
|Martinique||Disease not reported||OIE, 2009|
|Nicaragua||Disease never reported||OIE, 2009|
|Panama||Disease never reported||OIE, 2009|
|Saint Kitts and Nevis||CAB Abstracts data mining||OIE Handistatus, 2005|
|Saint Vincent and the Grenadines||Disease never reported||OIE Handistatus, 2005|
|Trinidad and Tobago||Disease never reported||OIE Handistatus, 2005|
|Argentina||Disease not reported||OIE, 2009|
|Bolivia||Disease never reported||OIE, 2009|
|Brazil||Disease never reported||OIE, 2009|
|Chile||Disease never reported||OIE, 2009|
|Colombia||Disease never reported||OIE, 2009|
|Ecuador||Disease never reported||OIE, 2009|
|Falkland Islands||Disease never reported||OIE Handistatus, 2005|
|French Guiana||Disease never reported||OIE, 2009|
|Guyana||Disease never reported||OIE Handistatus, 2005|
|Paraguay||Disease never reported||OIE Handistatus, 2005|
|Peru||Disease never reported||OIE, 2009|
|Uruguay||Disease never reported||OIE, 2009|
|Venezuela||Disease never reported||OIE, 2009|
|Albania||Disease never reported||OIE, 2009|
|Andorra||Disease never reported||OIE Handistatus, 2005|
|Austria||No information available||OIE, 2009|
|Belarus||Disease never reported||OIE, 2009|
|Belgium||Disease not reported||OIE, 2009|
|Bosnia-Hercegovina||Disease not reported||OIE Handistatus, 2005|
|Bulgaria||Disease never reported||OIE, 2009|
|Croatia||Disease never reported||OIE, 2009|
|Cyprus||Disease never reported||OIE, 2009|
|Czech Republic||Disease never reported||OIE, 2009|
|Denmark||Disease never reported||OIE, 2009|
|Estonia||Disease never reported||OIE, 2009|
|Finland||Disease never reported||OIE, 2009|
|France||No information available||OIE, 2009|
|Germany||Disease never reported||OIE, 2009|
|Greece||Disease never reported||OIE, 2009|
|Hungary||Disease never reported||OIE, 2009|
|Iceland||Disease never reported||OIE, 2009|
|Ireland||Disease never reported||OIE, 2009|
|Isle of Man (UK)||Disease never reported||OIE Handistatus, 2005|
|Italy||Disease never reported||OIE, 2009|
|Jersey||Disease never reported||OIE Handistatus, 2005|
|Latvia||Disease never reported||OIE, 2009|
|Liechtenstein||Disease not reported||OIE, 2009|
|Lithuania||Disease never reported||OIE, 2009|
|Luxembourg||Disease never reported||OIE, 2009|
|Macedonia||Disease never reported||OIE, 2009|
|Malta||Disease never reported||OIE, 2009|
|Moldova||Disease never reported||OIE Handistatus, 2005|
|Montenegro||Disease never reported||OIE, 2009|
|Netherlands||Disease never reported||OIE, 2009|
|Norway||Disease never reported||OIE, 2009|
|Poland||Disease never reported||OIE, 2009|
|Portugal||Disease not reported||OIE, 2009|
|Romania||Disease never reported||OIE, 2009|
|Russian Federation||Disease never reported||OIE, 2009|
|Serbia||Disease never reported||OIE, 2009|
|Slovakia||Disease not reported||OIE, 2009|
|Slovenia||Disease never reported||OIE, 2009|
|Spain||Disease never reported||OIE, 2009|
|Sweden||Disease never reported||OIE, 2009|
|Switzerland||Disease never reported||OIE, 2009|
|UK||Disease never reported||OIE, 2009|
|-Northern Ireland||Disease never reported||OIE Handistatus, 2005|
|Ukraine||Disease never reported||OIE, 2009|
|Yugoslavia (former)||No information available||OIE Handistatus, 2005|
|Yugoslavia (Serbia and Montenegro)||Disease never reported||OIE Handistatus, 2005|
|Australia||Disease never reported||OIE, 2009|
|French Polynesia||Disease never reported||OIE, 2009|
|New Caledonia||Disease never reported||OIE, 2009|
|New Zealand||Disease never reported||OIE, 2009|
|Samoa||Disease never reported||OIE Handistatus, 2005|
|Vanuatu||Disease never reported||OIE Handistatus, 2005|
|Wallis and Futuna Islands||No information available||OIE Handistatus, 2005|
PathologyTop of page
Lesions in cattle, sheep and goats are similar, although quite variable in extent and some changes are commoner in certain species than in others. Effusion of body cavities, (hydropericardium, hydrothorax, and, in some cases a degree of ascitis) is a very common change in most fatal cases of heartwater. The transudate is usually transparent or slightly turbid, light yellow fluid that often coagulates on exposure to air. The volume of fluid ranges from 20 ml in goats, about 0.5 litre in sheep to several litres in cattle (Steck, 1928). A hydropericardium, as indicated by the name ‘heartwater’ is a striking change in most animals that die of the disease and is usually more pronounced in sheep and goats than in cattle (Henning, 1956).
