Parasitological and molecular investigation of Trypanosoma evansi in dromedaries from Greater Cairo, Egypt

Abstract

In Egypt, camel trypanosomiasis is widespread. From October 2021 to March 2022, we collected 181 blood samples from apparently healthy one-humped camels (Camelus dromedarius) in Cairo and Giza Governates. The objective of this study was to assess infection rates of trypanosomes using blood smear examination and PCR-sequencing assays. Trypanosomes were detected in 8.3% (15/181) of camels by blood smear and in 23.8% (43/181) by PCR targeting the internal transcribed spacer (ITS). Based on blood smear and ITS-PCR results, and the absence of tsetse flies in the study area, we hypothesized that the Trypanosoma species was likely T. evansi. Validation using PCR based on the variant surface glycoprotein (VSG) of T. evansi Rode Trypanozoon antigen type (RoTat) 1.2 (RoTat 1.2 VSG gene) on ITS-PCR-positive samples (n=43) confirmed that 88.4% (38/43) were RoTat 1.2 T. evansi, while 11.6% (5/43) were non-RoTat 1.2 T. evansi. This marks the second report of non-RoTat 1.2 T. evansi in dromedary camels in Egypt. Considering the underestimated zoonotic risk of T. evansi in Egypt, there is a potential threat to humans, underscoring the need for a “One Health” approach to safeguard animal and human health.

Trypanosoma evansi, a member of the Trypanozoon subgenus, causes “surra,” a life-threatening disease in camels transmitted mechanically by biting insects. Surra manifests acute symptoms like fever, anaemia, and weakness, or chronic illness [16]. The disease is widespread from North Africa to Southeast Asia and is enzootic in Egypt [3, 5, 7, 17, 18, 40, 43, 46]. While T. evansi is the main pathogen causing camel trypanosomiasis, studies in Sudan [31] and Iran [6] also identified T. vivax as a second cause of camel trypanosomiasis. In Kenya, T. vivax, T. brucei, and T. congolense were found in camel blood [19]. Mixed infections with T. evansi, T. vivax, and T. congolense were reported in Saudi Arabia [2] and T. simiae in camels from Somalia [22].

Egypt’s escalating demand for affordable, high-quality protein has driven the large-scale import of camels from neighboring countries such as Sudan and Ethiopia. Upon arrival, these camels undergo quarantine at border facilities like Abu Simbel and Shalateen before being transported to slaughterhouses or dispersed across key animal markets such as Berkash near Cairo and Daraw near Aswan. These markets are crucial hubs in Egypt’s domestic animal trade, facilitating the movement of livestock across regions. However, the extensive cross-border trade of animals poses a significant risk for the spread of infectious diseases, including trypanosomiasis, potentially transforming localized outbreaks into global health threats.

Members of the Trypanozoon subgenus (T. evansi, T. brucei, and T. equiperdum) were considered distinct species, but phylogenetic studies suggest T. evansi and T. equiperdum should be subspecies of T. brucei [28, 54]. Trypanosomes are known for their kinetoplast DNA (kDNA). Trypanosoma brucei kDNA comprises 20–50 maxicircles (about 23 kb) and thousands of minicircles (about 1 kb), while T. evansi and T. equiperdum lack parts of kDNA. Trypanosoma evansi lacks maxicircles and exhibits either minicircle homogeneity or is akinetoplastic (Ak) [9]. The lack of kDNA stops T. evansi from completing its life cycle in the tsetse fly vector, yet it can spread mechanically through blood-sucking flies, enabling its distribution outside the tsetse belt. Based on minicircle restriction profiles, T. evansi is classified into type A, which is found in Africa, Asia, and South America, and type B, which is found in Africa [10].

Surra diagnosis traditionally relies on examining Giemsa-stained blood smears, which, though cost-effective, often miss T. evansi due to low parasitaemia in the chronic phase [14]. Molecular diagnostics targeting regions such as the internal transcribed spacer (ITS rRNA) [34] and the Variant Surface Glycoprotein (VSG) of T. evansi Rode Trypanozoon antigen type (RoTat) 1.2 (RoTat 1.2 VSG gene) [15] provide more specific detection of T. evansi infection. However, Ngaira and colleagues [32] found that some T. evansi isolates do not possess the RoTat 1.2 VSG gene, prompting the development of PCR assays specifically designed to detect these isolates [33].

The two main forms of human trypanosomiasis are sleeping sickness (caused by T. brucei gambiense or T. brucei rhodesiense), and Chagas disease (caused by T. cruzi). However, atypical human infections by other Trypanosoma species have been reported, including T. evansi [50]. Human susceptibility to T. evansi may be linked to a deficiency in apolipoprotein L-1 (apoL-1), a natural trypanolytic factor [52], as seen in an Indian patient [53], though a healthy Vietnamese woman with normal apoL-1 levels was also infected [51]. In Egypt, a suspected human infection with T. evansi was reported but lacked molecular confirmation [21].

Given the widespread presence of T. evansi among camels in Egypt, this study was focused on evaluating the infection rates using blood smear examinations and PCR-sequencing assays. This approach enhances the understanding of the epidemiology of trypanosomes in camels across Egypt.

