Definition
East Coast fever (ECF), a form of bovine theileriosis, is a
tick-transmitted protozoal disease of cattle characterized by high fever and
lymphadenopathy. The disease causes high mortalities in breeds nonindigenous to the
endemic areas, and is confined to eastern, central, and parts of southern Africa.
Etiology
The causative agent of classical ECF is Theileria parva.
Some previously recognized separate species and subspecies have been combined with T.
parva as a result of recent studies on their DNA (1,2).
The life cycle of T. parva is complex in its tick and
mammalian hosts (2). Sporozoite stages, produced in large numbers in the acinar cells of
the salivary glands of the infected tick vector, are inoculated along with saliva during
feeding and rapidly enter target lymphocytes, which become transformed after the Theileria
schizont is formed. The infected lymphocyte is transformed into a lymphoblast and divides
in conjunction with the schizont, giving rise to two infected daughter cells. This process
has been termed "parasite-induced reversible transformation" because, if the
cells are treated with antitheileria drugs, the transformed cells revert to quiescent
lymphocytes (3).
Within the infected lymphocytes, schizonts are associated with
microtubules involved in spindle formation during host cell division (2). Clonal expansion
of infected cells occurs with an approximate tenfold increase of schizonts every 3 days.
Schizonts, traditionally called macroschizonts or Koch's blue bodies vary in size and in
the number of nuclei. Early detectable forms are small with nuclei that, when
Giemsa-stained, appear as chromatic granules.
From day 14 after tick infection of cattle, individual schizonts
undergo merogony to produce merozoites (traditionally called microschizonts). Merozoites
invade the erythrocytes to become piroplasms, which may subsequently undergo limited
division also by merogony (4), Piroplasm-infected erythrocytes are ingested by ticks of
the larval or nymphal stages and undergo a sexual cycle in the gut of the replete tick to
produce zygotes, which in turn develop into motile kinete stages that infect the salivary
glands of the next instar, the nymph or adult (5,6).
Host Range
Cattle in endemic areas, particularly the zebu type (Bos
indicus), appear less susceptible to ECF, as do young animals. In addition, introduced
cattle, whether of a taurine, zebu, or sanga breed, are much more susceptible to
theileriosis than cattle from endemic areas. The Indian water buffalo (Bulbalis
bulbalis) is as susceptible to T. parva infection as cattle. The African
buffaloes (Syncerus caffer) are reservoirs of T. parva infection, and it has
recently been proved that waterbucks (Kobus spp.) are also reservoirs (2).
Buffaloes may suffer clinical disease from T. parva infection, but its effects on
waterbuck are unknown. Organisms isolated from buffalo, on repeated passage in cattle,
result in a parasite that produces disease characteristics indistinguishable from those
associated with ECF (2). Hence, the organism causing ECF is assumed to be a cattle-adapted
form of the buffalo parasite causing Corridor disease. Piroplasms can be demonstrated in
most wild antelopes in east Africa, but the relationship of most of them to T. parva is unclear.
Geographic Distribution
The distribution of ECF is strictly associated with the
distribution of the vector tick species. In the case of Rhipicephalus appendiculatus,
the area extends from southern Sudan to South Africa and as far west as Zaire. The range
of T. parva is less than the tick vector for T. parva-free populations of R.
appendiculatus occur in Zambia, Kenya, and South Africa. R. appendiculatus is found
from sea level to over 8,000 feet in areas where there is annual rainfall of over 20
inches (500 mm). Up to three generations of the tick vector can occur per year in
favorable areas of east Africa (Lake Victoria Basin), but there is only one generation a
year in southern and central Africa because of a behavioral diapause, controlled by
photoperiod, in the adult tick (2). This results in a strict seasonal occurrence of the
different tick stages on cattle and a seasonal occurrence of ECF. This behavioral diapause
allows the tick to survive during the long hot dry seasons occurring in the southern parts
of Africa. Tropical theileriosis caused by T. annulata infection does not overlap
in distribution with T. parva and is transmitted by ticks of the genus Hyalomma.
It is distributed in north Africa, down the Nile Valley to Sudan, southern Europe, the
Middle East, and parts of Asia, including the Indian subcontinent and China.
