Updated: 20.12.2005
Manual of Diagnostic Tests
and Vaccines for Terrestrial Animals
PART 2
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SECTION 2.5.
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Chapter 2.5.6.
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Summary
? - Index


CHAPTER 2.5.6.

EQUINE PIROPLASMOSIS


 

 

SUMMARY

Equine piroplasmosis or babesiosis is a tick-borne protozoal disease of horses, mules, donkeys and zebra. The aetiological agents are blood parasites named Theileria equi and Babesia caballi. Theileria equi was previously designated as Babesia equi. Infected animals may remain carriers of these parasites for long periods and act as sources of infection for ticks, which act as vectors. The parasites are found inside the red blood cells of the infected animals.
 
The introduction of carrier animals into areas where tick vectors are prevalent can lead to an epizootic spread of the disease.
 
Identification of the agent: Infected horses can be identified by demonstrating the parasites in stained blood or organ smears. Romanovsky-type staining methods, such as Giemsa, give the best results. In carrier animals, low parasitaemias make it extremely difficult to detect parasites, especially in the case of B. caballi infections, although they may sometimes be demonstrated by using a thick blood smear technique.
 
Paired merozoites, conected at one end, are a diagnostic feature of B. caballi infection. The merozoites of T. equi are less than 2–3 µm long, and are round or amoeboid. A characteristic of T. equi is the arrangement of four parasites in a tetrad or ‘Maltese cross’.
 
When equivocal results are encountered in serological tests, the inoculation of a large quantity of whole blood transfused into a susceptible splenectomised horse will assist diagnosis. The recipient horse is observed for clinical signs of disease and its red blood cells are examined for parasites. Alternatively, a specific tick vector is fed on a suspect animal and Babesia/Theileria may then be identified either in the vector or through transmission by the vector to another susceptible animal.
 
Serological tests: Infections in carrier animals are best demonstrated by testing their sera for the presence of specific antibodies.
 
Currently, the indirect fluorescent antibody (IFA) test and enzyme-linked immunosorbent assays (ELISAs) are the prescribed tests for international trade. The IFA test can be used to distinguish between T. equi and B. caballi infections. ELISA may be used to detect antibodies to both species in infected horses, although cross-reactions between T. equi and B. caballi occur, so that this cannot yet be recommended as a differential diagnostic test. However, recent advances using a recombinant T. equi and B. caballi merozoite proteins and monoclonal antibodies to these proteins in competitive inhibition ELISA appear to be very promising. The competitive ELISA may be superior to complement fixation (CF) tests, especially for detecting long-term infected animals in which the CF titre has waned but which remain IFA seropositive. Unlike the indirect ELISA, the competitive ELISA has been shown to be highly specific for each of the two species of piroplasmosis agent involved.
 
Requirements for vaccines and diagnostic biologicals: There are no biological products available.
 

A. INTRODUCTION

Equine piroplasmosis or babesiosis is a tick-borne protozoal disease of horses, mules, donkeys and zebra. The aetiological agents of equine piroplasmosis are Theileria equi and Babesia caballi. Twelve species of ixodid ticks in the genera Dermacentor, Rhipicephalus and Hyalomma have been identified as transstadial vectors of B. caballi and T. equi, while eight of these species were also able to transmit B. caballi infections transovarially (4, 28, 35, 36). Infected animals may remain carriers of these blood parasites for long periods and will act as sources of infection for tick vectors.
 
The parasites occur in southern Europe, Asia, countries of the Commonwealth of Independent States, Africa, Cuba, South and Central America, and certain parts of the southern United States of America. Theileria equi has also been reported from Australia (but, apparently never established itself in this region), and is now believed to have a wider general distribution than B. caballi.
 
During the life cycle of Babesia, the merozoites invade red blood cells (RBCs) where they transform into trophozoites (7, 29). In this situation the trophozoites grow and divide into two round, oval or pear-shaped merozoites. The mature merozoites are now capable of infecting new RBCs and the division process is then repeated.
 
Babesia caballi: the merozoites in the RBCs are pear-shaped, 2–5 µm long and 1.3–3.0 µm in diameter (20). The paired merozoites, connected at one end, are considered to be a diagnostic feature of B. caballi infection (25).
 
Theileria equi: the merozoites of this organism are relatively small, less than 2–3 µm long (20), and are round or amoeboid. Four parasites are to be found together in the form of a tetrad or so-called ‘Maltese cross’ arrangement. This is a characteristic feature of T. equi (10).
 
