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T. evansi tests in Kenya
Veterinary Parasitology 124 (2004) 187–199 www.elsevier.com/locate/vetpar
Detection of Trypanosoma evansi in camels using PCR and CATT/T. evansi tests in Kenya Z.K. Njirua,*, C.C. Constantinea, J.M. Ndung’uc, I. Robertsonb, S. Okayec, R.C.A. Thompsona, S.A. Reida a
Western Australian Biomedical Research Institute, Division of Health Sciences, School of Veterinary and Biomedical Sciences, Murdoch University, South Street, Murdoch, WA 6150, Australia b Division of Health Sciences, School of Veterinary and Biomedical Sciences, Murdoch University, South Street, Murdoch, WA 6150, Australia c Kenya Trypanosomiasis Research Institute (KETRI), P.O. box 362, Kikuyu, Kenya Received 27 February 2004; received in revised form 18 June 2004; accepted 26 June 2004
Abstract Camel trypanosomosis (Surra) causes high morbidity and is an impediment to the camel husbandry in Kenya. The lack of a sensitive diagnostic test has hindered the collection of accurate epidemiological data and institution of control programmes. A cross-sectional study was conducted in three districts of Kenya to estimate the prevalence of Trypanosoma evansi (T. evansi) and to compare four diagnostic tests: polymerase chain reaction (PCR), card agglutination test (CATT/ T. evansi), microhaematocrit centrifugation technique (MHCT) and mouse inoculation (MI). A total of 549 camels were randomly sampled. The overall prevalence of Surra was 5.3% using MHCT, 26.6% using PCR and 45.9% using CATT/T.evansi. There was a significant difference (P < 0.001) between PCR and CATT/T.evansi test, MHCT and MI in detection of T. evansi. The prevalence of T. evansi was 39.8% in Samburu, 24.7% in Nanyuki and 14.4% in Isiolo districts using PCR. A male camel was 2.6 times more likely to be infected with T. evansi compared to a female camel (OR = 3.0% CI: 1.6, 4.1), while an adult camel was 2.2 times more likely to be infected compared to non-adults (OR = 2.2; 95% CI: 1.2, 5.0). There was a poor association between the presence of the published clinical signs and seropositivity (k = 0.12), PCR (k = 0.11) and MHCT (k = 0.05). However, there was a higher agreement between farmers’ classification of disease with the PCR test (k = 0.5, n = 61). The mean PCV varied with age, presence of infection, locality and gender, with the lowest mean PCV being recorded in MHCT-positive * Corresponding author. Tel.: +61 08 9360 2690; fax: +61 08 9310 4144. E-mail address: [email protected] (Z.K. Njiru). 0304-4017/$ – see front matter # 2004 Elsevier B.V. All rights reserved. doi:10.1016/j.vetpar.2004.06.029
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animals (20.97 0.5) and from infected calves (19.5 1.2). This study shows that PCR was more sensitive in detecting T. evansi than other tests used. Further, the prevalence of T. evansi in the camel herds sampled is higher than that previously reported in Kenya, and that the judgment by camel keepers may be a reliable ‘‘pen-side’’ diagnostic test for Surra. Considering the low sensitivity of parasitological techniques in detection of chronic T. evansi infection and high cost of PCR, development of a sensitive pen side diagnostic test, with a low cost is still a priority. # 2004 Elsevier B.V. All rights reserved. Keywords: Trypanosoma evansi; Camel; PCR; CATT/T. evansi; Trypanosomosis
1. Introduction Trypanosma evansi (T. evansi), the cause of cameline trypanosomosis (Surra), is the most widespread pathogenic trypanosome in the world. It is mechanically transmitted by haemotophagus biting flies and therefore distributed widely outside the tsetse belts. The disease causes significant morbidity and mortality in camels in arid and semi arid regions in Kenya, which have a population of over 1 million camels and support over 25% of human population and over half of livestock production (Anonymous, 1997). Trypanosomosis in camels occurs both in chronic and acute forms (Wilson et al., 1983). The chronic form is most common and may present an association with secondary infections due to immunosuppression caused by T. evansi infection, which complicates clinical diagnosis. Published clinical signs (emaciation, fever, anaemia, lacramation, corneal opacity, diarrhoea and oedema of the dependent parts) (Chaudhary and Iqbal, 2000) are insufficient for diagnosis, while detection of parasites in the blood is difficult because parasitaemia is intermittent (Mahmound and Gray, 1980). Consequently, there is a need for alternative more sensitive diagnostic techniques. Serological tests have been developed and evaluated for diagnosis of trypanosomosis in camels. The antibody-detecting tests such as the CATT/T. evansi (Bajyana-Songa and Hamers, 1988) and enzyme-linked immunosorbent assay (Ab-ELISA) (Atarhouch et al., 2003) have been shown to be reliable while the antigen based Suratex1 (Nantulya, 1994) and Ag-ELISA (Olaho-Mukani et al., 1993) are seldom used. CATT/T. evansi, which is a commercially produced kit, is widely used in the field. However, it is unable to distinguish past (treated) from current infections because it detects antibodies that may persist for a long time following treatment. The polymerase chain reaction (PCR) is highly sensitive and specific and has been widely used in detection of trypanosomes. Primers targeting subgroup Trypanozoon (Moser et al., 1989; Masiga, 1994; Wuyts et al., 1994) and putatively T. evansi-specific (Artama et al., 1991; Masiga, 1994; Ventura et al., 2002) have been developed. The PCR has been used in surveys to determine the prevalence of T. evansi in buffaloes in Vietnam (Holland et al., 2001) and T. equiperdum in Mongolian horses (Clausen et al., 2003). However, PCR has not been systematically used for the detection of T. evansi in camels in Kenya (Masiga and Nyang’ao, 2001). In this study, we determined the prevalence of T. evansi in three camel-keeping regions of Kenya and compared the diagnostic utility of PCR, CATT/T. evansi, MHCT and mouse inoculation for the detection of T. evansi infections in camels.
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2. Materials and methods 2.1. Sampling sites A survey was conducted in three districts Samburu (Wamba and Ngurunit), Isiolo (Tractor, Eremet, Isiolo town and Ngare Ndare) and Nanyuki (Nanyuki town and Eldana) districts in August, 2003. The districts lie between longitudes 3684W and 3684 27E and between latitudes 0817S and 0845N. Rearing of camels is the major activity in these regions although small ruminants (sheep and goats) are also kept. The camels are moved by owners within a defined area depending on the availability of pasture. Rainfall is bimodal in pattern with long rains occurring between March and May and short rains from October to December. The annual rainfall and temperatures range from 500 to 900 mm and 20 to 37 8C, respectively. In Isiolo and Nanyuki, vegetation is predominantly savannah with Acacia species; communities of wooded grassland and bush land thickets are found along seasonal watercourses. In Samburu, most of the regions containing camels are grasslands with scattered Acacia species and short shrubs, with stretches of bare land. Numerous biting flies (Haematobia minuta) are prevalent in Nanyuki and Isiolo with few recorded occurrences of Tabanus taeniola, Hippobosca camelina and H. variegata (Njiru et al., 2002). 2.2. Study design and sampling A cross-sectional sampling procedure was used during this study. A total of 549 camels were sampled, 153 in Isiolo, 161 in Samburu and 235 in Nanyuki district, respectively. The sampling areas were chosen because information from respective district veterinary offices suggested the presence of Surra. Before the field trip, information was passed to the camel owners through their community leaders. On the day of sampling, animals were selected using a random number generator, marked using permanent spray, then information on age, gender, previous trypanocide treatment, history of abortions and weight estimations was recorded. Each animal was clinically examined, and 5 ml of blood was collected from the jugular vein. Two millilitres of blood was placed into a tube containing heparin for PCV, parasite examination and mouse inoculation, and 3 ml blood was placed in plain tubes and left to clot for serum collection. The serum was removed after 12 h and stored in liquid nitrogen. Collected heparinised blood samples were stored in cool boxes and transported to a temporary laboratory within the immediate vicinity. Approximately, 0.5 ml of blood was transferred into cryovials, mixed with phosphate saline glucose (PSG) and 10% glycerol, and stored in liquid nitrogen for subsequent extraction of genomic DNA. Animals were grouped into age groups; calves <12 months, immature 12 and 36 months and adults >36 months for purposes of analysis. 2.3. Parasitological examination and mouse inoculation Two capillary tubes were prepared from each heparinised blood sample and centrifuged in a microhaematocrit centrifuge for 5 min at 12,000 g. The packed cell volume (PCV) was
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determined, and both capillary tubes were examined for the presence of motile trypanosomes using the microhaematocrit centrifugation technique (MHCT) (Woo, 1970) and the buffy-coat technique (Murray et al., 1977). Four hundred microlitres of blood from animals with a PCVof 26% and/or showing clinical signs suggestive of Surra (Chaudhary and Iqbal, 2000) were inoculated into Swiss-bred mice, which were screened for infection with T. evansi by wet smear examination of tail-tip blood every day for 60 days. Any camel shown to be infected with T. evansi using MHCT and those animals showing clinical signs suggestive of Surra (emaciation, PCV 20% or a recent history of abortion) were treated with quinapyramine sulphate/chloride (Triquin1) intramuscular at 5 mg/kg body weight. 2.4. Serological tests Sera were tested for the presence of anti-T. evansi antibodies using the card agglutination test for T. evansi (CATT/T. evansi) (Institute of Tropical Medicine, Antwerp, Belgium). Approximately, 45 ml of the antigen was transferred into the test card and mixed with an equal amount of the test sera diluted at 1/4 with PBS pH 7.2 as per manufacturer’s instructions. The card was agitated for 5 min, and the reaction was checked in the clear light. Positive reaction was confirmed on recording agglutinations (blue granules) (Bajyana-Songa and Hamers, 1988). 2.5. Template preparation and DNA amplification Genomic DNA was extracted from samples of camel blood using commercially available QIAamp DNA mini kit (Qiagen, 2003), and the purified DNA template was stored at 20 8C for later use. PCR was carried out in 25 ml reaction mixtures containing 10 reaction buffer (Fischer Biotech), 1.5 mM MgCl2, 200 mM of each of the four deoxynucleoside triphosphates (dNTPs) (Promega), primers at 1 mM and 0.5 U of Taq DNA polymerase (Fischer Biotech). The cycles included an initial step at 94 8C for 4 min, followed by 30 cycles of denaturing at 948 C for 30 s, annealing at 608 C for 30 s and extension at 728 C for 30 s. PCR Elongation was continued at 728 C for 5 min. The amount of DNA template added per reaction was 3 ml. A positive (reference T. evansi DNA, 25– 50 ng/ml) and a negative control (without DNA) were included for each PCR reaction. Amplification products were resolved in 1.5% molecular grade agarose gel (Fischer Biotech) stained with ethidium bromide (5 mg/ml). 2.6. Primers Primers targeting a repetitive region specific for Trypanozoon subgenus (Wuyts et al., 1994) pMURTec.F, 50 -TGCAGACGACCTGACGCTACT; pMURTec.R, 50 -CTCCTAGAAGCTTCGGTGTCCT and T. evansi minicircle specific (Masiga, 1994) EVA1, 50 ACATATCAACAACGACAAAG, EVA2 50 -CCCTAGTATCTCCAATGAAT were used. Since the minicircle primers are less sensitive than the Wuyts primers, they were used only on those samples that gave strong reactions using the latter primers and/or on samples shown to be MHCT positive.