Transmission
Rhipicephalus appendiculatus is the main field vector of
ECF, although in certain areas other field vectors occur, such as R. zembeziensis in drier areas of southern Africa and R. duttoni in Angola. East coast fever is not
maintained in the absence of these field vectors. The rhipicephalid vectors are three-host
ticks, and transmission occurs from stage to stage; transovarian transmission does not
occur. Ticks can remain infected on the pasture for up to 2 years depending on the
climatic conditions and the stage of infection, for adults survive longer than nymphs (2).
The parasite dies out faster in hot climates and in nymphs compared with adults (2).
Normally, for transmission to occur, the infected tick has to attach for several days to
enable sporozoites to mature and be emitted in the saliva of the feeding tick. However,
under high ambient temperatures, ticks on the ground may develop infective Theileria
sporozoites, which can be transmitted to cattle within a few hours after attachment (2).
Unlike other Theileria species and Babesia species, T. parva is not easily transmitted experimentally by blood. Schizont-infected lymphoid tissues have
been used to initiate infection with variable results.
Incubation Period
Under experimental conditions, using either ticks of known
infection or sporozoite stabilate, the incubation period has a medium range of 8 to 12
days. The incubation period may be much more variable in the field owing to differences in
challenges experienced by the cattle and may extend to beyond 3 weeks after attachment of
infected ticks.
Clinical Signs
The first clinical sign of ECF in cattle appears 7 to 15 days
after attachment of infected ticks. This is seen as a swelling of the draining lymph node,
usually the parotid, for the ear is the preferred feeding site of the vector. This is
followed by a generalized lymphadenopathy in which superficial subcutaneous lymph nodes
such as the parotid, prescapular, and prefemoral lymph nodes, can easily be seen and
palpated (Fig. 50). Fever ensues and
continues throughout the course of infection. This rise in temperature is rapid and is
usually in excess of 103° F (39.5° C) but may reach 106° F (42° C). Anorexia develops,
and loss of condition follows. Other clinical signs may include lacrimation, corneal
opacity, nasal discharge, terminal dyspnea, and diarrhea. Before death the animal is
usually recumbent, the temperature falls, and there is a severe dyspnea due to pulmonary
edema that is frequently seen as a frothy nasal discharge. Death usually occurs 18 to 30
days after infestation of susceptible cattle by infected ticks. Mortality in fully
susceptible cattle can be nearly 100 percent. The severity and time course of the disease
depend on, among other factors, the magnitude of the infected tick challenge, for ECF is a
dose- dependent disease, and on the strain of parasites. Some stocks of parasites cause a
chronic wasting disease. A fatal condition called "turning sickness" is
associated with the blocking of brain capillaries by infected cells and results in
neurological signs.
In recovered cattle, chronic disease problems can occur that
result in stunted growth in calves and lack of productivity in adult cattle (17). However,
this syndrome tends to be in the minority of recovered clinical cases; in a majority of
cases, asymptomatic carriers can be recognized with apparently little or no effect on
their productivity (17). A review of the clinical disease is given by Irvin and Mwanachi
(11).
Gross Lesions
A frothy exudate is frequently seen around the nostrils of an
ECF-infected animal. Signs of diarrhea, emaciation, and dehydration may be seen. Lymph
nodes are greatly enlarged and may be hyperplastic, hemorrhagic, and edematous (Fig. 51). In acute cases of ECF, lymph
nodes are edematous and hyperemic but often become necrotic and shrunken in more chronic
disease. Generally, muscles and fat appear normal but, depending on relative acuteness of
infection, fat may become greatly depleted; serosal surfaces have petechial and ecchymotic
hemorrhages, and serous fluids may be present in body cavities. Hemorrhages and ulceration
may be seen throughout the gastrointestinal tract — particularly in the abomasum and
small intestine, where necrosis of Peyer's patches can be observed. Lymphoid cellular
infiltration appear in the liver and kidney as white foci that have been referred to as
pseudoinfarcts. The most striking changes are seen in the lungs. In most cases of ECF,
interlobular emphysema and severe pulmonary edema appear, the lungs are reddened and
filled with fluid, and the trachea and bronchi are filled with fluid and froth.
Morbidity and Mortality
Morbidity and mortality depend on, among other factors, the
magnitude of the infected tick challenge and susceptibility of the host and strain of
parasite. East Coast fever in susceptible cattle, which are not indigenous to the enzootic
area, is very severe with a mortality approaching 100 percent. Animals that recover are
often unthrifty and sickly. Zebu cattle residing for many generations in endemic areas
become infected (100 percent morbidity), but only a minor proportion succumb; however,
many become carriers, and early infection with T. parva can affect their growth and
productivity (17).