It has been shown that sporozoites inoculated into horses via a tick bite invade the lymphocytes (32). The sporozoites undergo development in the cytoplasm of these lymphocytes and eventually form Theileria-like schizonts. Merozoites released from these schizonts enter RBCs. The taxonomic position of T. equi has been controversial and only relatively recently has it been redescribed as a Theileria (26). Further support for the close relation with Theileria spp. also comes from the homology found between 30 and 34 kDa T. equi surface proteins and similar sized proteins of various Theileria spp. (15, 17). However, comparison of the small subunit ribosomal RNA genes of various Babesia, Theileria and Cytauxzoon parasites indicates that T. equi falls into a distinct group different from both the Babesia and Theileria groups (1).
 
The clinical signs of equine piroplasmosis are often nonspecific, and the disease can easily be confused with other conditions.
 
Piroplasmosis can occur in peracute, acute, subacute and chronic forms. The acute cases are more common, and are characterised by a fever that usually exceeds 40°C, reduced appetite and malaise, elevated respiratory and pulse rates, congestion of mucous membranes, and faecal balls that are smaller and drier than normal.
 
Clinical signs in subacute cases are similar. In addition, affected animals show loss of weight, and the fever is sometimes intermittent. The mucous membranes vary from pale pink to pink, or pale yellow to bright yellow. Petechiae and/or ecchymoses may also be visible on the mucous membranes. Normal bowel movements may be slightly depressed and the animals may show signs of mild colic. Mild oedematous swelling of the distal part of the limbs sometimes occurs.
 
Chronic cases usually present nonspecific clinical signs such as mild inappetence, poor performance and a drop in body mass. The spleen is usually found to be enlarged on rectal examination.
 
A rare peracute form where horses are found either dead or moribund has been reported (21).
 

B. DIAGNOSTIC TECHNIQUES

1.   Identification of the agent
 
     Horses that are already infected may be identified by demonstrating the parasites in stained blood or organ smears. Romanovsky-type staining methods, such as the Giemsa method, usually give the best results (34).
 
      The low parasitaemias of carrier animals make it extremely difficult to detect the parasites in smears, especially in the case of infections with B. caballi. When they occur at such low levels, experienced workers can sometimes detect them by the use of a thick blood smear technique (23). Thick films are made by placing a small drop (approximately 50 µl) of blood on to a clean glass slide. This droplet is then air-dried, heat fixed at 80°C for 5 minutes, and stained in 5% Giemsa for 20–30 minutes. An accurate identification of the species of the parasite is sometimes desirable, as mixed infections of T. equi and B. caballi probably occur frequently.
 
     It is very difficult to diagnose equine piroplasmosis in carrier animals by detection of parasites in the blood and serological methods are preferred for this (see below). However, false-negative or false-positive reactions may be encountered in the course of serological tests (5, 6, 38). In such cases, the passage of whole blood, although a cumbersome and expensive exercise, is a very useful technique to determine the true position. Large quantities of whole blood (500 ml) are transfused into a susceptible, preferably splenectomised, horse. This animal is then kept under close observation for clinical signs of disease. Diagnosis is confirmed by the presence of parasites in its RBCs.
 
     In an additional technique, a specific clean tick vector is fed on a suspect animal, and the organism can then either be identified in the tick itself, or through the transmission of the organism by the tick vector to another susceptible animal.
 
     Success in the establishment of long-term in-vitro cultures of T. equi and B. caballi may be one alternative to supplement the methods described above, in order to identify carriers of the parasites (11, 12, 42, 43). Babesia caballi parasites were successfully cultured from the blood of two horses that tested negative by the complement fixation (CF) test (12). Similarly, T. equi could be cultured from horses that did not show any patent parasitaemias at the time of the initiation of the cultures (42, 43).
 
     Molecular techniques for the detection of T. equi and B. caballi have been described (2) including a biotin-labelled DNA probe in a polymerase chain reaction-based assay for the detection of T. equi (31).
 
2.   Serological tests
 
     It is extremely difficult to diagnose the organisms in carrier animals by means of the microscopic examination of blood smears. Furthermore, it is by no means practical on a large scale. The serological testing of animals is therefore recommended as a preferred method of diagnosis, especially when horses are destined to be imported into countries where the disease does not occur, but the vector is present.
 
     Sera should be collected and dispatched to diagnostic laboratories in accordance with the specifications of that laboratory. Horses for export that have been subjected to serological tests and shown to be free from infection, should be kept free of ticks to prevent accidental infections.
 