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2.7. Weight measurements The live body weight of camels (kg) was estimated by measuring (in meters) shoulder height, thoracic girth and abdominal girth and multiplying these by 50 (Schwartz et al., 1983). Only camels from Isiolo and Samburu districts had their weights estimated. 2.8. Statistical analysis Statistical analysis was performed using SPSS v11.5. The level of agreement between diagnostic tests was determined using (Kappa statistics (k) and the influence on prevalence of various factors, such as gender, age, locality, abortion and presence of clinical signs were determined using chi-square test and odds ratios and their 95% confidence intervals. Analyses of variance (ANOVA) with Bonferoni mean-separation tests and t-tests were used to compare mean PCV values and weights in groups of camels.
3. Results 3.1. Diagnostic tests analysis A total of 549 camels were screened for infection with T. evansi using parasitological methods (MHCT and BCT), PCR and the CATT/T. evansi. Overall prevalence was 5.3% (95% CI: 3.4, 7.2) with the MHCT, 26.2% (95% CI: 22.5, 29.9) with the PCR and 45.9% (95% CI: 41.7, 50.1) with CATT/T. evansi. The prevalence of infection in each area sampled is presented in Table 1. There was a significant difference in the prevalence estimate determined using PCR and CATT/T. evansi (P < 0.001) while prevalence estimates using MHCT was significantly lower (P < 0.001) than those of PCR and CATT/T. evansi. There was fair agreement between PCR and MHCT (k = 0.3; 95% CI: 0.22, 0.38) and between PCR and CATT/T. evansi (k = 0.2; 95% CI: 0.13, 0.27). Camel owners identified the clinical signs of Surra as rough coat, drop in milk yield, abortion and by a characteristic urine odour. There was a moderate agreement between diagnosis by camel owners and PCR (k = 0.52; 95% CI: 0.3, 0.7, n = 61). Using PCR results, a camel reared in Samburu district was four times (OR = 3.9; 95% CI: 2.3, 6.8) and two times (OR = 2.0; 95% CI: 1.3, 3.1) more likely to be infected than a camel reared in either Isiolo or Nanyuki district, respectively. Table 1 T. evansi and seroprevalence in Samburu, Isiolo and Nanyuki district District
No. of animals
MHCT
PCR
Positive (%)
95% CI
Positive (%)
CATT test 95% CI
Positive (%)
95% CI
Samburu Isiolo Nanyuki
161 153 235
20 (12.4) 2 (1.3) 7 (3.0)
7.3, 17.5 0, 3.1 0.8, 5.2
64 (39.8) 22 (14.4) 58 (24.7)
32.2, 47.4 8.8, 20.0 19.2, 30.2
68 (42.2) 53 (34.6) 131 (55.7)
34.6, 49.8 27.1, 42.1 49.3, 62.1
Total
549
29 (5.3)
3.4, 7.2
144 (26.2)
22.5, 29.9
252 (45.9)
41.7, 50.1
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Fig. 1. Prevalence of T. evansi within age structure and sex.
3.2. Age and gender The prevalence of infection estimated using PCR was significantly higher in adult camels as compared to immature and calves (P < 0.05 and P < 0.01, respectively). The prevalence of T. evansi infection estimated using PCR and MHCT was also higher in male compared to female camels (x2 = 17.0, df = 1, P < 0.001 and (x2 = 9.1, df = 1, P < 0.01, respectively) (Fig. 1). Conversely, the seroprevalence was higher in adult camels compared to immature camels and calves (P < 0.001). Adult camels were 2.2 times more likely to be CATT/T. evansi-positive compared to non adult camel (OR = 2.2; 95% CI: 1.2, 5.0). There was no significant difference in the seroprevalence between the genders while male camels were 2.6 times more likely to be T. evansi-positive using PCR test compared to female camels (OR = 2.6; 95% CI: 1.6, 4.1). 3.3. Packed cell volume The lowest PCV means were recorded for MHCT-positive animals (20.9 0.5), infected calves (19.5 1.4) and animals from Samburu district (23.1 0.4). Overall the mean male camels had significantly lower PCVs compared to female camels (P < 0.05) and immature camels had significantly lower PCVs compared to adult camels (P < 0.001) (Table 2). The PCV of camels shown to have circulating T. evansi by PCR or MHCT was significantly lower than aparasitaemic animals (P < 0.001). There was no significant difference in mean PCV of seropositive and seronegative camels. 3.4. Sensitivity Of 29 MHCT-positive samples collected in this study, 28 (96.6%) were PCR-positive and 19 (65.5%) CATT/T. evansi-positive. Using both tests, 19 samples were positive and only one was negative on both tests. In earlier and separately collected samples, PCR
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Table 2 Packed cell volume for camels in Samburu and Isiolo district Category
n
Age Adult
Nanyuki
22.98 0.52 23.69 0.63
P = 0.4
Infected Non infected
103 351
24.28 0.43 25.51 0.18
P < 0.01
23.48 0.43a
Infected Non infected
454
b
25.23 0.17
n
397
25.55 0.18c
Infected Non infected
119 278
24.59 0.36 25.97 0.20
P < 0.001
84
24.95 0.35c
Infected Non infected
15 69
21.47 0.60 25.71 0.34
P < 0.001
Infected Non infected
10 58
19.50 1.39 21.36 0.41
P = 0.12
Infected Non infected
64 97
22.48 0.62 23.53 0.43
P = 0.16
Infected Non infected
22 131
26.77 0.75 25.33 0.29
P = 0.06
Infected Non infected
58 177
24.40 0.28 26.17 0.23
P < 0.001
Calves
Isiolo
41 54
95
Immature
Districts Samburu
t-test
Status
Gender Male Female
PCV S.E.M.
Mean PCV
68
d
21.09 0.4
161
23.11 0.36b
153
a
235
25.54 0.28
a
25.73 0.19
Superscripts that are different are significantly different within each group for mean PCV (mean PCV for age and district was analyzed with ANOVA, P < 0.05, while gender was done with t-test, P < 0.05).
sensitivity was 70/72 (97.2%) and 48/72 (66.7%) for CATT/T. evansi test. Mouse inoculation sensitivity was done for 22 positive samples from Samburu and Isiolo of which 19 (86.4%) were positive. There was a poor association but still significant difference between MI and PCR in detecting T. evansi infections in 175 aparasitaemic samples (k = 0.19; P < 0.001) as shown in Table 3. 3.5. Weights The estimated mean weights for adults, immature and calves were 484 9.1, 264 13.3 and 203 16.4 kg, respectively. Significant difference weights were only recorded Table 3 Relationship between PCR and mouse inoculation in detection of T. evansi infections in aparasitaemic camels with PCV of 26% and showing clinical signs of Surra Mouse inoculation
PCR Positive
Negative
Total
Positive Negative
8 42
2 123
10 165
Total
50
125
175
Fischer’s exact test, P < 0.001, k = 0.19, 95% CI: 0.06, 0.3.
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between seropositive (501 8.75) and seronegative adults (462 17.3) (P < 0.05) and between estimated weights of camels from Isiolo and Samburu (433.75 14.30 and (381.36 12.8; P < 0.001).