Diagnosis
Field Diagnosis
East coast fever is only found in association with its known tick
vectors, Rhipicephalus appendiculatus, R. zembeziensis and possibly R.
duttoni and R. nitens (2). A febrile disease with signs of enlarged lymph nodes
associated with infestation by tick vectors is suggestive of ECF. An acute disease with
high mortality on farms, where tick control is not effectively applied, also is suggestive
of ECF. In many epidemiological situations, high mortality occurs only in calves; the
adult cattle represent immune survivors.
In the field, diagnosis is usually achieved by finding Theileria
parasites in Giemsa-stained blood smears and lymph node needle biopsy smears (Fig. 52).
Specimens for Laboratory
Specimens consisting of buffy coat smears air-dried and fixed in
methanol; lymph node impressions air-dried and fixed in methanol; lymph nodes, spleen,
lung, liver, and kidney samples for histopathology; and serum should be collected.
Laboratory Diagnosis
The demonstration of schizont-infected cells in lymph node samples
is diagnostic of ECF. Small piroplasms in erythrocytes are suggestive of ECF, but
diagnosis must be confirmed by the detection of schizonts. Schizonts can be detected in
sections but are best seen in smears
Antibody in the mammals can be detected by a variety of
serological tests of which the most widely used is the indirect fluorescent antibody test
employing cell culture schizont antigen. Enzyme-linked assays have been developed using
whole parasite lysates or specific antigens isolated by monoclonal antibodies (14).
Because of the often acute nature of the disease, serological tests are useful in
detecting a changed immune status of recovered animals within an exposed herd. Now DNA
technologies can be applied to material from cattle and ticks, including the use of probes
and the polymerase chain reaction (1,2,15,16).
Differential Diagnosis
Identification of schizonts in lymphoid cells is considered to be
pathognomonic of ECF. However, it must be realized that in an area such as Kenya, five
species of Theileria have been recognized in cattle (T. parva, T. mutans, T. velifera, T. taurotragi and T. buffeli) and it is possible for an
individual animal to harbor all these parasites at once (2). Also, all these species
produce schizonts which, except for those of T. mutans, are not morphologically
distinct (2). Piroplasms of Theileria spp. have similar morphology and thus are
difficult to differentiate on blood slides. In addition, recovered animals, particularly
in areas with endemic stability, become carriers of parasites and may show both T.
parva schizont and piroplasm stages without clinical ECF (2).
Theileria parva derived from African buffalo (Syncerus
caffer), which causes Corridor disease in cattle, is characterized by production of
low parasitosis and parasitaemia in cattle although it can result in high fatality rates
(2). Because enzyme-linked immunosorbant assay antibody and antigen tests and DNA probes
are being developed for this range of Theiletia species, it will become easier to
differentiate them in the field. Other species tend to either be of low pathogenicity (T.
mutans, T. taurotragi, T. buffeli) or avirulent (T. velifera) in
cattle.
T. annulata is the cause of Mediterranean or tropical
theileriosis, which is also a severe disease of cattle; although it is endemic in northern
Africa, there is no evidence that its distribution overlaps with that of T. parva (2).
Gross postmortem lesions of ECF may be confused with those of a
variety of diseases such as the following:
1. Heartwater because of pulmonary edema and hydrothrax.
Examination of brain smears and lymph node or spleen impression smears can differentiate
between the two diseases.
2. Trypanosomiasis because of edema, lymphadenopathy, and anemia.
Blood and lymph node smear examination will normally differentiate between the two
diseases.
3. Babesiosis and anaplasmosis because of anemia. These diseases
can easily be differentiated from ECF on examination of blood smears.
4. Malignant catarrhal fever because of lymphadenopathy and
corneal opacity. Examination of blood and lymph node smears will clearly differentiate
between the two diseases.
Treatment
There are currently three effective drugs for the treatment of
ECF: parvaquone (Clexon), buparvaquone (Butalex), and halofuginone lactate (Terit). Each
of these drugs has been introduced to the market within the last 15 years (2). The
availability of a therapeutic means of controlling ECF is a significant development.
However, there are two constraints to widespread use of medication: the drugs are too
expensive for most African farmers, and rapid, accurate diagnosis is required for
effective therapy (2).