      A number of serological techniques have been used in the diagnosis of piroplasmosis, such as the indirect fluorescent antibody (IFA) test, the enzyme-linked immunosorbent assay (ELISA) and the CF test.
 
      a)    Indirect fluorescent antibody test (a prescribed test for international trade)
 
            The IFA test has been successfully applied to the differential diagnosis of T. equi and B. caballi infections (22). The recognition of a strong positive reaction is relatively simple, but any differentiation between weak positive and negative reactions requires considerable experience in interpretation. A detailed description of the protocol of the IFA test has been given (22, 27). An example of an IFA protocol is given below.
 
                Antigen production
 
            Blood for antigen is obtained from horses with a rising parasitaemia, ideally 2–5%. Carrier animals are not suitable for antigen production as they have already produced antibodies. Blood (about 15 ml) is collected into 235 ml of phosphate buffered saline (PBS), pH 7.2. The RBCs are washed three times in cold PBS (1000 g for 10 minutes at 4°C). The supernatant fluid and the white cell layer are removed after each wash. After the last wash, the packed RBCs are reconstituted to normal volume with 4% bovine serum albumin fraction V made up in PBS, i.e. the original packed cell volume = 30%, so that one-third consists of RBCs. If the original RBC volume is 15 ml, then 5 ml of packed RBCs + 10 ml of 4% bovine albumin in PBS constitutes the antigen. After thorough mixing, the antigen is placed in prepared wells on a glass slide using a template or a syringe (27). Alternatively, the cells can be spread smoothly on to microscope slides, covering the entire slide with an even, moderately thick film. These slides are allowed to dry, wrapped in soft paper and sealed in plastic bags or wrapped in aluminium foil, and stored at –20°C for up to 1 year.
 
                Test procedure
 
            i)    Each sample of serum is tested against an antigen of B. caballi and of T. equi.
 
            ii)    Prior to use, the antigen smears are removed from storage at –20°C and incubated at 37°C for 10 minutes.
 
            iii)    The antigen smears are then removed from their protective covering and fixed in cold dry acetone (–20°C) for 1 minute. Commercially produced slides are available that are pre-fixed.
 
            iv)    If smears were prepared, squares (14–21 in number, i.e. 2–3 rows of 7 each) are formed on the antigen smears with nail varnish or rapidly drying mounting medium (i.e. Cystoseal).
 
            v)    Test, positive and negative control sera are diluted from 1/80 to 1/1280 in PBS. Negative and positive control sera are included in each test.
 
            vi)    Sera are applied (10 µl each) at appropriate dilutions to the different wells or squares on the antigen smear, incubated at 37°C for 30 minutes, and washed several times in PBS and once in water.
 
            vii)    An anti-horse immunoglobulin prepared in rabbits and conjugated with fluorescein isothiocyanate (this conjugate is available commercially) is diluted in PBS and applied to the smear, which is then incubated and washed as before.
 
            viii)    After the final wash, two drops of a solution containing equal parts of glycerin and PBS are placed on each smears and mounted with a cover-slip.
 
            ix)    The smear are then examined under the microscope for the fluorescing parasites. Sera diluted 1/80 or more that show strong fluorescence are usually considered to be positive, although due consideration is also given to the patterns of fluorescence of the positive and negative controls.
 
      b)    Enzyme-linked immunosorbent assay (a prescribed test for international trade)
 
            The production of recombinant antigens for the use in ELISAs has been described. The recombinant T. equi merozoite protein (EMA-1) has been produced in Escherichia coli (18) and in insect cells by baculovirus (41). Recombinant T. equi Be 82 gene product fused with glutathione S-transferase fusion protein antigen has also been produced in E. coli (9). Recombinant B. caballi rhoptry-associated protein antigen has been produced in E. coli (13, 14). Recombinant antigens produced in E. coli or by baculovirus have the obvious advantage of removing the need to infect horses for antigen production, and they provide a consistent source of antigen for international distribution and standardisation. Recombinant antigens have been used in the indirect ELISA (13) and the competitive inhibition ELISA (C-ELISA) (40). EMA-1 and a specific monoclonal antibody (MAb) that defines this merozoite surface protein epitope, have been used in a C-ELISA for T. equi (18). This C-ELISA overcomes the problem of antigen purity, as the specificity of this test depends only on the MAb used. A 94% correlation was shown between the C-ELISA and the CF test in detecting antibodies to T. equi. Sera that gave discrepant results were evaluated for their ability to immunoprecipitate 35S-methionine-labelled in-vitro translated products of T. equi merozoite mRNA. Samples that were C-ELISA positive and CF test negative clearly precipitated multiple T. equi proteins. However, immunoprecipitation results with serum samples that were C-ELISA negative and CF test positive were inconclusive (19). Limited data at this stage would suggest that the C-ELISA is specific for T. equi (19). This C-ELISA for T. equi was also recently validated in Morocco and Israel, giving a concordance of 91% and 95.7% with the IFA test, respectively (30, 33).
 