Oedema of the lungs is a regular finding and appears to be more severe in most animals that die peracutely from the disease (Pypekamp and Prozesky, 1987). The interlobular septa of the lungs, mediastinum and associated lymph nodes are oedematous and serous frothy fluid oozes from the cut surface of the lung. The trachea and bronchi often contain serofibrinous exudates, and their mucosae are congested, with petechiae and ecchymoses.
Splenomegaly is present although less strikingly in sheep and goats. The cut surface is dark red in colour and has a pulpy consistency. In animals that die peracutely it is often impossible to make a diagnosis on macroscopical lesions alone; splenomegaly, epi- and endocardial haemorrhages are sometimes the only significant changes (Alexander, 1931). Hepatic lesions are less striking with only a mild hepatomegaly present and the gallbladder slightly distended.
Congestion and/or oedema of the mucosa of the abomasum are regularly seen in cattle, but are less common in sheep and goats. Enterorrhagia (small and large intestine) is present in a small percentage of domestic ruminants, particularly Jersey cattle.
The lymph nodes are moderately swollen in most animals. The cut surface is moist and petechiae are often present, especially in the retropharyngeal, submaxillary, cervical, bronchial and mediastinal lymph nodes (Alexander, 1931). Petechiae are frequently visible on mucous membranes of tissues including those of the urinary bladder, vagina, epi- and endocardium and conjunctiva.
The nervous symptoms observed in affected animals are usually attributed to oedema of the brain, although it is often difficult and sometimes impossible to detect swelling of the brain macroscopically. Occasionally, the entire brain, but particularly the gyri of the cerebellum may be strikingly swollen and severe oedema of the brain may even result in a partial prolapse (herniation) of the cerebellum through the foramen magnum. Most animals that die of heartwater show congestion and oedema of the meninges. There is an accumulation of excessive fluid in the subarachnoid space and thickening of the choroids plexus, which has a dull greyish appearance. In some animals petechiae and ecchymoses and sometimes sugillations are evident in the midbrain, brain stem and cerebellum (Pienaar et al., 1966).
Lungs: An alveolar and interstitial oedema occurs in most animals but is not always discernible histopathologically.
Kidneys: Nephrosis of varying degree, is a common change in domestic ruminants that die of heartwater. The observations of (Steck, 1928) of a multifocal lymphocytic interstitial nephritis occurring in cattle, sheep and goats could not be confirmed in subsequent studies (Uilenberg, 1983).
Variable numbers of E. ruminantium colonies are discernable in the cytoplasm of endothelial cells, particularly those of the brain and lungs. Cowdry (1926) and Steck (1928) frequently also observed colonies in the endothelial cells of glomerular capillaries. As a general rule, however, these colonies are difficult to find in haematoxylin- and eosin-stained sections.
Brain: Lesions in the brain of cattle, sheep and goats were described by (Pienaar et al., 1966) and are characterized by changes compatible with oedema, such as widened perivascular spaces which sometimes contain oedematous fluid or protein droplets; swollen, often necrotic, astrocytes; swollen axons, and multifocal microcavitations and haemorrhages affecting mainly the midbrain, brain stem, cerebral white matter and cerebral peduncles. A perivascular accumulation of cells, mainly macrophages and a few neutrophils, and occasionally a vasculitis were observed in all the bovines and in only about 50% of the sheep. A diffuse meningitis, mainly macrophages, was present in a few bovines only. In the majority of animals a fibrinuous chorioditis occurred and occasionally mutifocal glial nodules, mainly confined to the neutrophil around small blood vessels, were apparent in sheep and cattle. Brain lesions in recumbent animals often comprise different degrees of status spongiosus and in severe cases the white matter of the entire brain may be affected.
Other organs: in most animals that die of heartwater the hepatic changes are inconspicuous; the lymph nodes are congested and oedematous; and congestion is the only splenic change.
Several species of game are susceptible to heartwater, but reports on the pathological changes in game that died of heartwater are limited and in most cases lesions are very similar to that described in domestic animals (Young and Basson, 1973; Prozesky, 1987).
Transmission electron microscopical studies of the lung lesions in sheep and goats reveal the presence of minor cytopathic changes in endothelial cells. Apart from mild swelling of mitochondria and endoplasmic reticulum, no other changes occur in most parasitized alveolar endothelial cells. Non-parasitized endothelial cells are sometimes swollen, or even necrotic, and are separated from their basement membranes. Oedema of blood vessel walls is infrequently seen (Prozesky and Plessis, 1985a; Prozesky and Plessis, 1985b).
In all suspected cases a diagnosis of heartwater must be confirmed by the demonstration of Ehrlichia organisms in Giemsa-stained preparations made from the hippocampus.
DiagnosisTop of page
Infected domestic ruminants exhibit a wide range of clinical signs varying from a peracute to mild (clinically inapparent) form. The incubation period in naturally infected cattle ranges 9-29 days (average 18 days) and that of sheep and goats 7-35 days (average 14 days). Peracutely affected animals die within a few hours after the initial fever, either with or without any clinical signs (Alexander, 1931; Neitz, 1968; Uilenberg, 1983).