Vaccination
Methods of immunization using live parasites have been summarized
by Cunningham (19). The most successful involves an "infection and treatment
method" initially using oxytetracycline and more recently the newer drugs mentioned
above. Animals are inoculated with a potentially lethal dose of infective sporozoite
stabilate prepared from ticks and treated either simultaneously (tetracyclines,
buparvaquone) or subsequently (parvaquone and halofuginone) with a drug. Sporozoite
stabilates are produced from adult ticks fed as nymphs on infected cattle; the adult ticks
are ground up in a medium after prefeeding on rabbits for 4 days, and the sporozoite
suspension is prepared by centrifugation and cryopreserved as a stabilate. Extensive
trials have been carried out using infection and treatment immunization in the ECF zones,
and this method is now approved for field use by several countries in Africa (2). Problems
do occur in the recognition of suitable antigenic stocks for immunization (2), and any
vaccination scheme can only follow after a careful assessment of the local complex of T.
parva parasites. In any population of T. parva parasites in the field, an
isolate may constitute several strains. This method of immunization requires a reliable
cold chain and extensive monitoring. Recovery from ECF usually results in an excellent
immunity to homologous or related stocks of the parasite, lasting for over 3 1/2 years in
the absence of reinfection.
The nature of the immunity has been studied in some detail (2,20).
Neutralizing antibodies to sporozoites have been recognized and related to a 67Kda antigen
on the surface of the sporozoites. This has been synthesized by recombinant technology and
has been shown to provide a degree of protection to cattle immunized with it (21). Other
potential protective antigens have been recognized associated with the surfaces of
sporozoites and schizont-infected cells (2).
Control
Preventive Measures
The current primary method of controlling ECF in cattle is
immunization (see vaccination) and treatment of cattle with chemical acaricides. A number
of acaricides, mainly organochlorides and organophosphorus compounds but recently
synthetic pyrethroids and amidens, are applied in dips, spray races, or by handspraying.
More recently, "pour on" or "spot on" formulations have been
introduced. The application is usually on a weekly basis, but this rate has to be
increased when the challenge is high. The cost of this control measure is becoming
exorbitant, and the farming economies in many countries in Africa are not able to afford
it. This may well be an advantage because this level of acaricide exposure leads to
resistance of vectors, unacceptable residues in milk and meat, and, where successful, the
creation of an epidemic instability with a large proportion of the cattle population
becoming susceptible. A more rational approach using integrated control has been suggested
by Young et al (18). These measures include effective fencing, pasture management,
rotational grazing to reduce the level of challenge, selection of tick resistant cattle,
and new methods of immunization; with strategic acaricide application, this approach
offers a more satisfactory method of ECF control (18).
Sanitation and disinfection measures other than those associated
with tick control are not applicable to ECF .
Public Health
Theileria spp. show a high degree of host specificity for
both the vector and the mammalian host. Theileria parva does not infect man.
GUIDE TO LITERATURE
1. CONRAD, P.A., ole-MOIYOI, O.K., BALDWIN, C. L., DOLAN, T.T.,
O'CALLAGHAN, C.J., NJAMUNGGEH, R.E.G., GROOTENHUIS, I.G., STAGG, D.A., LEITCH, B.L., and
YOUNG, A.S. 1989. Characterization of buffalo-derived theilerial parasites with monoclonal
antibodies and DNA probes. Parasitology, 98: 179-188.
2. NORVAL, R.A.I., PERRY, B.D., and YOUNG, A.S. 1992. The
Epidemiology of Theileriosis in Africa. London:Academic Press, 481 pp.
3. ole-MOIYOI, O.K., 1989. Theileria parva: An intracellular
parasite that induces reversible Iymphocyte transformation. Exptl. Parasitol., 69:
204-210.
4. CONRAD, P.A., DENHAM, D., and BROWN, C.G.D. 1986.
Intraerythrocytic multiplication of Theileria parva in vitro: An ultrastructural study.
Internat. J. Parasitol., 16: 223-230.
5. MEHLHORN, H., and SCHEIN, E. 1984. The piroplasms: Life cycle
and sexual stages. Adv. Parasitol., 23: 37-103.
6. FAWCET, D.W., YOUNG, A.S., and LEITCH, B.L. 1985. Sporogony in
Theileria (Apicomplexa: Piroplasmida): A comparative ultrastructural study. J. Submicro.
Cytol., 17: 299-314.