            A similar C-ELISA has been developed using the recombinant B. caballi rhoptry-associated protein 1 (RAP-1) and an MAb reactive with a peptide epitope of a 60 kDa B. caballi antigen (14). The results of 302 serum samples tested with this C-ELISA and the CF test showed a 73% concordance. Of the 72 samples that were CF test negative and C-ELISA positive, 48 (67%) were shown to be positive on the IFA test, while four of the five samples that tested CF test positive and C-ELISA negative were positive on the IFA test (14).
 
            A test protocol for an equine piroplasmosis C-ELISA has been described and used for additional validation studies (16, 40). The apparent specificity of the B. equi and B. caballi C-ELISAs lay between 99.2% and 99.5% using sera from 1000 horses presumed to be piroplasmosis-free. One thousand foreign-origin horses of unknown infection status were tested by the C-ELISA and the CF test with an apparent greater sensitivity of the C-ELISA. The results were 1.1% (B. equi) and 1.3% (B. caballi) more seropositive animals detected by C-ELISA than by the CF test; the additional positive results were confirmed by IFA testing. Eight experimentally infected horses (four for B. equi, four for B. caballi) were serially tested from 4 to 90 days post-exposure. Both C-ELISA procedures were again found to more sensitive than the CF test for the detection of infected animals; the results were confirmed by IFA testing. Seroconversion was detected by C-ELISA as soon as or sooner than by the CF test. Both tests were highly reproducible well-to-well, plate-to-plate, and day-to-day, with overall variances of ± 1.2% and ±1.6% for the B. equi and B. caballi tests, respectively.
 
            An example of a C-ELISA protocol is given below.
 
                Solutions
 
            Antigen coating buffer: prepare the volume of antigen coating buffer required by using the following amounts of ingredients per litre: 2.93 g sodium bicarbonate; 1.59 g sodium carbonate; sufficient ultra-pure water to dissolve, and make up to 1 litre with ultra-pure water. Adjust to pH 9.6.
 
            C-ELISA wash solution (high salt diluent): prepare the volume of C-ELISA wash solution required by using the following amounts of ingredients per litre: 29.5 g sodium chloride; 0.22 g monobasic sodium phosphate; 1.19 g dibasic sodium phosphate; 2.0 ml Tween 20; sufficient ultra-pure water to dissolve, and make up to 1 litre with ultra-pure water. Mix well. Adjust pH to 7.4. Sterilise by autoclaving at 121°C.
 
                Antigen production
 
            Frozen transformed E. coli culture is inoculated at a 1/10,000 dilution into any standard non-selective bacterial growth broth (e.g. Luria broth) containing added carbenicillin (100 µg/ml) and isopropyl-thiogalactoside (IPTG, 1 mM). Cultures are incubated on an orbital shaker set at 200 rpm at 37°C overnight. Cells grown overnight are harvested by centrifugation (5000 g for 10 minutes), washed in 50 mM Tris/HCl and 5 mM ethylene diamine tetra-acetic acid (EDTA) buffer, pH 8.0, and harvested again as before. (Antigen is available from the National Veterinary Services Laboratories, P.O. Box 844, Ames, Iowa 50010, USA.)
 
            Cells are resuspended to 10% of the original volume in the Tris/EDTA buffer to which 1 mg/ml of lysozyme has been added, and incubated on ice for 20 minutes. Nonidet P-40 detergent (NP-40) is then added to a final 1% concentration (v/v), vortexed, and the mixture is incubated on ice for 10 minutes. The material is next sonicated four times for 30 seconds each time at 100 watts, on ice, allowing 2 minutes between sonications for the material to remain cool. The sonicate is centrifuged at 10,000 g for 20 minutes. The resulting supernatant is dispensed in 0.5 ml aliquots in microcentrifuge tubes and may then be stored at –70°C for several years. The presence of heterologous host bacterial antigens does not interfere with the binding of specific equine anti-piroplasma antibodies or the binding of the paired MAbs to their respective expressed recombinant antigen epitopes and is confirmed by the following procedures. The antigen-containing supernatants are quality controlled by titrating them with their paired MAbs and with reference monospecific equine antisera to verify both an adequate level of expression and complete specificity for the homologous species of piroplasmosis agent. Normal serum (negative serum) controls must not interfere with binding of the monoclonals or positive equine reference sera to the expressed antigen preparation.
 