4. Discussion In this study, infection with T. evansi infection was demonstrated in the three districts sampled. The prevalence estimates from this study are higher than those of previous studies (Ngaira et al., 2003). There is no obvious reason why the prevalence of T. evansi in camels in Samburu district was higher than in other districts sampled. However, it is worth noting that the areas visited in Samburu district were difficult to access (poor roads especially in the rainy seasons) as compared to Nanyuki and Isiolo, which have better communication and transport network. Therefore, it is possible that some of the difference in prevalence may be because camel keepers in the latter districts have better access to trypanocidal drugs and veterinary services. The results of this study indicate that PCR is the most accurate test for the detection of T. evansi parasitaemia in camels. PCR has been used successfully in detecting infection with T. evansi in buffaloes (Omanwar et al., 1999; Holland et al., 2001), horses (Clausen et al., 2003) and in camels (Masiga and Nyang’ao, 2001). It is interesting to note that the prevalence estimate determined using PCR varied in the different districts visited with the highest prevalence in Samburu district. This may indicate differences in the epidemiology of infection because seroprevalence did not vary significantly with district, which might indicate a higher level of active infections in Samburu compared to other districts. These differences may also be due to genetic differences in T. evansi isolated from each area. Animals in Samburu district also weighed less and showed poor body conditions due to factors, which were associated with higher disease burden and poor foliage. The CATT/T.evansi test is a well-validated field assay for the detection of T. evansi in camels (Dia et al., 1997; Ngaira et al., 2003) buffalo (Davison et al., 1999) and cattle (Reid and Copeman, 2003)). It has been the test of choice for the field surveys, although being an antibody-detecting assay, it has inherent shortcomings of not being able to differentiate between past and present infections. The sensitivity of CATT/T. evansi in this study was 65.5%, which compared well with 68.6% reported in an earlier study by Ngaira et al. (2003) in Isiolo district. So far, there is no comprehensive data on the use of PCR for detecting infection in camels and/or comparison of CATT/T. evansi test with PCR. Such data are necessary in the face of camel husbandry strengthening in Kenya. A comparison of the PCR with CATT/T. evansi showed a significant difference (P < 0.001) and only a fair association (k = 0.3), implying that the two tests were categorizing infections independently. The CATT/T. evansi test failed to detect 51/144 (35.4%) of PCR positive samples and 10/29 (3.5%) of MHCT positive samples. Not only the phenomenon of parasitological positive samples but CATT/T. evansi negative test were also observed by Ngaira et al. (2003), Elsaid et al. (1998) and Davison et al. (1999). The 51 PCR positive cases missed by CATT/T. evansi may indicate early infections where animals have not yet formed detectable antibody levels or isolates that lack Ro Tat 1.2 gene and/or do not express Ro Tat 1.2 VSG. If treatment had been instigated based on the results of CATT/T.
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evansi, 252 animals would have been treated as opposed to 144 animals if the results had been based on the PCR. Assuming no false positive with PCR, this indicates 108 (43%) unnecessary treatments. This means that in an area where treatment is frequent, CATT/T. evansi test needs to be used with second confirmatory test. In our case, PCR, although expensive, it was the most appropriate in detecting both chronic and current infections. The CATT/T. evansi is based on variable surface antigen (VSG) designated as RoTat 1.2, which is thought to be expressed in all T. evansi isolates (Verloo et al., 2001). However, recently Ngaira et al. (2004) have shown isolates from Kenya that lack both Ro Tat 1.2 gene and their associated VSG and another group of isolates with Ro Tat 1.2 gene but absence of expressed VSG. Earlier studies showed variable expression of VSG genes in the closely related species T. b. gambiense (Dukes et al., 1992). If the recently observed variations are widespread in Kenya camel-rearing regions, then it may explain the low sensitivity of the CATT/T. evansi observed. Moreover, isolates of T. evansi from Kenya have also been shown to differ genetically (Gibson et al., 1983) and the existence of T.evansi type B suggests some genetic variation occurring in the Kenyan northern frontier (Borst et al., 1987). Further ongoing studies of isolates from the same study areas reveal regional genetic differences in T. evansi isolates (Njiru, unpublished results). The possible occurrence of infections with other Trypanozoon parasites was ruled out by testing most of the PCRpositive isolates (n = 88) with T. evansi-specific minicircle primers (Masiga, 1994). Ongoing work will confirm whether our 10 MHCT-positive but CATT/T. evansi-negative isolates express the RoTat 1.2 gene. However, one way to improve CATT/T. evansi sensitivity would be to add other not additional to other antigens in the test. The PCR test had a sensitivity of 97% in this study and 97.2% in a separate study using Trypanozoon spp primers (Wuyts et al., 1994). The one missed case in this study could either be due to degraded DNA and/or loss of DNA during extraction. The strength of PCR was further shown in detection of infection in aparasitaemic animals showing clinical symptoms (Table 3). Analysis of animals with PCV of 26% and some with clinical signs of Surra (lachrimation, diarrhoea, emaciation, enlargement of lymph nodes and recent aborted females) showed T. evansi prevalence of 50/175 (28.6%) and 10/175 (5.7%) with PCR and MI. When all clinical symptoms were combined together, a fair association between clinical signs and seroprelevalence (k = 0.3; 95% CI: 0.22, 0.38) and low association with PCR (k = 0.1; 95% CI: 0.05, 0.19) were recorded. These results are suggestive that Surra may not be the only cause of the observed clinical signs. This argument is reflected in the low numbers of infected animals showing each clinical symptom, e.g. 1/7 (diarrhoea), 5/15 (lacrimation) and 10/26 (emaciated) were positive either with MHCT or PCR. Documented clinical signs of Surra are general for most of the susceptible animals. In the absence of other diagnostic techniques, this leaves treatment of camels based on personal opinion/experience. An interesting strong association (k = 0.5; 95% CI: 0.32, 0.72, n = 61) was shown in this study between signs recognized by camel keepers (urine with a characteristic smell, drying up of lactating camels, reluctance to move, rough coat and abortions) and infection in animals (MHCT and PCR). The characteristic smell of urine is due to ketones produced after breakdown of amino acids by parasite in infected animals (Hunter, 1986). Though the sample size was small, it appears that the camel owners have dependable diagnostic knowledge of Surra having lived with these animals for generations. Use of parasitological technique in T. evansi detection still
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remains wanting because of the chronic nature of T. evansi in the field. Both BCT and MHCT will not be done away soon considering the cost of other diagnostic techniques. However, these techniques may be made more useful if clinical sign guidelines are established with disease prevalence and are used side by side. In this study, 36 camels were treated in Samburu and Isiolo district based on results of PCV, MHCT and the presence of clinical signs in the field. However, the PCR results indicated that 86 of the sampled animals were infected in these regions. If the PCR indicated active infection, then 58.1% of infected animals could not be detected through the most commonly used method. If we included CATT/T. evansi test results, the number treated could have gone up to 52 still leaving 39.2% animals as untreated but also increasing the number of treated ‘‘false’’ negative. These undetected cases would eventually act as a source for future infection in the herds. MI is a frequently used technique in the field to detect latent infections. In our study, MI showed a sensitivity of 86.4% of parasitaemic cases, however it picked only 16% (8/50) of PCR-positive aparasitaemic cases. The low prevalence identified with the MI may be due to trypanosomes being lost through the mice immune systems. Sensitivity of MI may be improved by use of irradiated mice though expensive. The observation that seropositivity increases with age is in agreement with previous studies (Dia et al., 1997; Atarhouch et al., 2003; Gutierrez et al., 2000) and is probably related to the chronic nature of infection and possibility that antibodies persist after a camel is treated. However, it was also noted that the camel owners prefer to graze calves within the vicinity of human dwellings as opposed to older camels which are grazed and watered in the open and hence have greater potential for contact with vectors. There is insufficient data to explain the observed differences in the prevalence of infections between males and female camels. One possibility is that hormonal differences may influence vector behaviour and increase the likelihood of infection in male compared to female camels. Twenty-six animals were reported to have aborted between 1 and 6 months period prior to this study. When camel keepers routinely treat these animals with trypanocides, immediately an abortion occurs. Of these animals, 14/26 (53.8%) were still positive for T. evansi with PCR (including 3 MHCT-positive cases) and the same number were seropositive. Ten animals with active infection also were seropositive and eight animals were negative with both PCR and CATT/T.evansi test. Active infection in these already treated camels indicates either re-infection or treatment failure. Misuses of drugs through under dosing (often self-administrated by the camel keepers without correct weight adjustment or using a single vial to treat several animals) and fake drugs (Ngaira et al., 2002) may have consequential effects on T. evansi prevalence and the development of resistance. There was a significant relationship between infection and low PCV. Previous authors have tried to determine a cut-off value for PCVs that can be used for diagnostic purposes. For example, <23% was proposed by Ngaira et al. (2002) in Kenya, 20% by Chartier et al. (1986), in Mauritania and 18% by Diall et al. (1993) in Mali. Though anaemia could result from T. evansi infection, other factors such as infection with Haemonchus spp and the effects of drought could cause similar effects. In this study, we found that camels shown to be PCR and MHCT-positive had significantly lower PCVs compared to uninfected camels.
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We further found significant differences in PCV by locality, gender and age (Table 2). Therefore, it is difficult to use PCV as the sole indicator of infection with T. evansi. Trypanosomosis continues to pose a great risk to camel keeping in Kenya. Low sensitivity of diagnostic techniques is a major factor impeding the control of this disease. This study has shown that Surra is prevalent in all three districts sampled at significantly different levels. The PCR test showed the best sensitivity compared with CATT/T. evansi using MHCT positive samples as the gold standard. Noting the chronic nature of the disease and apparent low sensitivity of CATT/T. evansi in Kenya, the use of PCR is recommended for inclusion in survey and control programmes. In instances where treatment has to be carried out immediately in the field, consideration of the farmers’ diagnosis with respect to documented signs by veterinarian is necessary. However, we recommend a further comprehensive study on farmers’ diagnostic knowledge versus disease prevalence in the camel herds. In cases where low PCV as a measure of infection is used, considerations of other factors like age and genders are necessary.