Acute heartwater is the most common form of the disease in endemic areas. Fever of 40°C or higher, which usually persists for 3-6 days is followed by a drop to subnormal levels shortly before death. Animals gradually show inappetance and eventually stop feeding. Cessation of rumination and difficult breathing follow. Petechiae are visible on the mucous membranes of the conjunctiva (mainly cattle). During the latter stage of acute heartwater, the majority of animals manifest nervous symptoms ranging from a mild incoordination to pronounced convulsions (Alexander, 1931). They are hypersensitive when handled or startled. The gait of affected animals becomes progressively more unsteady, whereas some animals show hypermetria, especially of the forelegs (mainly cattle). They eventually become prostrate, assume a position of lateral recumbency and show intermittent leg-paddling, chewing movements, opisthotonus, licking of the lips and nystagmus. A large amount of froth is usually present at the mouth and nostrils. Diarrhoea is occasionally seen in cattle.
Less severe cases (subacute and mild) occur with clinical signs ranging from slightly less intense than the acute form to little or no signs at all.
Numerous conditions causing nervous symptoms or acute death must be differentiated from heartwater. Diseases such as rabies, cerebral babesiosis/theileriosis, bacterial meningitis/encephalitis, numerous plant, pesticide and heavy metal poisonings (Bezuidenhout et al., 1994).
The confirmation of a diagnosis based on clinical signs and postmortem lesions requires the demonstration of the organisms in the cytoplasm of endothelial cells of blood vessels. The easiest, most efficient and quickest way of doing this is to visualize them in stained smears of the brain (Purchase, 1945) although they may also be found in histological sections such as the brain and kidneys. The examination of brain biopsies in live animals for the confirmation of a diagnosis of heartwater is useful in experimental animals, but is not practical under field conditions (Synge, 1978; Camus and Barré, 1982; Amstel, 1987).
Smears should be air-dried before staining, and stains such as Giemsa or the CAM’s Quick stain give the best results.
The indirect immunofluorescence test (IFA test) is not used anymore (Plessis and Malan, 1987a, b). Many attempts at developing a diagnostic serological test for heartwater have failed due to the high degree of cross-reaction occurring between antigens from different stocks of Ehrlichia and antibodies against Cytoecetes phagocytophila, and some Ehrlichia spp. (Ehrlichia equi, E. canis, E. ovina and E. bovis) (Logan et al., 1986; Camus, 1987; Holland et al., 1987; Plessis and Malan, 1987b). To minimize the degree of cross-reaction, two ELISA were developed using recombinant MAP-1. The first one is an indirect ELISA, ELISA MAP1-B using an immunogenic fraction of MAP-1, the recombinant antigen MAP1-B (Van Vliet et al, 1985). The second one is a competitive ELISA using MAP-1 gene cloned in baculovirus and monoclonal antibodies raised against MAP1 (Katz et al., 1997). Both tests improved the specificity but there is still some cross reactivity with E. canis and E. chaffeensis.
Low sero-positivity of cattle (even cattle that had previously also been vaccinated) occurs in a heartwater endemic areas. The antibody detection is possible 2 weeks after natural infection and lasts few months in naturally infected domestic ruminants moreover the period is shorter for bovine than for small ruminants. Serology as a diagnostic tool for detecting individual animals exposed specifically to E. ruminantium is unreliable. Serological analysis should be considered at herd level taking into account the epidemiological environment and should be complemented by molecular diagnosis. )
There have been significant improvements in the development of molecular tools for the diagnosis of heartwater and the genetic typing of the different strains of E. ruminantium. Concerning the molecular diagnosis, two primers, AB128 and AB129 have been designed to target a fragment of a unique and specific gene of E. ruminantium, pCS20. These primers amplify a 280 pb pCS20 fragment which is revealed by a labelled pCS20 probe (Peter et al., 1995). The PCR/hybridization allows increasing the sensitivity of the method with an experimental detection threshold of 1 to 10 organisms per sample. However the sensitivity of the PCR assay is lower and drops to 61% and 28% with tick samples containing 103 and 102 organisms, respectively (Peter et al., 2000).
A hemi-nested PCR targeting pCS20 fragment of gene was developed using as external primers AB128 and a new primer AB130 followed by a second run on first PCR product using AB128 and AB129 (Martinez et al., 2004). The detection limit (6 organisms per sample) is similar to the PCR/hybridization method described above but the nested PCR method is easier and less time consuming. The diagnosis of heartwater based on examination of brain smears from dead ruminants is much less sensitive than by molecular diagnosis. As an example, the comparison of methods, brain smear observations and pCS20 nested PCR on the same brain samples, demonstrated the improvement of detection threshold with a percentage of heartwater positive cases after brain smears observations of 75% compare to 97% by pCS20 nested PCR (Adakal et al., 2010a). The main disadvantage with nested PCR is the higher contamination risk. The range of strain detection was increased by the use of primers including universal nucleotides AB128’ AB130’ and AB129’ and this method is used routinely for E. ruminantium detection in field samples, especially in ticks (Molia et al., 2008, Adakal et al., 2009, 2010b). The pCS20 hemi-nested PCR allowed detection in organs (lung and brain) from infected dead animals, blood from infected animals during hyperthermia and ticks fresh, frozen or preserved in 70% ethanol. The detection of E. ruminantium by nested PCR is possible in the blood of animals 1 or 2 days before hyperthermia and during the hyperthermia period but not on asymptomatic animals. PCR based methods appear to be more reliable in detecting infection in ticks and this could have epidemiological value in determining the E. ruminantium prevalence in ticks and the geographical distribution of E. ruminantium.