                Test procedure
 
            i)    Microtitration plates are prepared by coating the wells with 50 µl of either B. equi antigen or B. caballi antigen diluted in antigen-coating buffer. The dilution used is determined by standard serological titration techniques. The plate is sealed with sealing tape, stored overnight at 4°C, and frozen at –70°C.
 
            ii)    The biotin-labelled anti-murine IgG is diluted in sterile water as directed by the manufacturer, stored at 4°C, and further diluted at the time of use in C-ELISA wash solution to which 2% (v/v) normal equine serum has been added. The avidin–alkaline phosphatase enzyme conjugate is also diluted in C-ELISA wash solution, and the chromogenic enzyme substrate is mixed according to the manufacturer’s instructions.
 
            iii)    Plates are thawed at room temperature, the coating solution is decanted, and the plates are washed twice with C-ELISA wash solution.
 
            iv)    Undiluted equine sera (50 µl/well) is added to wells. Serum should not be heat-treated. Each serum is tested in duplicate wells. Plates are incubated at 37°C for 40 minutes in a humid chamber.
 
            v)    All wells then receive 50 µl/well of diluted anti-B. equi or anti-B. caballi monoclonal antibody. (The MAb is produced in a cell culture bioreactor and is available from the National Veterinary Services Laboratories, P.O. Box 844, Ames, Iowa 50010, USA.) Plates are incubated for 30 minutes at 37°C in a humid chamber, and then washed three times in C-ELISA wash solution.
 
            vi)    Diluted biotinylated anti-murine IgG (50 µl/well) is added to wells. Plates are incubated for 20 minutes at 37°C in a humid chamber, and then washed four times in C-ELISA wash solution.
 
            vii)    Avidin–alkaline phosphatase conjugate (50 µl/well) is added to all wells. Plates are incubated, covered, for 15 minutes at room temperature, and then washed three times in C-ELISA wash solution.
 
            viii)    Chromogenic enzyme substrate (50 µl/well) is added to wells, and plates are incubated with shaking at room temperature during colour development.
 
            ix)    The colour development is stopped by adding 50 µl EDTA stop solution (2.5% [w/v] solution of EDTA in ultra-pure water) to all wells when the negative serum control wells have an optical density of 0.2–0.4 at 590 nm wavelength (OD590).
 
            x)    The plates are read at 590 nm. The average OD590 is calculated for the duplicate wells for all sera. For a valid test, the positive control serum must reduce the amount of the colour by 70–90% compared with the negative control serum and the coefficients of variation of the negative and positive control sera cannot exceed 10%. Duplicate serum sample well values must be within 10% of each other or the sample must be retested, except for strongly positive sera that give OD590 values so close to zero that duplicate numerical precision is unlikely.
 
    xi) If the OD590 of the test serum is reduced by 70% or more relative to the negative control serum, the test serum is considered to be positive. For example, if the mean negative control serum OD590 is 0.04, then test sera with mean OD590 values of 0.12 or less would be positive.
 
      c)    Complement fixation
 
            The CF test is the primary test used by some countries to qualify horses for importation (37). A detailed description of antigen production and a test protocol has been given, for example by the United States Department of Agriculture (USDA) (3, 6, 39). Because the CF test may not identify all infected animals, especially those that have been treated, and because of the anti-complementary reactions produced by some sera, and of the inability of IgG(T) (the major immunoglobulin isotype of equids) to fix complement (24), the IFA test and the ELISA have been accepted for use as prescribed tests for international trade. An example of a CF test protocol is given below.
 
                Solutions
 
            Alsever's solution: prepare 1 litre of Alsever's solution by dissolving 20.5 g glucose; 8.0 g sodium citrate; 4.2 g sodium chloride in sufficient distilled water. Adjust to pH 6.1 using citric acid, and make up the volume to 1 litre with distilled water. Sterilise by filtration.
 
            Stock veronal buffer (5x): dissolve the following in 1 litre of distilled water: 85.0 g sodium chloride; 3.75 g sodium 5,5 diethyl barbituric; 1.68 g magnesium chloride (MgCl2.6H2O); 0.28 g calcium chloride. Dissolve 5.75 g of 5,5 diethyl barbituric acid in 0.5 litre hot (near boiling) distilled water. Cool this acid solution and add to the salt solution. Make up to 2 litres with distilled water and store at 
4°C. To prepare a working dilution, add one part stock solution to four parts distilled water. The final pH should be from 7.4 to 7.6.
 