Acknowledgements The authors gratefully acknowledge the financial support from the International Foundation for Science (IFS grant number B3372 to Z.K.Njiru), Kenya government and Murdoch University. Supplemental funds were contributed by Western Australia Biomedical Research Institute (WABRI). We also thank Dr. Thuita, Dr. Chemuliti, Rashid Farah and Veterinary department, Isiolo district for helping with field sampling, Purity Njoka, John Ndichu and Samuel Guya for excellent technical assistance.
References Artama, W.T., Agey, M.W., Danelson, J.E., 1991. DNA comparison of Trypanosoma evansi (Indonesia) and Trypanosoma brucei spp. Parasitology 104, 67–74. Atarhouch, T., Rami, M., Bendahman, M.N., Dakkak, A., 2003. Camel trypanosomosis in Morocco 1: results of a first epidemiological survey. Vet. Parasitol. 111, 277–286. Bajyana Songa, E., Hamers, R., 1988. A card agglutination test (CATT) for veterinary use based on early VAT RoTat 1.2 of Trypanosoma evansi. Ann. Soc. Belg. Med. Trop. 68, 233–240. Borst, P., Fase-Fowler, F., Gibson, W.C., 1987. Kinetoplast DNA of Trypanosoma evansi. Mol. Biochem. Parasitol. 23, 31–38. Clausen, P.H., Chuluun, S., Sodnomdarjaa, R., Greiner, M., Noeckler, K., Staak, C., Zessin, K.H., Schein, E., 2003. A field study to estimate the prevalence of Trypanosoma equiperdum in Mongolian horses. Vet. Parasitol. 115, 9–18. Chartier, C., Chartier, F., Lepers, J.P., Pesce, J.L., 1986. Preliminary studies of common blood parameters of the Mauritanian dromedary (Camelus dromedaries). Rev. Elev. Med. Vet. Pays. Trop. 39, 395–401. Chaudhary, Z.I., Iqbal, M.J., 2000. Incidence, biochemical and haematological alterations induced by natural trypanosomosis in racing dromedary camels. Acta Trop. 77, 209–213. Davison, H.C., Thrusfield, M.V., Muharsini, S., Husein, A., Partoutomo, S., Rae, P.F., Masake, R., Luckins, A.G., 1999. Evaluation of antigen and antibody detection tests for Trypanosoma evansi infection of buffaloes in Indonesia. Epidemiol. Infect. 123, 149–155. Dia, M.L., Diop, C., Aminetou, M., Jacquiet, P., Thiam, A., 1997. Some factors affecting the prevalence of Trypanosoma evansi in camels in Mauritania. Vet. Parasitol. 72, 111–120.
198
Z.K. Njiru et al. / Veterinary Parasitology 124 (2004) 187–199
Diall, O., Bocoum, Z., Diarra, B., Sanogo, Y., Coulibaly, Z., Waigalo, Y., 1993. Epidemiology of trypanosomosis caused by T. evansi in camels in Mali: results of parasitological and clinical survey. Rev. Elev. Med. Vet. Pays. Trop. 46, 455–461. Dukes, P., Gibson, W.C., Gashumba, J.K., Hudson, K.M., Bromidge, T.J., Kaukus, A., Asonganyi, T., Magnus, E., 1992. Absence of the LiTat 1.3 (CATT antigen) gene in Trypanosoma brucei gambiense stocks from Cameroon. Acta Trop. 51, 123–134. Elsaid, H.M., Nantulya, V.M., Hilali, M., 1998. Diagnosis of Trypanosoma evansi infection among Sudanese camels imported to Egypt using card agglutination test (CATT) and antigen detection latex agglutination test (Suratex). J. Protozool. Res. 8, 194–200. Gibson, W.C., Wilson, A.J., Moloo, S.K., 1983. Characterization of Trypanosoma (Trypanozoon) evansi from camels in Kenya using isoenzyme electrophoresis. Res. Vet. Sci. 34, 114–118. Gutierrez, C., Juste, M.C., Corbera, J.A., Magnus, E., Verloo, D., Montoya, J.A., 2000. Camel Trypanosomosis in the Canary Islands: assessment of seroprevalence and infection rates using the card agglutination test (CATT/ T. evansi) and parasite tests. Vet. Parasitol. 90, 155–159. Holland, W.G., Claes, F., My, L.N., Thanh, N.G., Tam, P.T., Verloo, D., Buscher, P., Goddeeris, B., Vercruysse, J., 2001. A comparative evaluation of parasitological tests and a PCR for Trypanosoma evansi diagnosis in experimentally infected water buffaloes. Vet. Parasitol. 97, 23–33. Hunter, A.G., 1986. Urine odour in a camel suffering from Surra T. evansi infection. Trop. Anim. Health Prod. 18, 146–148. Anonymous, 1997. The Eighth National Development Plan, Government printers, Nairobi, Kenya, pp. 76–77. Masiga, D.K., 1994. The Development and Application of the Polymerase Chain Reaction Methodology for the Identification of African Trypanosomes, Ph.D. Thesis, University of Bristol. Masiga, R.C., Nyang’ao, J.M., 2001. Identification of trypanosome species from camel using polymerase chain reaction and procyclic transformation test. Camel Pract. Res. 8, 17–22. Mahmound, M.M., Gray, A.R., 1980. Trypanosomosis due to T. evansi (Istell, 1885; Balbiani, 1885). A review of recent research. Trop. Anim. Health Prod. 12, 35–47. Moser, D.R., Cook, G.A., Ochs, D.E., Bailey, C.P., McKane, M.R., Donelson, J.E., 1989. Detection of Trypanosoma congolense and T. brucei ssp by DNA amplification using the polymerase chain reaction. Parasitology 99, 57–66. Murray, M., Murray, P.K., McIntyre, I.M., 1977. An improved parasitological technique for the diagnosis of African Trypanosomosis. Trans. R. Soc. Trop. Med. Hyg. 71, 325–326. Nantulya, V.M., 1994. Suratex: a simple latex agglutination antigen test for diagnosis of Trypanosoma evansi infections (Surra). Trop. Med. Parasitol. 45, 9–12. Olaho-Mukani, W., Munyua, W.K, Mutugi, M.W., Njogu, A.R., 1993. Comparison of antigen and antibody selection enzyme immunoassays for the diagnosis of Trypanosoma evansi infections in camels. Vet. Parasitol. 45, 231–240. Ngaira, J.M., Bett, B., Karanja, S.M., 2002. Animal-level factors for Trypanosoma evansi infection in camels in eastern and central parts of Kenya. Onderst. Vet. Res. 69, 263–271. Ngaira, J.M., Bett, B., Karanja, S.M., Njagi, E.N.M., 2003. Evaluation of antigen and antibody rapid detection tests for Trypanosoma evansi infection in camels in Kenya. Vet. Parasitol. 114, 131–141. Ngaira, J.M., Njagi, E.N.M., Ngeranwa, J.J.N., Olembo, N.K., 2004. PCR amplification of Ro Tat 1.2 VSG gene in Trypanosoma evansi isolates in Kenya. Vet. Parasitol. 120, 23–33. Njiru, Z.K., Bett, B., Mapeny-Ole, I.M., Githiori, J.B., Ndung’u, J.M., 2002. Trypanosomosis and helminthosis in camels: comparison of ranch and traditional camel management systems in Kenya. Camel Pract. Res. 9, 67– 71. Omanwar, S., Rao, J.R., Basagoudanavar, S.H., Singh, R.K., Butchaiah, G., 1999. Direct and sensitive detection of Trypanosoma evansi by polymerase chain reaction. Acta Vet. Hung. 47, 351–359. Reid, S.A., Copeman, D.B., 2003. The development and validation of an antibody-ELISA to detect Trypanosoma evansi infection in cattle in Australia and Papua New Guinea. Prev. Vet. Med. 61, 195–208. Schwartz, H.J., Dolan, R.B., Wilson, A.J., 1983. Camel production in Kenya and its constraints. Trop. Anim. Health Prod. 15, 169–178. Verloo, D., Magnus, E., Buscher, P., 2001. General expression of Ro Tat variable antigen type in Trypanosoma evansi isolates from different origin. Vet. Parasitol. 97, 183–189.
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Ventura, R.M., Takeda, G.F., Silva, R.A.M.S., Nunes, V.L.B., Buck, G.A., Teixeira, M.M.G., 2002. Genetic relatedness among Trypanosoma stocks by random amplification of polymorphic DNA and evaluation of a synapomorphic DNA fragment for species-specific diagnosis. Int. J. Parasitol. 2, 53–63. Woo, P.T.R., 1970. The haemotocrit centrifuge technique for the diagnosis of African trypanosomes. Acta Trop. 27, 384–386. Wilson, A.J., Schwartz, H.J., Dolan, R., Olaho-Mukani, W., 1983. A simple classification of different types of trypanosomosis occurring in four camel herds in selected areas of Kenya. Tropenmedizin und Parasitologie 34, 220–224. Wuyts, N., Chokesajjawatee, N., Panyim, S., 1994. A simplified and highly sensitive detection of Trypanosoma evansi by DNA amplification. S.E. Asian J. Trop. Med. Publ. Health 25, 266–271.