Several quantitative real time PCRs have been developed for the detection of E. ruminantium targeting map-1, map1-1 and pCS20 (Peixoto et al., 2005, Postigo et al., 2002, Steyn et al., 2008). These methods allow the quantification of the pathogen with a similar sensitivity to nested PCR. pCS20 real time PCR can be used for diagnosis due to its ability to detect up to 18 different E. ruminantium strains.
The genetic characterization and structure of E. ruminantium population at regional scale is essential in order to select potential vaccinal strains. The genetic typing of strains was previously done using RFLP on the polymorphic gene map-1 after PCR amplification (Faburay et al., 2007b; Adakal et al., 2010a). Based on the genome analysis of 2 different strains Gardel and Welgevonden, truncated and unique coding sequences specific of strains have been identified. This analysis allows the development of a differential strain-specific diagnosis using nested PCRs targeting 6 unique and 4 truncated CDS (Vachiéry et al., 2008a). New multi-locus methods adapted to E. ruminantium recently have been validated such as multi-locus sequence typing (Adakal et al., 2009) and multi-locus variable number of tandem repeated sequence analysis (Pilet et al., 2012). These tools are recently used on field samples for molecular epidemiological studies.
Protective immunity to E. ruminantium seems to be predominantly cell mediated. Transfer of immune T cells to naïve mice protect them against heartwater and knockout mice studies demonstrate the importance of memory T cells in protection (Plessis et al., 1991, Byrom et al., 2000). In vitro assays on peripheral blood mononuclear cells from vaccinated animals showed the induction of IFNg by both CD8+ and CD4+ T cells in response to total E. ruminantium antigens (Esteves et al., 2004). PBMC from immune animals vaccinated with live vaccine generated CD4+T cell lines after MAP-1 antigen stimulation which expressed IFNg, IFNa, TNFa (Mwangi et al., 2002). Similarly, the protective immune response induced in sheep by four E. ruminantium genes corresponded with increased IFNg expression (Pretorius et al., 2008). Moreover, IFNg has been shown to inhibit the growth of E. ruminantium (Totté et al., 1996).
List of Symptoms/SignsTop of page
|Cardiovascular Signs / Muffled, decreased, heart sounds||Sign|
|Cardiovascular Signs / Tachycardia, rapid pulse, high heart rate||Sign|
|Cardiovascular Signs / Weak pulse, small pulse||Sign|
|Digestive Signs / Abdominal distention||Sign|
|Digestive Signs / Anorexia, loss or decreased appetite, not nursing, off feed||Cattle & Buffaloes:All Stages,Sheep & Goats:All Stages||Sign|
|Digestive Signs / Ascites, fluid abdomen||Sign|
|Digestive Signs / Bloody stools, faeces, haematochezia||Sign|
|Digestive Signs / Diarrhoea||Cattle & Buffaloes:All Stages||Sign|
|Digestive Signs / Excessive salivation, frothing at the mouth, ptyalism||Sign|
|Digestive Signs / Grinding teeth, bruxism, odontoprisis||Sign|
|Digestive Signs / Melena or occult blood in faeces, stools||Cattle & Buffaloes:All Stages||Sign|
|Digestive Signs / Mucous, mucoid stools, faeces||Cattle & Buffaloes:All Stages||Sign|
|Digestive Signs / Rumen hypomotility or atony, decreased rate, motility, strength||Sign|
|Digestive Signs / Tongue protrusion||Sign|
|General Signs / Ataxia, incoordination, staggering, falling||Cattle & Buffaloes:All Stages||Diagnosis|
|General Signs / Dysmetria, hypermetria, hypometria||Sign|
|General Signs / Fever, pyrexia, hyperthermia||Cattle & Buffaloes:All Stages,Sheep & Goats:All Stages||Diagnosis|
|General Signs / Generalized weakness, paresis, paralysis||Sign|
|General Signs / Head, face, ears, jaw weakness, droop, paresis, paralysis||Sheep & Goats:All Stages||Sign|
|General Signs / Hypothermia, low temperature||Cattle & Buffaloes:All Stages||Diagnosis|
|General Signs / Inability to stand, downer, prostration||Sign|
|General Signs / Opisthotonus||Cattle & Buffaloes:All Stages,Sheep & Goats:All Stages||Diagnosis|
|General Signs / Reluctant to move, refusal to move||Sign|
|General Signs / Sudden death, found dead||Cattle & Buffaloes:All Stages,Sheep & Goats:All Stages||Sign|
|General Signs / Tenesmus, straining, dyschezia||Sheep & Goats:All Stages||Sign|
|General Signs / Torticollis, twisted neck||Sign|
|General Signs / Trembling, shivering, fasciculations, chilling||Sign|
|General Signs / Underweight, poor condition, thin, emaciated, unthriftiness, ill thrift||Sign|
|General Signs / Weight loss||Sign|
|Musculoskeletal Signs / Spasms of the limbs, legs, foot, feet in birds||Cattle & Buffaloes:All Stages,Sheep & Goats:All Stages||Diagnosis|
|Nervous Signs / Abnormal anal, perineal, tail reflexes, increased or decreased||Sheep & Goats:All Stages||Sign|