                Antigen production
 
            Blood is obtained from horses with a high parasitaemia (e.g. 3–7% parasitaemia for B. caballi and 60–85% for T. equi), and mixed with equal volumes of Alsever's solution as an anticoagulant. The plasma/Alsever's supernatant and buffy coat are removed when the RBCs have settled to the bottom of the flask. The RBCs are washed several times with cold veronal buffer and then disrupted. The antigen is recovered from the lysate by centrifugation at 30,900 g for 30 minutes.
 
            The recovered antigen is washed several times in cold veronal buffer by centrifugation at 20,000 g for 15 minutes. Polyvinyl pyrrolidone 40,000 (1–5%[ w/v]) is added as a stabiliser and the preparation is mixed on a magnetic stirrer for 30 minutes, strained through two thicknesses of sterile gauze, dispensed into 2 ml volumes and freeze-dried. The antigen can then be stored at below –50°C for several years.
 
                Test procedure
 
            i)    The specificity and potency of each batch of antigen should be checked against standard antisera of known specificity and potency. Optimal antigen dilutions are also determined in a preliminary checkerboard titration.
 
            ii)    Test sera are inactivated for 30 minutes at 58°C (donkey and mule sera are inactivated at
62.5°C for 35 minutes) and tested in dilutions of 1/5 to 1/5120. Veronal buffer is used for all dilutions.
 
            iii)    Complement is prepared and titrated spectrophotometrically to determine the 50% haemolytic dose (C'H50) (36) and used in the test at five times C'H50. The haemolytic system consists of equal parts of a 2% sheep (RBC) suspension and veronal buffer with 5 minimum haemolytic doses (MHDs) of haemolysin (amboceptor) (40). Some laboratories use twice the 100% haemolytic dose, which gives equivalent sensitivity.
 
            iv)    The test has been adapted to microtitration plates (8). The total volume of the test is 0.125 ml, made up of equal portions (0.025 ml) of antigen, complement (five times C'H50) and diluted serum. Incubation is performed for 1 hour at 37°C.
 
            v)    A double portion (0.05 ml) of the haemolytic system is added and the plates are incubated for a further 45 minutes at 37°C with shaking after 20 minutes.
 
            vi)    The plates are centrifuged for 1 minute at 200 g before being read over a mirror.
 
            vii)    A lysis of 50% is recorded as positive, with the titre being the greatest serum dilution giving 50% lysis. A titre of 1/5 is regarded as positive. A full set of controls (positive and negative sera) must be included in each test as well as control antigen prepared from normal (uninfected) horse RBCs.
 
            Anticomplementary samples are examined by the IFA test. Donkey sera are frequently anticomplementary.
 

C. REQUIREMENTS FOR VACCINES AND DIAGNOSTIC BIOLOGICALS

No biological products are available currently.
 

REFERENCES

1.   Allsopp M.T.E.P., Cavalier-Smith T., De Waal D.T. & Allsopp B.A. (1994). Phylogeny and evolution of the piroplasms. Parasitol., 108, 147–152.
 
2.   Bashiruddin J.B., Camma C. & Rebelo E. (1999). Molecular detection of Babesia equi and Babesia caballi in horse blood by PCR amplification of part of the 16S rRNA gene. Vet. Parasitol., 84, 75–83.
 
3.   Brown G.M. (1979). Equine piroplasmosis complement fixation test antigen production. USDA, APHIS, National Veterinary Services Laboratories. Diagnostic Reagents Laboratory, Ames, Iowa. USA NVSL Diagnostic Production Guide No. R–72/73/74.
 
4.   De Waal D.T. & Potgieter F.T. (1987). The transstadial transmission of Babesia caballi by Rhipicephalus evertsi evertsi. Onderstepoort J. Vet. Res., 54, 655–656.
 
5.   Donnelly J., Joyner L.P., Graham-Jones O. & Ellis C.P. (1980). A comparison of the complement fixation and immunofluorescent antibody titres in a survey of the prevalence of Babesia equi and Babesia caballi in horses in the Sultanate of Oman. Trop. Anim. Health Prod., 12, 50–60.
 
6.   Frerichs W.M., Holbrook A.A. & Johnson A.J. (1969). Equine piroplasmosis: Complement-fixation titres of horses infected with Babesia caballi. Aust. J. Vet. Res., 30, 697–702.
 
7.   Friedhoff K. (1970). Studies on the fine structure of Babesia bigemina, B. divergens and B. ovis. Proceedings of the Second International Congress of Parasitology, J. Parasitol., 56, Section 2 (1), 110.
 