Detection of Trypanosoma evansi in camels using PCR and CATT/T. evansi tests in Kenya Z.K. Njirua,*, C.C. Constantinea, J.M. Ndung’uc, I. Robertsonb, S. Okayec, R.C.A. Thompsona, S.A. Reida a
Western Australian Biomedical Research Institute, Division of Health Sciences, School of Veterinary and Biomedical Sciences, Murdoch University, South Street, Murdoch, WA 6150, Australia b Division of Health Sciences, School of Veterinary and Biomedical Sciences, Murdoch University, South Street, Murdoch, WA 6150, Australia c Kenya Trypanosomiasis Research Institute (KETRI), P.O. box 362, Kikuyu, Kenya Received 27 February 2004; received in revised form 18 June 2004; accepted 26 June 2004
Abstract Camel trypanosomosis (Surra) causes high morbidity and is an impediment to the camel husbandry in Kenya. The lack of a sensitive diagnostic test has hindered the collection of accurate epidemiological data and institution of control programmes. A cross-sectional study was conducted in three districts of Kenya to estimate the prevalence of Trypanosoma evansi (T. evansi) and to compare four diagnostic tests: polymerase chain reaction (PCR), card agglutination test (CATT/ T. evansi), microhaematocrit centrifugation technique (MHCT) and mouse inoculation (MI). A total of 549 camels were randomly sampled. The overall prevalence of Surra was 5.3% using MHCT, 26.6% using PCR and 45.9% using CATT/T.evansi. There was a significant difference (P < 0.001) between PCR and CATT/T.evansi test, MHCT and MI in detection of T. evansi. The prevalence of T. evansi was 39.8% in Samburu, 24.7% in Nanyuki and 14.4% in Isiolo districts using PCR. A male camel was 2.6 times more likely to be infected with T. evansi compared to a female camel (OR = 3.0% CI: 1.6, 4.1), while an adult camel was 2.2 times more likely to be infected compared to non-adults (OR = 2.2; 95% CI: 1.2, 5.0). There was a poor association between the presence of the published clinical signs and seropositivity (k = 0.12), PCR (k = 0.11) and MHCT (k = 0.05). However, there was a higher agreement between farmers’ classification of disease with the PCR test (k = 0.5, n = 61). The mean PCV varied with age, presence of infection, locality and gender, with the lowest mean PCV being recorded in MHCT-positive * Corresponding author. Tel.: +61 08 9360 2690; fax: +61 08 9310 4144. E-mail address: [email protected] (Z.K. Njiru). 0304-4017/$ – see front matter # 2004 Elsevier B.V. All rights reserved. doi:10.1016/j.vetpar.2004.06.029
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animals (20.97 0.5) and from infected calves (19.5 1.2). This study shows that PCR was more sensitive in detecting T. evansi than other tests used. Further, the prevalence of T. evansi in the camel herds sampled is higher than that previously reported in Kenya, and that the judgment by camel keepers may be a reliable ‘‘pen-side’’ diagnostic test for Surra. Considering the low sensitivity of parasitological techniques in detection of chronic T. evansi infection and high cost of PCR, development of a sensitive pen side diagnostic test, with a low cost is still a priority. # 2004 Elsevier B.V. All rights reserved. Keywords: Trypanosoma evansi; Camel; PCR; CATT/T. evansi; Trypanosomosis
1. Introduction Trypanosma evansi (T. evansi), the cause of cameline trypanosomosis (Surra), is the most widespread pathogenic trypanosome in the world. It is mechanically transmitted by haemotophagus biting flies and therefore distributed widely outside the tsetse belts. The disease causes significant morbidity and mortality in camels in arid and semi arid regions in Kenya, which have a population of over 1 million camels and support over 25% of human population and over half of livestock production (Anonymous, 1997). Trypanosomosis in camels occurs both in chronic and acute forms (Wilson et al., 1983). The chronic form is most common and may present an association with secondary infections due to immunosuppression caused by T. evansi infection, which complicates clinical diagnosis. Published clinical signs (emaciation, fever, anaemia, lacramation, corneal opacity, diarrhoea and oedema of the dependent parts) (Chaudhary and Iqbal, 2000) are insufficient for diagnosis, while detection of parasites in the blood is difficult because parasitaemia is intermittent (Mahmound and Gray, 1980). Consequently, there is a need for alternative more sensitive diagnostic techniques. Serological tests have been developed and evaluated for diagnosis of trypanosomosis in camels. The antibody-detecting tests such as the CATT/T. evansi (Bajyana-Songa and Hamers, 1988) and enzyme-linked immunosorbent assay (Ab-ELISA) (Atarhouch et al., 2003) have been shown to be reliable while the antigen based Suratex1 (Nantulya, 1994) and Ag-ELISA (Olaho-Mukani et al., 1993) are seldom used. CATT/T. evansi, which is a commercially produced kit, is widely used in the field. However, it is unable to distinguish past (treated) from current infections because it detects antibodies that may persist for a long time following treatment. The polymerase chain reaction (PCR) is highly sensitive and specific and has been widely used in detection of trypanosomes. Primers targeting subgroup Trypanozoon (Moser et al., 1989; Masiga, 1994; Wuyts et al., 1994) and putatively T. evansi-specific (Artama et al., 1991; Masiga, 1994; Ventura et al., 2002) have been developed. The PCR has been used in surveys to determine the prevalence of T. evansi in buffaloes in Vietnam (Holland et al., 2001) and T. equiperdum in Mongolian horses (Clausen et al., 2003). However, PCR has not been systematically used for the detection of T. evansi in camels in Kenya (Masiga and Nyang’ao, 2001). In this study, we determined the prevalence of T. evansi in three camel-keeping regions of Kenya and compared the diagnostic utility of PCR, CATT/T. evansi, MHCT and mouse inoculation for the detection of T. evansi infections in camels.
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2. Materials and methods 2.1. Sampling sites A survey was conducted in three districts Samburu (Wamba and Ngurunit), Isiolo (Tractor, Eremet, Isiolo town and Ngare Ndare) and Nanyuki (Nanyuki town and Eldana) districts in August, 2003. The districts lie between longitudes 3684W and 3684 27E and between latitudes 0817S and 0845N. Rearing of camels is the major activity in these regions although small ruminants (sheep and goats) are also kept. The camels are moved by owners within a defined area depending on the availability of pasture. Rainfall is bimodal in pattern with long rains occurring between March and May and short rains from October to December. The annual rainfall and temperatures range from 500 to 900 mm and 20 to 37 8C, respectively. In Isiolo and Nanyuki, vegetation is predominantly savannah with Acacia species; communities of wooded grassland and bush land thickets are found along seasonal watercourses. In Samburu, most of the regions containing camels are grasslands with scattered Acacia species and short shrubs, with stretches of bare land. Numerous biting flies (Haematobia minuta) are prevalent in Nanyuki and Isiolo with few recorded occurrences of Tabanus taeniola, Hippobosca camelina and H. variegata (Njiru et al., 2002). 2.2. Study design and sampling A cross-sectional sampling procedure was used during this study. A total of 549 camels were sampled, 153 in Isiolo, 161 in Samburu and 235 in Nanyuki district, respectively. The sampling areas were chosen because information from respective district veterinary offices suggested the presence of Surra. Before the field trip, information was passed to the camel owners through their community leaders. On the day of sampling, animals were selected using a random number generator, marked using permanent spray, then information on age, gender, previous trypanocide treatment, history of abortions and weight estimations was recorded. Each animal was clinically examined, and 5 ml of blood was collected from the jugular vein. Two millilitres of blood was placed into a tube containing heparin for PCV, parasite examination and mouse inoculation, and 3 ml blood was placed in plain tubes and left to clot for serum collection. The serum was removed after 12 h and stored in liquid nitrogen. Collected heparinised blood samples were stored in cool boxes and transported to a temporary laboratory within the immediate vicinity. Approximately, 0.5 ml of blood was transferred into cryovials, mixed with phosphate saline glucose (PSG) and 10% glycerol, and stored in liquid nitrogen for subsequent extraction of genomic DNA. Animals were grouped into age groups; calves <12 months, immature 12 and 36 months and adults >36 months for purposes of analysis. 2.3. Parasitological examination and mouse inoculation Two capillary tubes were prepared from each heparinised blood sample and centrifuged in a microhaematocrit centrifuge for 5 min at 12,000 g. The packed cell volume (PCV) was
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determined, and both capillary tubes were examined for the presence of motile trypanosomes using the microhaematocrit centrifugation technique (MHCT) (Woo, 1970) and the buffy-coat technique (Murray et al., 1977). Four hundred microlitres of blood from animals with a PCVof 26% and/or showing clinical signs suggestive of Surra (Chaudhary and Iqbal, 2000) were inoculated into Swiss-bred mice, which were screened for infection with T. evansi by wet smear examination of tail-tip blood every day for 60 days. Any camel shown to be infected with T. evansi using MHCT and those animals showing clinical signs suggestive of Surra (emaciation, PCV 20% or a recent history of abortion) were treated with quinapyramine sulphate/chloride (Triquin1) intramuscular at 5 mg/kg body weight. 2.4. Serological tests Sera were tested for the presence of anti-T. evansi antibodies using the card agglutination test for T. evansi (CATT/T. evansi) (Institute of Tropical Medicine, Antwerp, Belgium). Approximately, 45 ml of the antigen was transferred into the test card and mixed with an equal amount of the test sera diluted at 1/4 with PBS pH 7.2 as per manufacturer’s instructions. The card was agitated for 5 min, and the reaction was checked in the clear light. Positive reaction was confirmed on recording agglutinations (blue granules) (Bajyana-Songa and Hamers, 1988). 2.5. Template preparation and DNA amplification Genomic DNA was extracted from samples of camel blood using commercially available QIAamp DNA mini kit (Qiagen, 2003), and the purified DNA template was stored at 20 8C for later use. PCR was carried out in 25 ml reaction mixtures containing 10 reaction buffer (Fischer Biotech), 1.5 mM MgCl2, 200 mM of each of the four deoxynucleoside triphosphates (dNTPs) (Promega), primers at 1 mM and 0.5 U of Taq DNA polymerase (Fischer Biotech). The cycles included an initial step at 94 8C for 4 min, followed by 30 cycles of denaturing at 948 C for 30 s, annealing at 608 C for 30 s and extension at 728 C for 30 s. PCR Elongation was continued at 728 C for 5 min. The amount of DNA template added per reaction was 3 ml. A positive (reference T. evansi DNA, 25– 50 ng/ml) and a negative control (without DNA) were included for each PCR reaction. Amplification products were resolved in 1.5% molecular grade agarose gel (Fischer Biotech) stained with ethidium bromide (5 mg/ml). 2.6. Primers Primers targeting a repetitive region specific for Trypanozoon subgenus (Wuyts et al., 1994) pMURTec.F, 50 -TGCAGACGACCTGACGCTACT; pMURTec.R, 50 -CTCCTAGAAGCTTCGGTGTCCT and T. evansi minicircle specific (Masiga, 1994) EVA1, 50 ACATATCAACAACGACAAAG, EVA2 50 -CCCTAGTATCTCCAATGAAT were used. Since the minicircle primers are less sensitive than the Wuyts primers, they were used only on those samples that gave strong reactions using the latter primers and/or on samples shown to be MHCT positive.