|Nervous Signs / Abnormal behavior, aggression, changing habits||Cattle & Buffaloes:Calf||Sign|
|Nervous Signs / Abnormal forelimb reflexes, increased or decreased||Cattle & Buffaloes:All Stages,Sheep & Goats:All Stages||Diagnosis|
|Nervous Signs / Circling||Sign|
|Nervous Signs / Coma, stupor||Sign|
|Nervous Signs / Constant or increased vocalization||Sheep & Goats:All Stages||Sign|
|Nervous Signs / Dullness, depression, lethargy, depressed, lethargic, listless||Sheep & Goats:Lamb||Sign|
|Nervous Signs / Excitement, delirium, mania||Cattle & Buffaloes:All Stages||Diagnosis|
|Nervous Signs / Head pressing||Cattle & Buffaloes:All Stages||Sign|
|Nervous Signs / Head tilt||Cattle & Buffaloes:All Stages,Sheep & Goats:All Stages||Sign|
|Nervous Signs / Hyperesthesia, irritable, hyperactive||Cattle & Buffaloes:All Stages,Sheep & Goats:All Stages||Diagnosis|
|Nervous Signs / Propulsion, aimless wandering||Cattle & Buffaloes:Calf||Sign|
|Nervous Signs / Seizures or syncope, convulsions, fits, collapse||Cattle & Buffaloes:All Stages,Sheep & Goats:All Stages||Diagnosis|
|Nervous Signs / Tremor||Sign|
|Ophthalmology Signs / Abnormal pupillary response to light||Sign|
|Ophthalmology Signs / Blindness||Sign|
|Ophthalmology Signs / Mydriasis, dilated pupil||Sign|
|Ophthalmology Signs / Nystagmus||Sheep & Goats:All Stages||Sign|
|Reproductive Signs / Abortion or weak newborns, stillbirth||Sign|
|Reproductive Signs / Agalactia, decreased, absent milk production||Sign|
|Respiratory Signs / Abnormal lung or pleural sounds, rales, crackles, wheezes, friction rubs||Sign|
|Respiratory Signs / Coughing, coughs||Cattle & Buffaloes:All Stages,Sheep & Goats:All Stages||Sign|
|Respiratory Signs / Decreased, muffled, lung sounds, absent respiratory sounds||Sign|
|Respiratory Signs / Dull areas on percussion of chest, thorax||Sign|
|Respiratory Signs / Dyspnea, difficult, open mouth breathing, grunt, gasping||Cattle & Buffaloes:All Stages,Sheep & Goats:All Stages||Sign|
|Respiratory Signs / Increased respiratory rate, polypnea, tachypnea, hyperpnea||Cattle & Buffaloes:All Stages,Sheep & Goats:All Stages||Sign|
|Respiratory Signs / Mucoid nasal discharge, serous, watery||Sign|
|Respiratory Signs / Purulent nasal discharge||Sign|
|Skin / Integumentary Signs / Rough hair coat, dull, standing on end||Sign|
|Urinary Signs / Polyuria, increased urine output||Sign|
Disease CourseTop of page
The pathogenesis of the disease is still poorly understood, but the following hypotheses have been proposed.
After infection of the host with E. ruminantium, initial replication of the organisms appears to take place in reticuloendothelial cells and macrophages in the regional lymph nodes. From here the organisms are disseminated via the blood stream to invade endothelial cells of blood vessels of various organs where further multiplication occurs (Plessis, 1970). Endothelial cell parasitization coincides with the onset of fever. There is an increased vascular permeability allowing the seepage of plasma proteins which result in transudation through the serous membranes with resultant tissue oedema (Brown and Skowronek, 1990) and effusion into body cavities. This causes the drastic fall in blood volume before death (Clark, 1962). Oedema of the brain is responsible for the nervous signs, hydropericardium contributes to cardiac dysfunction during the terminal stages of the disease and progressive pulmonary oedema and hydrothorax result in asphyxiation (Uilenberg, 1971; Owen et al., 1973). Amstel et al. (1988a, b) found normal arterial carbon dioxide tension in calves with experimentally induced heartwater, with a tendency towards alkalosis, an increased pulmonary dead space and fluctuations in venous admixture. In the terminal stages of the disease there was a marked decrease in stroke volume and cardiac output.
The pathogenesis of vascular permeability remains speculative as the intracytoplasmic development of the organisms seems to have little detectable cytopathic effect upon the endothelial cells (Pienaar, 1970), and there is also no apparent correlation between the number of parasitized cells in the pulmonary blood vessels and the severity of the pulmonary oedema (Prozesky and Plessis, 1985a; Jackson and Neitz, 1932). It has been proposed that an endotoxin (Amstel et al., 1988a) and increased cerebrospinal fluid pressure (Brown and Skowronek, 1990) play a role in the development of lung oedema.
The course of the disease can vary from a peracute form marked by sudden death with little or no clinical signs, to a chronic form, characterized by a transitory fever, followed by natural recovery. Small ruminants (sheep and goats, particularly Angora goats) appear to be most susceptible. A similar course, to a varying degree, is also seen in cattle.