7. MALMQUIST, W. A., NYINDO, M.B.A., and BROWN, C.G.D. 1970. East
Coast Fever. Cultivation in vitro of bovine spleen cells lines infected and transformed by
Theileria parva. Trop. Anim. Hlth Prod., 2:139-145.
8. CUNNINGHAM, M.P., BROWN, C.G.D., BURRIDGE, M.J., and PURNIELL,
R.E. 1973. Cryopreservation of infective particles of Theileria parva. Internat. J.
Parasitol., 3:583-587.
9. BROWN, C.G.D., STAGG, D.A., PURNELL, R.E., KANHAI, G.K., and
PAYNIE, R.C. 1973. Infection and transformation of bovine Iymphoid cells in vitro by
infective particles of Theileria parva. Nature, 245: 101-103.
10. DOLAN, T.T., TEALE, A.J., STAGG, D.A., KEMP, S.J., COWAN,
K.M., YOUNG, A S., GROOCOCK, C.M., LEITCH, B.L., SPOONER, R.L., and BROWN, C.G.D. 1984.
Histcompatability barrier to immunization against East Coast fever using Theileria parva
infected lymphoblastoid cell lines. Parasit. Immunol., 6: 243-250.
11. IRVIN, A.D., and MWANACHI, D. M. 1983. Clinical and diagnostic
features of East Coast fever (Theileria parva) infection of cattle. Vet. Rec., 113:
191-198.
12. De KOCK, G. 1957. Studies on the lesion and pathogenesis of
East Coast fever (Theileria parva infection) in cattle with special reference to the
lymphoid tissues. Onderst. J. Vet. Sci., 27:431.
13. MORRISON, W.I., BOSCHER, G., MURRAY, M., EMERY, D.L., MASAKE,
R.A., COOK, R.H., and WELLS, P.W. 1981. Theileria parva: Kinetics of infection in lymphoid
system of cattle. Exptl. Parasit., 52: 248-260.
14. KATENDE, J.M., GODDEERIS, B.M., MORZARIA, S.P., NKONGE, C.G.,
and MUSOKE, A.J. 1990. Identification of Theileria mutans — specific antigen for use
in an antibody and antigen detection ELISA. Parasit. Immunol., 12: 419-433.
15. CHEN, P.P., CONRAD, P.A., ole-MOIYOI, O.K., BROWN, W.C., and
DOLAN, T.T. 1991. DNA probes detect Theileria parva in salivary glands of Rhipicephalus
appendiculatus ticks. Parasitol. Res., 77: 590-594.
16. BISHOP, R., SOHANPAL, B., KARIUKI, D.P., YOUNG, A.S., NENE,
V., BAYLIS, H., ALLSOPP, B.A., SPOONER P.R., DOLAN, T.T., and MORZARIA, S. P. 1992.
Detection of a carrier state in Theileria parva infected cattle using the polymerase chain
reaction. Parasitology, 104:
17. MOLL, G., LOHDING, A., YOUNG, A.S., and LEITCH, B.L. 1986.
Epidemiology of theileriosis in calves in an edemic area of Kenya. Vet. Parasitol., 19:
255-273.
18. YOUNG, A.S., GROOCOCK, C.M., and KARIUKI, D.P. 1988.
Integrated control of ticks and tick-home diseases of cattle in Africa. Parasitology, 96:
403-441.
19. CUNNINGHAM, M.P. 1977. Immunization of Cattle Against
Theileria parva. In Immunity to Blood Parasites of Animals and Man. L.H. Miller, J.
A. Pino, and J.J. McKelvey, Jr., eds., New york:Plenum Press, pp. 189-207.
20. MORRISON, W.L., GODDEERIS, B.M., BROWN, W.C., BALDWIN, C.L.,
and TEALE, A.J. 1989. Theileria parva in cattle: Characterization of infected lymphocytes
and the immune response they provoke. Vet. Immun. Immuno. Path., 20: 213-217.
21. MUSOKE, A.J., MORZARIA, S. P., NKONGE, C., JONES, E., and
NENE, V. 1992. A recombinant sporozoite surface antigen of Theileria parva induces
protection in cattle. Proc. Nat. Acad. Sci. USA., 89: 514-519.
A. S. Young, Ph.D., ARCS, ILRAD, Nairobi (deceased)
C. M. Groocock, D.V.M. Ph.D., USDA-APHIS-IS, Vienna, Austria
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