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2.7. Weight measurements The live body weight of camels (kg) was estimated by measuring (in meters) shoulder height, thoracic girth and abdominal girth and multiplying these by 50 (Schwartz et al., 1983). Only camels from Isiolo and Samburu districts had their weights estimated. 2.8. Statistical analysis Statistical analysis was performed using SPSS v11.5. The level of agreement between diagnostic tests was determined using (Kappa statistics (k) and the influence on prevalence of various factors, such as gender, age, locality, abortion and presence of clinical signs were determined using chi-square test and odds ratios and their 95% confidence intervals. Analyses of variance (ANOVA) with Bonferoni mean-separation tests and t-tests were used to compare mean PCV values and weights in groups of camels.
3. Results 3.1. Diagnostic tests analysis A total of 549 camels were screened for infection with T. evansi using parasitological methods (MHCT and BCT), PCR and the CATT/T. evansi. Overall prevalence was 5.3% (95% CI: 3.4, 7.2) with the MHCT, 26.2% (95% CI: 22.5, 29.9) with the PCR and 45.9% (95% CI: 41.7, 50.1) with CATT/T. evansi. The prevalence of infection in each area sampled is presented in Table 1. There was a significant difference in the prevalence estimate determined using PCR and CATT/T. evansi (P < 0.001) while prevalence estimates using MHCT was significantly lower (P < 0.001) than those of PCR and CATT/T. evansi. There was fair agreement between PCR and MHCT (k = 0.3; 95% CI: 0.22, 0.38) and between PCR and CATT/T. evansi (k = 0.2; 95% CI: 0.13, 0.27). Camel owners identified the clinical signs of Surra as rough coat, drop in milk yield, abortion and by a characteristic urine odour. There was a moderate agreement between diagnosis by camel owners and PCR (k = 0.52; 95% CI: 0.3, 0.7, n = 61). Using PCR results, a camel reared in Samburu district was four times (OR = 3.9; 95% CI: 2.3, 6.8) and two times (OR = 2.0; 95% CI: 1.3, 3.1) more likely to be infected than a camel reared in either Isiolo or Nanyuki district, respectively. Table 1 T. evansi and seroprevalence in Samburu, Isiolo and Nanyuki district District
No. of animals
MHCT
PCR
Positive (%)
95% CI
Positive (%)
CATT test 95% CI
Positive (%)
95% CI
Samburu Isiolo Nanyuki
161 153 235
20 (12.4) 2 (1.3) 7 (3.0)
7.3, 17.5 0, 3.1 0.8, 5.2
64 (39.8) 22 (14.4) 58 (24.7)
32.2, 47.4 8.8, 20.0 19.2, 30.2
68 (42.2) 53 (34.6) 131 (55.7)
34.6, 49.8 27.1, 42.1 49.3, 62.1
Total
549
29 (5.3)
3.4, 7.2
144 (26.2)
22.5, 29.9
252 (45.9)
41.7, 50.1
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Fig. 1. Prevalence of T. evansi within age structure and sex.
3.2. Age and gender The prevalence of infection estimated using PCR was significantly higher in adult camels as compared to immature and calves (P < 0.05 and P < 0.01, respectively). The prevalence of T. evansi infection estimated using PCR and MHCT was also higher in male compared to female camels (x2 = 17.0, df = 1, P < 0.001 and (x2 = 9.1, df = 1, P < 0.01, respectively) (Fig. 1). Conversely, the seroprevalence was higher in adult camels compared to immature camels and calves (P < 0.001). Adult camels were 2.2 times more likely to be CATT/T. evansi-positive compared to non adult camel (OR = 2.2; 95% CI: 1.2, 5.0). There was no significant difference in the seroprevalence between the genders while male camels were 2.6 times more likely to be T. evansi-positive using PCR test compared to female camels (OR = 2.6; 95% CI: 1.6, 4.1). 3.3. Packed cell volume The lowest PCV means were recorded for MHCT-positive animals (20.9 0.5), infected calves (19.5 1.4) and animals from Samburu district (23.1 0.4). Overall the mean male camels had significantly lower PCVs compared to female camels (P < 0.05) and immature camels had significantly lower PCVs compared to adult camels (P < 0.001) (Table 2). The PCV of camels shown to have circulating T. evansi by PCR or MHCT was significantly lower than aparasitaemic animals (P < 0.001). There was no significant difference in mean PCV of seropositive and seronegative camels. 3.4. Sensitivity Of 29 MHCT-positive samples collected in this study, 28 (96.6%) were PCR-positive and 19 (65.5%) CATT/T. evansi-positive. Using both tests, 19 samples were positive and only one was negative on both tests. In earlier and separately collected samples, PCR
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Table 2 Packed cell volume for camels in Samburu and Isiolo district Category
n
Age Adult
Nanyuki
22.98 0.52 23.69 0.63
P = 0.4
Infected Non infected
103 351
24.28 0.43 25.51 0.18
P < 0.01
23.48 0.43a
Infected Non infected
454
b
25.23 0.17
n
397
25.55 0.18c
Infected Non infected
119 278
24.59 0.36 25.97 0.20
P < 0.001
84
24.95 0.35c
Infected Non infected
15 69
21.47 0.60 25.71 0.34
P < 0.001
Infected Non infected
10 58
19.50 1.39 21.36 0.41
P = 0.12
Infected Non infected
64 97
22.48 0.62 23.53 0.43
P = 0.16
Infected Non infected
22 131
26.77 0.75 25.33 0.29
P = 0.06
Infected Non infected
58 177
24.40 0.28 26.17 0.23
P < 0.001
Calves
Isiolo
41 54
95
Immature
Districts Samburu
t-test
Status
Gender Male Female
PCV S.E.M.
Mean PCV
68
d
21.09 0.4
161
23.11 0.36b
153
a
235
25.54 0.28
a
25.73 0.19
Superscripts that are different are significantly different within each group for mean PCV (mean PCV for age and district was analyzed with ANOVA, P < 0.05, while gender was done with t-test, P < 0.05).
sensitivity was 70/72 (97.2%) and 48/72 (66.7%) for CATT/T. evansi test. Mouse inoculation sensitivity was done for 22 positive samples from Samburu and Isiolo of which 19 (86.4%) were positive. There was a poor association but still significant difference between MI and PCR in detecting T. evansi infections in 175 aparasitaemic samples (k = 0.19; P < 0.001) as shown in Table 3. 3.5. Weights The estimated mean weights for adults, immature and calves were 484 9.1, 264 13.3 and 203 16.4 kg, respectively. Significant difference weights were only recorded Table 3 Relationship between PCR and mouse inoculation in detection of T. evansi infections in aparasitaemic camels with PCV of 26% and showing clinical signs of Surra Mouse inoculation
PCR Positive
Negative
Total
Positive Negative
8 42
2 123
10 165
Total
50
125
175
Fischer’s exact test, P < 0.001, k = 0.19, 95% CI: 0.06, 0.3.
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between seropositive (501 8.75) and seronegative adults (462 17.3) (P < 0.05) and between estimated weights of camels from Isiolo and Samburu (433.75 14.30 and (381.36 12.8; P < 0.001).