In general the prognosis is especially poor for imported or exotic cattle and small ruminants. Peracute and acute forms are usually fatal. Few ruminants survive once nervous symptoms have appeared. Case mortalities vary from 5% to virtually 100% depending on the strain of E. ruminantium involved, the locality, season and host breed.
EpidemiologyTop of page
Factors relating to the tick vector, causative organism, and vertebrate host are important in the epidemiology of heartwater. These include possible immunological strain differences of E. ruminantium, availability of wild animal reservoir hosts or vectors for the organisms, infection rate in ticks, age and genetic resistance of domestic ruminant populations, seasonal changes influencing tick abundance and activity, and the intensity of tick control (Uilenberg, 1983).
Although there is a lack of information on the development of E. ruminantium there is some evidence that the parasite undergoes a sequential development in both the vertebrate and invertebrate hosts (Plessis, 1982; Kocan et al., 1987a). The organism replicates mainly by binary fission, and possibly by endosporulation (Pienaar, 1970). It appears that the reticulate bodies are predominately proliferative, while the elementary bodies represent the infective stage (Jongejan et al., 1990). Transmission electron microscope studies of in vitro-cultivated organisms demonstrated the presence of intracellular reticulate bodies 2 to 4 days after infection and intermediate bodies 4 to 5 days after infection. Large numbers of elementary bodies are seen after rupture of endothelial cells 5 to 6 days after infection (Jongejan et al., 1990 and 1991).
The host Amblyomma spp. ticks become infected during the larval and nymphal stages when they feed on infected domestic and wild ruminants, and possibly also on certain game birds and reptiles while E. ruminantium is circulating in the blood of these hosts. Nymphae and adult ticks transmit E. ruminantium to susceptible hosts without losing the infection. Intrastadial transmission has been demonstrated (Andrew and Norval, 1989), but transovarial transmission has only been reported once under laboratory conditions and probably does not occur in nature (Bezuidenhout and Jacobsz, 1986). The development cycle of the organism in the tick and the infectivity of successive stages of the tick are poorly understood. It is thought that after an infected blood meal, initial replication of the organism takes place in the intestinal epithelium of the tick and that the salivary glands eventually become parasitized (Kocan et al., 1987b). Transmission of the parasite to the vertebrate host probably takes place either by regurgitation of their gut contents or through the saliva of the tick while feeding. The minimum period required for transmission of the parasite after tick attachment is between 27-38 hours in nymphs and 21-75 h in adults (Bezuidenhout, 1988).
The main vectors of heartwater are Amblyomma variegatum and A. hebraeum, although a number of other Amblyomma spp. have been shown experimentally to be able to transmit the organism. A. maculatum, occurring in the USA is also capable of transmitting the disease (Uilenberg, 1982b). Not all are equally good vectors, and their importance in the transmission of heartwater depends not only on their vector competence, but also on their distribution and association with domestic stock (Uilenberg, 1983). Furthermore, the activity and abundance of the ticks is influenced by temperature and humidity (Petney et al., 1987). Ticks can acquire the infection from the host from about the time of the febrile reaction for up to 361 days, or even longer (Andrew and Norval, 1989) and probably retain their infectivity for life (Neitz, 1968; Ilemobade, 1976). Infection rates in ticks vary, from 0-44.9% for males, 20-36.1% for females and 0-13.4% for nymphs, depending on the season and the locality in which they are collected (Plessis, 1985; Plessis and Malan, 1987b; Norval et al., 1990).
The existence of antigenically different stocks of E. ruminantium with varying virulence has been demonstrated. There is also variable cross-protection between these different varieties (Jongejan et al., 1988; Plessis et al., 1989). The introduction of animals which are immune to a particular variant of E. ruminantium into an endemic area where a different variety occurs may therefore result in cases of heartwater.
Various factors such as species, breed, age, degree of natural resistance and immune status play a role in determining whether asymptomatic or overt disease will develop in a susceptible host after infection. Young calves, lambs and kids possess a non-specific resistance which is independent of the immune status of the dam and is of short duration: the first 4-6 weeks of life in calves and only the first week in lambs and kids (Neitz and Alexander, 1941; Alexander et al., 1946; Uilenberg, 1981; Plessis et al., 1987). The susceptibilities of different breeds of cattle and sheep vary. Some sheep breeds, such as the Blackhead Persian, possess a certain degree of natural resistance (Alexander, 1931; Uilenberg, 1983). Angora goats are highly susceptible to heartwater and their immunity is of short duration (Plessis et al., 1983). Genetic resistance, which is due to a recessive sex-linked gene, has been demonstrated in Creole goats in Guadeloupe (Matheron et al., 1987).
Wild ruminants such as the blesbok (Damaliscus dorcas phillipsi) and black wildebeest, as well as helmeted guinea fowl, leopard tortoise (Geochelone pardalis) and scrub hare have been shown to harbour E. ruminantium subclinically for long periods and may therefore play a role as source of infection for ticks (Petney and Horak, 1988).