8.   Herr S., Huchzermeyer H.F.K.A., Te Brugge L.A., Williamson C.C., Roos J.A. & Schiele G.J. (1985). The use of a single complement fixation test technique in bovine brucellosis, Johne’s disease, dourine, equine piroplasmosis and Q fever serology. Onderstepoort J. Vet. Res., 52, 279–282.
 
9.   Hirata H., Xhau X., Yokoyama N., Yousifumi N., Kozo F., Suzuki N. & Igarashi I. (2003). Identification of a specific antigenic region of the P82 protein of Babesia equi and its potential use in serodiagnosis. J. Clin. Microbiol., 41, 547–551.
 
10.   Holbrook A.A., Johnson A.J. & Madden B.S. (1968). Equine piroplasmosis: Intraerythrocytic development of Babesia caballi (Nuttall) and Babesia equi (Laveran). Am. J. Vet. Res., 29, 297–303.
 
11.   Holman P.J., Chieves L., Frerichs W.M., Olson D. & Wagner G.G., 1994. Babesia equi erythrocytic stage continuously cultured in an enriched medium. J. Parasitol., 80, 232–236.
 
12.   Holman P.J., Frerichs W.M., Chieves L. & Wagner G.G. 1993. Culture confirmation of the carrier status of Babesia caballi-infected horses. J. Clin. Microbiol., 31, 698–701.
 
13.   Ikadai H., Nagai A., Xuan X., Igarashi I., Kamino T., Tsuji N., Oyamada T., Suzuki N. & Fujisaki K. (2002). Seroepidemiologic studies on Babesia caballi and Babesia equi infection in Japan. J. Vet. Med. Sci., 64, 325–328.
 
14.   Kappmeyer L.S., Perryman L.E., Hines S.A., Baszler T.V., Katz J.B., Hennager S.G. & Knowles D.P. (1999). Detection of equine antibodies to Babesia caballi recombinant B. caballi rhoptry-associated protein 1 in a competitive-inhibition enzyme-linked immunosorbent assay. J. Clin. Microbiol., 37, 2285–2290.
 
15.   Kappmeyer L.S., Perryman L.E. & Knowles D.P (1993). A Babesia equi gene encodes a surface protein with homology to Theileria species. Mol. Biochem. Parasitol., 62, 121–124.
 
16.   Katz J., Dewald R. & Nicholson J. (2000). Procedurally similar competitive immunoassay systems for the serodiagnosis of Babesia equi, Babesia caballi, Trypanosoma equiperdum and Burkholderia mallei infection in horses. J. Vet. Diagn. Invest., 12, 46–50.
 
17.   Knowles D.P. Kappmeyer, L.S. & Perryman L.E. (1997). Genetic and biochemical analysis of erythrocyte-stage surface antigens belonging to a family of highly conserved proteins of Babesia equi and Theileria species. Mol. Biochem. Parasitol., 90, 69–79.
 
18.   Knowles D.P., Kappmeyer, L.S., Stiller D., Hennager S.G. & Perryman L.E. (1992). Antibody to a recombinant merozoite protein epitope identifies horses infected with Babesia equi. J. Clin. Microbiol., 30, 3122–3126.
 
19.   Knowles D.P., Perryman L.E. & Kappmeyer L.S. (1991). Detection of equine antibody to Babesia equi merozoite proteins by a monoclonal antibody-based competitive inhibition enzyme-linked immunosorbent assay. J. Clin. Microbiol., 29, 2056–2058.
 
20.   Levine N.D. (1985). Veterinary protozoology. Iowa State University Press, Ames, Iowa, USA.
 
21.   Littlejohn A. (1963). Babesiosis. In: Equine Medicine and Surgery, Bone J.F., Catcott E.J., Gabel A.A., Johnson L.E. & Riley W.F., eds. American Veterinary Publications, California, USA, 211–220.
 
22.   Madden P.A. & Holbrook A.A. (1968). Equine piroplasmosis: Indirect fluorescent antibody test for Babesia caballi. Am. J. Vet. Res., 29, 117–123.
 
23.   Mahoney D.F. & Saal J.R. (1961). Bovine babesiosis: Thick blood films for the detection of parasitaemia. Aust. Vet. J., 37, 44–47.
 
24.   McGuire T.C., van Hoosier G.L. Jr & Henson J.B. (1971). The complement-fixation reaction in equine infectious anemia: demonstration of inhibition by IgG (T). J. Immunol., 107, 1738–1744.
 
25.   Mehlhorn H. & Schein E. (1984). The piroplasms: Life cycle and sexual stages. Adv. Parasitol., 23, 37–103.
 
26.   Melhorn h. & Schein E. (1998). Redescription of Babesia equi Laveran, 1901 as Theileria equi. Parasitol Res., 84, 467-475.
 