4. Discussion In this study, infection with T. evansi infection was demonstrated in the three districts sampled. The prevalence estimates from this study are higher than those of previous studies (Ngaira et al., 2003). There is no obvious reason why the prevalence of T. evansi in camels in Samburu district was higher than in other districts sampled. However, it is worth noting that the areas visited in Samburu district were difficult to access (poor roads especially in the rainy seasons) as compared to Nanyuki and Isiolo, which have better communication and transport network. Therefore, it is possible that some of the difference in prevalence may be because camel keepers in the latter districts have better access to trypanocidal drugs and veterinary services. The results of this study indicate that PCR is the most accurate test for the detection of T. evansi parasitaemia in camels. PCR has been used successfully in detecting infection with T. evansi in buffaloes (Omanwar et al., 1999; Holland et al., 2001), horses (Clausen et al., 2003) and in camels (Masiga and Nyang’ao, 2001). It is interesting to note that the prevalence estimate determined using PCR varied in the different districts visited with the highest prevalence in Samburu district. This may indicate differences in the epidemiology of infection because seroprevalence did not vary significantly with district, which might indicate a higher level of active infections in Samburu compared to other districts. These differences may also be due to genetic differences in T. evansi isolated from each area. Animals in Samburu district also weighed less and showed poor body conditions due to factors, which were associated with higher disease burden and poor foliage. The CATT/T.evansi test is a well-validated field assay for the detection of T. evansi in camels (Dia et al., 1997; Ngaira et al., 2003) buffalo (Davison et al., 1999) and cattle (Reid and Copeman, 2003)). It has been the test of choice for the field surveys, although being an antibody-detecting assay, it has inherent shortcomings of not being able to differentiate between past and present infections. The sensitivity of CATT/T. evansi in this study was 65.5%, which compared well with 68.6% reported in an earlier study by Ngaira et al. (2003) in Isiolo district. So far, there is no comprehensive data on the use of PCR for detecting infection in camels and/or comparison of CATT/T. evansi test with PCR. Such data are necessary in the face of camel husbandry strengthening in Kenya. A comparison of the PCR with CATT/T. evansi showed a significant difference (P < 0.001) and only a fair association (k = 0.3), implying that the two tests were categorizing infections independently. The CATT/T. evansi test failed to detect 51/144 (35.4%) of PCR positive samples and 10/29 (3.5%) of MHCT positive samples. Not only the phenomenon of parasitological positive samples but CATT/T. evansi negative test were also observed by Ngaira et al. (2003), Elsaid et al. (1998) and Davison et al. (1999). The 51 PCR positive cases missed by CATT/T. evansi may indicate early infections where animals have not yet formed detectable antibody levels or isolates that lack Ro Tat 1.2 gene and/or do not express Ro Tat 1.2 VSG. If treatment had been instigated based on the results of CATT/T.
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evansi, 252 animals would have been treated as opposed to 144 animals if the results had been based on the PCR. Assuming no false positive with PCR, this indicates 108 (43%) unnecessary treatments. This means that in an area where treatment is frequent, CATT/T. evansi test needs to be used with second confirmatory test. In our case, PCR, although expensive, it was the most appropriate in detecting both chronic and current infections. The CATT/T. evansi is based on variable surface antigen (VSG) designated as RoTat 1.2, which is thought to be expressed in all T. evansi isolates (Verloo et al., 2001). However, recently Ngaira et al. (2004) have shown isolates from Kenya that lack both Ro Tat 1.2 gene and their associated VSG and another group of isolates with Ro Tat 1.2 gene but absence of expressed VSG. Earlier studies showed variable expression of VSG genes in the closely related species T. b. gambiense (Dukes et al., 1992). If the recently observed variations are widespread in Kenya camel-rearing regions, then it may explain the low sensitivity of the CATT/T. evansi observed. Moreover, isolates of T. evansi from Kenya have also been shown to differ genetically (Gibson et al., 1983) and the existence of T.evansi type B suggests some genetic variation occurring in the Kenyan northern frontier (Borst et al., 1987). Further ongoing studies of isolates from the same study areas reveal regional genetic differences in T. evansi isolates (Njiru, unpublished results). The possible occurrence of infections with other Trypanozoon parasites was ruled out by testing most of the PCRpositive isolates (n = 88) with T. evansi-specific minicircle primers (Masiga, 1994). Ongoing work will confirm whether our 10 MHCT-positive but CATT/T. evansi-negative isolates express the RoTat 1.2 gene. However, one way to improve CATT/T. evansi sensitivity would be to add other not additional to other antigens in the test. The PCR test had a sensitivity of 97% in this study and 97.2% in a separate study using Trypanozoon spp primers (Wuyts et al., 1994). The one missed case in this study could either be due to degraded DNA and/or loss of DNA during extraction. The strength of PCR was further shown in detection of infection in aparasitaemic animals showing clinical symptoms (Table 3). Analysis of animals with PCV of 26% and some with clinical signs of Surra (lachrimation, diarrhoea, emaciation, enlargement of lymph nodes and recent aborted females) showed T. evansi prevalence of 50/175 (28.6%) and 10/175 (5.7%) with PCR and MI. When all clinical symptoms were combined together, a fair association between clinical signs and seroprelevalence (k = 0.3; 95% CI: 0.22, 0.38) and low association with PCR (k = 0.1; 95% CI: 0.05, 0.19) were recorded. These results are suggestive that Surra may not be the only cause of the observed clinical signs. This argument is reflected in the low numbers of infected animals showing each clinical symptom, e.g. 1/7 (diarrhoea), 5/15 (lacrimation) and 10/26 (emaciated) were positive either with MHCT or PCR. Documented clinical signs of Surra are general for most of the susceptible animals. In the absence of other diagnostic techniques, this leaves treatment of camels based on personal opinion/experience. An interesting strong association (k = 0.5; 95% CI: 0.32, 0.72, n = 61) was shown in this study between signs recognized by camel keepers (urine with a characteristic smell, drying up of lactating camels, reluctance to move, rough coat and abortions) and infection in animals (MHCT and PCR). The characteristic smell of urine is due to ketones produced after breakdown of amino acids by parasite in infected animals (Hunter, 1986). Though the sample size was small, it appears that the camel owners have dependable diagnostic knowledge of Surra having lived with these animals for generations. Use of parasitological technique in T. evansi detection still
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remains wanting because of the chronic nature of T. evansi in the field. Both BCT and MHCT will not be done away soon considering the cost of other diagnostic techniques. However, these techniques may be made more useful if clinical sign guidelines are established with disease prevalence and are used side by side. In this study, 36 camels were treated in Samburu and Isiolo district based on results of PCV, MHCT and the presence of clinical signs in the field. However, the PCR results indicated that 86 of the sampled animals were infected in these regions. If the PCR indicated active infection, then 58.1% of infected animals could not be detected through the most commonly used method. If we included CATT/T. evansi test results, the number treated could have gone up to 52 still leaving 39.2% animals as untreated but also increasing the number of treated ‘‘false’’ negative. These undetected cases would eventually act as a source for future infection in the herds. MI is a frequently used technique in the field to detect latent infections. In our study, MI showed a sensitivity of 86.4% of parasitaemic cases, however it picked only 16% (8/50) of PCR-positive aparasitaemic cases. The low prevalence identified with the MI may be due to trypanosomes being lost through the mice immune systems. Sensitivity of MI may be improved by use of irradiated mice though expensive. The observation that seropositivity increases with age is in agreement with previous studies (Dia et al., 1997; Atarhouch et al., 2003; Gutierrez et al., 2000) and is probably related to the chronic nature of infection and possibility that antibodies persist after a camel is treated. However, it was also noted that the camel owners prefer to graze calves within the vicinity of human dwellings as opposed to older camels which are grazed and watered in the open and hence have greater potential for contact with vectors. There is insufficient data to explain the observed differences in the prevalence of infections between males and female camels. One possibility is that hormonal differences may influence vector behaviour and increase the likelihood of infection in male compared to female camels. Twenty-six animals were reported to have aborted between 1 and 6 months period prior to this study. When camel keepers routinely treat these animals with trypanocides, immediately an abortion occurs. Of these animals, 14/26 (53.8%) were still positive for T. evansi with PCR (including 3 MHCT-positive cases) and the same number were seropositive. Ten animals with active infection also were seropositive and eight animals were negative with both PCR and CATT/T.evansi test. Active infection in these already treated camels indicates either re-infection or treatment failure. Misuses of drugs through under dosing (often self-administrated by the camel keepers without correct weight adjustment or using a single vial to treat several animals) and fake drugs (Ngaira et al., 2002) may have consequential effects on T. evansi prevalence and the development of resistance. There was a significant relationship between infection and low PCV. Previous authors have tried to determine a cut-off value for PCVs that can be used for diagnostic purposes. For example, <23% was proposed by Ngaira et al. (2002) in Kenya, 20% by Chartier et al. (1986), in Mauritania and 18% by Diall et al. (1993) in Mali. Though anaemia could result from T. evansi infection, other factors such as infection with Haemonchus spp and the effects of drought could cause similar effects. In this study, we found that camels shown to be PCR and MHCT-positive had significantly lower PCVs compared to uninfected camels.
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We further found significant differences in PCV by locality, gender and age (Table 2). Therefore, it is difficult to use PCV as the sole indicator of infection with T. evansi. Trypanosomosis continues to pose a great risk to camel keeping in Kenya. Low sensitivity of diagnostic techniques is a major factor impeding the control of this disease. This study has shown that Surra is prevalent in all three districts sampled at significantly different levels. The PCR test showed the best sensitivity compared with CATT/T. evansi using MHCT positive samples as the gold standard. Noting the chronic nature of the disease and apparent low sensitivity of CATT/T. evansi in Kenya, the use of PCR is recommended for inclusion in survey and control programmes. In instances where treatment has to be carried out immediately in the field, consideration of the farmers’ diagnosis with respect to documented signs by veterinarian is necessary. However, we recommend a further comprehensive study on farmers’ diagnostic knowledge versus disease prevalence in the camel herds. In cases where low PCV as a measure of infection is used, considerations of other factors like age and genders are necessary.