Impact: EconomicTop of page
Most authorities regard heartwater in southern Africa as an economically important disease. Uilenberg (1983) ranked it second only to East Coast fever and tsetse-transmitted trypanosomosis. Neitz (1968) stated that in endemic areas, mortalities due to heartwater were three times as great as those due to babesiosis and anaplasmosis. However, there have been no definitive studies designed to quantify this importance. Recently, under the auspices of the UF/USAID/SADC heartwater research project, a study was undertaken to evaluate the economic impact of heartwater in Zimbabwe. The total annual losses were estimated at US $5.6 million (Mukhebi et al., 1999). Annual economic losses per animal in the commercial production system in Zimbabwe were 25 times greater than losses in the communal system. The greatest components of economic loss were acaricide costs (76%), followed by milk loss (18%) and treatment cost (5%). However, no other reliable figures are available on the economic impact of heartwater in the region.
Zoonoses and Food SafetyTop of page
The fact that E. ruminantium can be grown in human endothelial cells (Totté et al., 1993) has led to speculation that cowdriosis might be a zoonosis. But to date, no evidence from the field supports this (Kelly et al., 1992).
Disease TreatmentTop of page
A variety of drugs have been used with varying success against E. ruminantium (Amstel and Oberem, 1987). Treatment of heartwater during the early febrile stages presents very few problems and recovery can be expected when either sulphonamides or tetracyclines are used at generally accepted dose rates. Tetracyclines are nowadays recognized as more effective than sulfonamides. The successful treatment of field cases of heartwater remains a problem because of the advanced stage of the disease in which the animal is usually presented and because of ineffective supportive therapy. Drugs active in reducing oedema (Shakespeare et al., 1998), stabilization of membranes and blocking effect of vasoactive compounds released with cellular death could be considered (Amstel and Oberem, 1987).
This is a procedure by which a series of tetracycline injections is used to protect susceptible animals against heartwater when they are introduced into an endemic area (Purnell, 1987). In goats it is advocated that short-acting tetracyclines be administered at a dosage rate of 3 mg/kg body weight on 10, 20, 30, 45 and 60 days after introduction, but that the animals should not be dipped until after 60 days (Gruss, 1981). Similarly, injections of long-acting tetracycline formulations (10-20 mg/kg body weight) given on days 7, 14 and 21, or even on only two occasions (days 7 and 14) in cattle are sufficient to protect them from contracting heartwater, while at the same time allowing them to develop a natural immunity (Purnell, 1987). The success of this regime is dependent on all the animals becoming naturally infected with heartwater during the time that they are protected by the drug. With fluctuating infection rates in ticks under different ecological conditions this approach may fail.
Prevention and ControlTop of page
Heartwater can be controlled by immunization of calves, lambs or kids (generally no treatment is necessary following the immunization), treatment of sick animals infected by ticks, and the strategic control of the number of bont ticks to which livestock are exposed.
Sustained, intensive tick control measures may, under certain conditions, succeed in preventing outbreaks of heartwater, even in endemic areas. This, however, should be considered a temporary measure which is accompanied by risks of later outbreaks of the disease if control measures should be relaxed. It is important to remember that, since E. ruminantium replicates in the tissue of the tick, the infection is amplified so that a single tick can transmit the disease to a large ruminant. Strict tick control can succeed in epidemic areas, where the disease normally does not occur and bont ticks could be considered to be only temporary invaders.
In endemic areas, where the disease normally occurs and the tick vector is permanently established, control is more difficult to accomplish and also more costly. The disease can only be controlled successfully if all the animals on the farm can be dipped regularly throughout the year and if there are no, or an absolute minimum, of game and birds on which ticks can survive. Intensive dipping programmes (high frequency dipping) also carry a high risk as far as the development of tick resistance to the dipping compound is concerned. This approach is practically impossible in any extensive farming enterprise.
In marginal (transitional) areas where veld suitable for bont ticks changes to veld in which they cannot survive, control may be difficult. This is because the transitional veld very often consists of bushy gorges and valleys (where the bont tick may occur) that connect heartwater-free middle- or highveld with lower-lying bushveld where the disease occurs regularly. In cases like these the disease can be prevented by a combination of an intensive dipping programme, particularly at strategic times (October-November and March-April) and management aimed at avoiding the grazing of these danger areas.
Strategic Tick Control
This is the level of control that prevents ticks from becoming a nuisance to the animals, but allows sufficient numbers to maintain the animals’ immunity through regular re-infection, is the only practical approach recommended in the vast majority of the heartwater endemic areas of southern Africa. This approach implies that animals become naturally infected by tick exposure, or are immunized, and that their immunity is maintained by regular re-infection through the tick at intervals not exceeding 6-9 months. A dipping compound is applied only when the number of ticks on the animals reaches such a level that it causes tick worry or becomes a threat to the general health of the animal (such as causing damage to the udders of cows).
In the Caribbean Amblyomma program, the acaricide treatment of ruminants with dorsal pour on using remanent acaricide flumethrine such as Bayticol® had been recommended with a frequency of twice treatment per month during 2 years with an initial objective of eradication of the ticks. This method diminished the tick population in several islands and eliminated the tick from 4 of them. An integrated tick control strategy taking into account the recent data on the heterogeneous drop off rhythm of Amblyoma variegatum nymphs has been proposed to reduce pasture infestation by adult ticks (Stachurski et al., 2010). Mathematical models of Amblyomma population dynamics based on biotic and abiotic parameters are being develop with an objective of developing maps of habitat suitability and should allow testing of different control strategies.