27.   Morzaria S.P., Brocklesby D.W. & Harradine D.L. (1977). Evaluation of the indirect fluorescent antibody test for Babesia major and Theileria mutans in Britain. Vet. Rec., 100, 484–487.
 
28.   Neitz W.O. (1956). Classification, transmission and biology of piroplasms of domestic animals. Ann. N.Y. Acad. Sci., 64, 56–111.
 
29.   Potgieter F.T. & Els H.J. (1977). The fine structure of intra-erythrocytic stages of Babesia bigemina. Onderstepoort J. Vet. Res., 44, 157–168.
 
30.   Rhalem A., Sahibi H., Lasri S., Johnson W.C., Kappmeyer L.S., Hamidouch A., Knowles D.P. & Goff W.L. (2001). Validation of a competitive enzyme-linked immunosorbent assay for diagnosing Babesia equi infections of Moroccan origin and its use in determining the seroprevalence of B. equi in Morocco. J. Vet. Diagn. Invest., 13, 249–251.
 
31.   Sahagun-Ruiz A., Wagnela S.D., Holman P.J., Chieves L.P. & Wagner G.G. (1997). Biotin-labeled DNA probe in a PCR-based assay increases detection sensitivity for the equine hemoparasite Babesia caballi. Vet. Parasitol., 73, 53–63.
 
32.   Schein E., Rehbein G., Voigt W.P. & Zweygarth E. (1981). Babesia equi (Leveran, 1901). Development in horses and in lymphocyte culture. Tropenmed Parasitol., 32, 223–227.
 
33.   Shkap V., Cohen I., Leibovitz B., Savitsky, Pipano E., Avni G., Shofer S., Giger U., Kappmeyer L. & Knowles D. (1998). Seroprevalence of Babesia equi among horses in Israel using competitive inhibition ELISA and IFA assays. Vet. Parasitol., 76, 251–259.
 
34.   Shute P.G. (1966). The staining of malaria parasites. Trans. R. Soc. Trop. Med. Hyg., 60, 412–416.
 
35.   Stiller D. & Frerich S.W.M. (1979). Experimental transmission of Babesia caballi to equids by different stages of the tropical horse tick, Anocentor nitens. Recent Adv. Acarol., 2, 262–268.
 
36.   Stiller D., Frerichs W.M., Leatch G. & Kuttler K.L. (1980). Transmission of equine babesiosis and bovine anaplasmosis by Dermacentor albipictus (Packard) (Acari: Ixodidae). J. N.Y. Ent. Soc., 88, 75–76.
 
37.   Taylor W.M., Bryant J.E. Anderson J.B. & Willers K.H. (1969). Equine piroplasmosis in the United States – A review. J. Am. Vet. Med. Assoc., 155, 915–919.
 
38.   Tenter A.M. & Freidhoff K.T. (1986). Serodiagnosis of experimental and natural Babesia equi and B. caballi infections. Vet. Parasitol., 20, 49–61.
 
39.   United States Department of Agriculture (USDA) Animal and Plant Health Inspection Service, Veterinary Services (1997). Complement fixation test for the detection of antibodies to Babesia caballi and Babesia equi – microtitration test. USDA, National Veterinary Services Laboratories, Ames, Iowa, USA.
 
40.   United States Department of Agriculture (USDA) Animal and Plant Health Inspection Service, Veterinary Services (2003). Competitive ELISA for Serodiagnosis of Equine Piroplasmosis (Babesia equi and Babesia caballi), and Production of Recombinant Babesia equi and Babesia caballi cELISA Antigens. USDA, National Veterinary Services Laboratories, Ames, Iowa, USA.
 
41.   Xuan X., Larsen A., Idadai H., Tnanka T., Igarashi I., Nagasawa H., Fujisaki K., Toyoda Y., Suzuki N. & Mikami T. (2001). Expression of Babesia equi merozoite antigen 1 in insect cells by recombinant baculovirus and evaluation of its diagnostic potential in an enzyme-linked immunosorbant assay. J. Clin. Microbiol., 39, 705–709.
 
42.   Zweygarth E., Just M.C. & De Waal D.T. (1995). Continuous in vitro cultivation of erythrocytic stages of Babesia equi. Parasitol. Res., 81, 355-358.
 
43.   Zweygarth E., Just M.C. & De Waal D.T. (1997). In vitro cultivation of Babesia equi: detection of carrier animals and isolation of parasites. Onderstepoort J. Vet Res., 64, 51–56.
 

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