Acknowledgements The authors gratefully acknowledge the financial support from the International Foundation for Science (IFS grant number B3372 to Z.K.Njiru), Kenya government and Murdoch University. Supplemental funds were contributed by Western Australia Biomedical Research Institute (WABRI). We also thank Dr. Thuita, Dr. Chemuliti, Rashid Farah and Veterinary department, Isiolo district for helping with field sampling, Purity Njoka, John Ndichu and Samuel Guya for excellent technical assistance.
References Artama, W.T., Agey, M.W., Danelson, J.E., 1991. DNA comparison of Trypanosoma evansi (Indonesia) and Trypanosoma brucei spp. Parasitology 104, 67–74. Atarhouch, T., Rami, M., Bendahman, M.N., Dakkak, A., 2003. Camel trypanosomosis in Morocco 1: results of a first epidemiological survey. Vet. Parasitol. 111, 277–286. Bajyana Songa, E., Hamers, R., 1988. A card agglutination test (CATT) for veterinary use based on early VAT RoTat 1.2 of Trypanosoma evansi. Ann. Soc. Belg. Med. Trop. 68, 233–240. Borst, P., Fase-Fowler, F., Gibson, W.C., 1987. Kinetoplast DNA of Trypanosoma evansi. Mol. Biochem. Parasitol. 23, 31–38. Clausen, P.H., Chuluun, S., Sodnomdarjaa, R., Greiner, M., Noeckler, K., Staak, C., Zessin, K.H., Schein, E., 2003. A field study to estimate the prevalence of Trypanosoma equiperdum in Mongolian horses. Vet. Parasitol. 115, 9–18. Chartier, C., Chartier, F., Lepers, J.P., Pesce, J.L., 1986. Preliminary studies of common blood parameters of the Mauritanian dromedary (Camelus dromedaries). Rev. Elev. Med. Vet. Pays. Trop. 39, 395–401. Chaudhary, Z.I., Iqbal, M.J., 2000. Incidence, biochemical and haematological alterations induced by natural trypanosomosis in racing dromedary camels. Acta Trop. 77, 209–213. Davison, H.C., Thrusfield, M.V., Muharsini, S., Husein, A., Partoutomo, S., Rae, P.F., Masake, R., Luckins, A.G., 1999. Evaluation of antigen and antibody detection tests for Trypanosoma evansi infection of buffaloes in Indonesia. Epidemiol. Infect. 123, 149–155. Dia, M.L., Diop, C., Aminetou, M., Jacquiet, P., Thiam, A., 1997. Some factors affecting the prevalence of Trypanosoma evansi in camels in Mauritania. Vet. Parasitol. 72, 111–120.
198
Z.K. Njiru et al. / Veterinary Parasitology 124 (2004) 187–199
Diall, O., Bocoum, Z., Diarra, B., Sanogo, Y., Coulibaly, Z., Waigalo, Y., 1993. Epidemiology of trypanosomosis caused by T. evansi in camels in Mali: results of parasitological and clinical survey. Rev. Elev. Med. Vet. Pays. Trop. 46, 455–461. Dukes, P., Gibson, W.C., Gashumba, J.K., Hudson, K.M., Bromidge, T.J., Kaukus, A., Asonganyi, T., Magnus, E., 1992. Absence of the LiTat 1.3 (CATT antigen) gene in Trypanosoma brucei gambiense stocks from Cameroon. Acta Trop. 51, 123–134. Elsaid, H.M., Nantulya, V.M., Hilali, M., 1998. Diagnosis of Trypanosoma evansi infection among Sudanese camels imported to Egypt using card agglutination test (CATT) and antigen detection latex agglutination test (Suratex). J. Protozool. Res. 8, 194–200. Gibson, W.C., Wilson, A.J., Moloo, S.K., 1983. Characterization of Trypanosoma (Trypanozoon) evansi from camels in Kenya using isoenzyme electrophoresis. Res. Vet. Sci. 34, 114–118. Gutierrez, C., Juste, M.C., Corbera, J.A., Magnus, E., Verloo, D., Montoya, J.A., 2000. Camel Trypanosomosis in the Canary Islands: assessment of seroprevalence and infection rates using the card agglutination test (CATT/ T. evansi) and parasite tests. Vet. Parasitol. 90, 155–159. Holland, W.G., Claes, F., My, L.N., Thanh, N.G., Tam, P.T., Verloo, D., Buscher, P., Goddeeris, B., Vercruysse, J., 2001. A comparative evaluation of parasitological tests and a PCR for Trypanosoma evansi diagnosis in experimentally infected water buffaloes. Vet. Parasitol. 97, 23–33. Hunter, A.G., 1986. Urine odour in a camel suffering from Surra T. evansi infection. Trop. Anim. Health Prod. 18, 146–148. Anonymous, 1997. The Eighth National Development Plan, Government printers, Nairobi, Kenya, pp. 76–77. Masiga, D.K., 1994. The Development and Application of the Polymerase Chain Reaction Methodology for the Identification of African Trypanosomes, Ph.D. Thesis, University of Bristol. Masiga, R.C., Nyang’ao, J.M., 2001. Identification of trypanosome species from camel using polymerase chain reaction and procyclic transformation test. Camel Pract. Res. 8, 17–22. Mahmound, M.M., Gray, A.R., 1980. Trypanosomosis due to T. evansi (Istell, 1885; Balbiani, 1885). A review of recent research. Trop. Anim. Health Prod. 12, 35–47. Moser, D.R., Cook, G.A., Ochs, D.E., Bailey, C.P., McKane, M.R., Donelson, J.E., 1989. Detection of Trypanosoma congolense and T. brucei ssp by DNA amplification using the polymerase chain reaction. Parasitology 99, 57–66. Murray, M., Murray, P.K., McIntyre, I.M., 1977. An improved parasitological technique for the diagnosis of African Trypanosomosis. Trans. R. Soc. Trop. Med. Hyg. 71, 325–326. Nantulya, V.M., 1994. Suratex: a simple latex agglutination antigen test for diagnosis of Trypanosoma evansi infections (Surra). Trop. Med. Parasitol. 45, 9–12. Olaho-Mukani, W., Munyua, W.K, Mutugi, M.W., Njogu, A.R., 1993. Comparison of antigen and antibody selection enzyme immunoassays for the diagnosis of Trypanosoma evansi infections in camels. Vet. Parasitol. 45, 231–240. Ngaira, J.M., Bett, B., Karanja, S.M., 2002. Animal-level factors for Trypanosoma evansi infection in camels in eastern and central parts of Kenya. Onderst. Vet. Res. 69, 263–271. Ngaira, J.M., Bett, B., Karanja, S.M., Njagi, E.N.M., 2003. Evaluation of antigen and antibody rapid detection tests for Trypanosoma evansi infection in camels in Kenya. Vet. Parasitol. 114, 131–141. Ngaira, J.M., Njagi, E.N.M., Ngeranwa, J.J.N., Olembo, N.K., 2004. PCR amplification of Ro Tat 1.2 VSG gene in Trypanosoma evansi isolates in Kenya. Vet. Parasitol. 120, 23–33. Njiru, Z.K., Bett, B., Mapeny-Ole, I.M., Githiori, J.B., Ndung’u, J.M., 2002. Trypanosomosis and helminthosis in camels: comparison of ranch and traditional camel management systems in Kenya. Camel Pract. Res. 9, 67– 71. Omanwar, S., Rao, J.R., Basagoudanavar, S.H., Singh, R.K., Butchaiah, G., 1999. Direct and sensitive detection of Trypanosoma evansi by polymerase chain reaction. Acta Vet. Hung. 47, 351–359. Reid, S.A., Copeman, D.B., 2003. The development and validation of an antibody-ELISA to detect Trypanosoma evansi infection in cattle in Australia and Papua New Guinea. Prev. Vet. Med. 61, 195–208. Schwartz, H.J., Dolan, R.B., Wilson, A.J., 1983. Camel production in Kenya and its constraints. Trop. Anim. Health Prod. 15, 169–178. Verloo, D., Magnus, E., Buscher, P., 2001. General expression of Ro Tat variable antigen type in Trypanosoma evansi isolates from different origin. Vet. Parasitol. 97, 183–189.
Z.K. Njiru et al. / Veterinary Parasitology 124 (2004) 187–199
199
Ventura, R.M., Takeda, G.F., Silva, R.A.M.S., Nunes, V.L.B., Buck, G.A., Teixeira, M.M.G., 2002. Genetic relatedness among Trypanosoma stocks by random amplification of polymorphic DNA and evaluation of a synapomorphic DNA fragment for species-specific diagnosis. Int. J. Parasitol. 2, 53–63. Woo, P.T.R., 1970. The haemotocrit centrifuge technique for the diagnosis of African trypanosomes. Acta Trop. 27, 384–386. Wilson, A.J., Schwartz, H.J., Dolan, R., Olaho-Mukani, W., 1983. A simple classification of different types of trypanosomosis occurring in four camel herds in selected areas of Kenya. Tropenmedizin und Parasitologie 34, 220–224. Wuyts, N., Chokesajjawatee, N., Panyim, S., 1994. A simplified and highly sensitive detection of Trypanosoma evansi by DNA amplification. S.E. Asian J. Trop. Med. Publ. Health 25, 266–271.