Immunization and Vaccines
The only commercial vaccine available consists of the blood of sheep infected with live virulent E. ruminantium organisms (BALL3-strain) (Bezuidenhout, 1989). This vaccine is only used on a large scale in South Africa and to a very limited extent in other African countries.
Vaccination with heartwater-infected sheep blood (=heartwater vaccine) will complement natural tick infection of young animals and also ensure immunity in those animals which escape natural infection. Since the vaccine contains live heartwater organisms, vaccinated, susceptible animals will develop variable degrees of heartwater, which may or may not require appropriate treatment. A number of different vaccination methods are used.
Calves vaccinated before the age of 4-6 weeks usually show no clinical signs but will nevertheless develop an immunity. Although young calves generally do not require any treatment, approximately 10-20% of calves may show a temperature reaction (fever), and a few of these may even develop clinical signs and therefore require treatment. The immunity which thus develops in calves is usually of shorter duration and it is important that the calves be exposed to infected ticks within 3 months. Newborn lambs and kids must be vaccinated within the first week of their lives to avoid clinical reactions and therefore the need for treatment.
It is important that animals be kept under daily observation for at least one month after vaccination, so that additional treatment can be given should any of the animals develop clinical signs. Vaccinated animals should be exposed to bont ticks as soon as possible to boost their immunity. Pregnant animals should not be vaccinated. Susceptible pregnant animals introduced into heartwater endemic areas should be kept free from ticks until they have lambed/calved and thereafter they should be immunized.
The vaccine is administered intravenously into the jugular vein at a dosage of 3 ml for cattle, sheep and goats, irrespective of their size or age (Combrink et al., 1997). The vaccine reaction usually occurs 1-3 weeks after vaccination. The peak of the reaction is normally accompanied by fever (40°C or higher, measured early in the morning), and maybe symptoms such as listlessness, poor appetite, and decreased milk yield, and even nervous symptoms. Animals which develop vaccine reactions must immediately receive appropriate treatment. Immunity to heartwater develops within approximately 4-6 weeks after immunization and immunized animals will be protected against serious disease caused by most (but not all) naturally-occurring strains of heartwater.
This method is risky (possible transmission of other pathogens and possible loss of animals), expensive (cold chain required for storage, close monitoring of animals during one month) and thus inadequate in low-input farming systems. Other experimental vaccines have been developed such as inactivated, attenuated vaccine and recombinant vaccines. The first one is the entire killed bacteria emulsified in oily adjuvant, ISA50. Inactivated vaccines have several advantages as they contain killed bacteria and their storage conditions are compatible with field use (-20°C or refrigerated). Two injections are necessary with one month delay and animals should be protected from tick infestation during at least these 2 months. After natural or experimental challenges, animals develop hyperthermia and are clinically affected. Inactivated vaccine emulsified in ISA50 had been tested both in experimental and field conditions and demonstrated its efficiency (Martinez et al., 1994; Mahan et al., 1995, 1998, 2001; Adakal et al., 2010a).
Lack of vaccine efficiency is related to the diversity of strains within a restricted area. A vaccine including a second local strain improved significantly the protection in a field trial in Burkina Faso (Adakal et al., 2010a). Improvement of the production of E. ruminantium antigen at industrial scale and evaluation of the minimal efficient dose of vaccine (35µg) gives the opportunity to produce this vaccine at low cost (0.11 euros per dose) (Marcelino et al., 2006; Vachiéry et al., 2006; Marcelino et al., 2007). As soon as regional isolates are available in culture after isolation, it could be possible to produce an inactivated vaccine including a cocktail of regional strains. The main difficulty is to choose the strains which could protect against other circulating strains. The choice will depend on genetic characteristics and markers which are not yet defined.
The second experimental vaccine is the in vitro attenuated vaccine. Senegal, Welgevonden and Gardel strains were attenuated by successive in vitro passages and demonstrated their efficiency in experimental conditions (Jongejan et al., 1991b; Zweygarth et al., 2005, 2008; Faburay et al., 2007b). Senegal attenuated vaccine confers good protection against homologous strains but poor protection against heterologous strains. Welgevonden attenuated vaccine confers good protection in controlled conditions against homologous and four different strains but is not yet tested in field conditions. The main disadvantage of attenuated vaccines is the possible reversion to virulence. Moreover as any live vaccine, it should be stored in liquid nitrogen during cold chain storage.
The third vaccine has been developed based on prime DNA/boost recombinant protein vaccine (Pretorius et al., 2008). There was an efficient protective effect of a cocktail of 4 ORFs against homologous challenge but it did not give satisfactory results during field tick challenge. Moreover, simple intramuscular immunisation is not sufficient to induce protection, and the use of gene gun is necessary for prime DNA injection which is not suitable for large vaccination campaign. One polymorphic gene has been identified as an efficient component of a recombinant vaccine against heartwater using prime/boost method (Pretorius et al., 2010). However, as this gene is polymorphic, a recombinant vaccine should include almost 3 different genotypes.
For any kind of vaccine, live, inactivated, attenuated or recombinant vaccines, the main problem is the presence of numerous strains in the field with high genetic diversity and the choice of vaccinal strains genotypes depending on the region.
ReferencesTop of page
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