MATERIALS AND METHODS

Sampling

We conducted a cross-sectional survey from October 2021 to March 2022, using a convenience sampling approach to collect blood samples from 181 apparently healthy one-humped camels (Camelus dromedarius). Sample collection relied on owner cooperation without specific criteria. Sampling took place at three sites in Cairo and Giza Governorates, Egypt: El-Basateen Abattoir (n=19) in Cairo Governorate (30°00’08.9”N, 31°16’27.4”E), El-Waraq Abattoir (n=32) (30°06’38.0”N 31°12’39.3”E), and Berkash Market (n=130) (30°08’56.8”N, 30°59’42.7”E) (Fig. 1). Blood was collected into 2 mL tubes coated with ethylene diamine tetraacetic acid (EDTA) (BD Bioscience, Bergen County, NJ, USA); at abattoirs, blood was taken post-incision of jugular vessels, and at Berkash market, it was drawn using a syringe after careful restraining. Samples were transported in an icebox to the Biotechnology Department, Animal Health Research Institute (AHRI), Dokki, Egypt, for processing.

Fig. 1.

Map of Egypt highlighting the sampling locations.

Parasitological examination

We followed the protocol described by [44] to prepare thin blood smears, which included air drying, methanol fixation, and Giemsa staining. These smears were then examined under an oil-immersion lens of a light microscope to detect the presence of blood parasites.

Genomic DNA extraction

We extracted genomic DNA from blood samples using the QIAamp^® DNA Blood Mini Kit (Qiagen, Hilden, Germany) according to the manufacturer’s instructions. A total of 200 μL of whole blood was used for DNA extraction, with a final elution volume of 60 μL. The quality and concentration of DNA were assessed using a NanoDrop™ 2000 spectrophotometer (Thermo Fisher Scientific). The extracted DNA was stored at −30°C until further use.

Polymerase chain reaction

We screened genomic DNA for trypanosomes using primer sets listed in Table 1. Each PCR was conducted in a 10 μL reaction volume with 5 μL 2x Ampdirect^® Plus (Shimadzu Corp., Kyoto, Japan), 0.05 μL BIOTAQ^TM HS DNA Polymerase (5 U/μL) (Bioline, London, UK), 0.3 μL of each primer (10 μM), 1.5 μL template DNA, and 2.85 μL UltraPure^TM DNase/RNase-Free distilled water (Invitrogen, Waltham, MA, USA). Positive controls with confirmed DNA samples and negative controls with UltraPure^TM distilled water were included in each reaction. Thermal cycling conditions were retrieved from previous studies [13, 34]. PCR products were electrophoresed on 1.5% agarose gel, stained with ethidium bromide, and visualized under UV light.

Table 1. Primer sets and PCR conditions.

Sequencing and phylogenetic analyses

We randomly selected at least 10% of positive samples per pathogen for sequencing. Positive amplicons were purified using the NucleoSpin^® Gel and PCR Clean-up kit (Macherey Nagel, Düren, Germany). Purified PCR product concentration was measured with a NanoDrop^TM 2000 spectrophotometer (Thermo Fisher Scientific, Waltham, MA, USA). Samples with sufficient DNA concentration were used for direct sequencing using the BigDye^TM Terminator v3.1 Cycle Sequencing Kit (Applied Biosystems) and ABI Prism 3100 Genetic Analyzer (Applied Biosystems, Foster City, CA, USA). Sequence reads were analysed and trimmed with SnapGene^® software (http://www.snapgene.com/). Trimmed sequences were aligned against GenBank sequences with the BLASTn tool (https://blast.ncbi.nlm.nih.gov/Blast) to determine identity percentages. Pairwise distances, best DNA substitution models, and phylogenetic analyses were conducted using the maximum likelihood method with 1,000 replications, utilizing MEGA X software [27].

GenBank accession numbers

Accession numbers were obtained by submitting sequences to the GenBank database of NCBI using the BankIt tool for coding sequences (https://www.ncbi.nlm.nih.gov/WebSub/; accessed on May 22, 2024), and the GenBank submission portal for non-coding sequences (https://submit.ncbi.nlm.nih.gov/subs/genbank/; accessed on May 22, 2024).

Statistical analyses

The study’s background data were entered and analysed using SPSS version 20.0 (SPSS Inc., Chicago, IL, USA). A binary regression model was used to explore the relationship between trypanosome infection (0 =absent, 1 =present) and factors such as camel sex, sampling month, and location. Associations were considered significant at P<0.05. Agreement between PCR and blood smear results was assessed with Cohen’s kappa coefficient using VassarStats (http://www.vassarstats.net/kappa.html).

RESULTS

Microscopic examination of Giemsa-stained blood smears detected trypanosomes in 8.3% (15/181) of the examined camels, with most cases from the Berkash market (Table 2). Trypanosomes were identified as belonging to the subgenus Trypanozoon based on morphological features described by [25, 47]. We observed various morphological features, including double kinetoplast, elongated subterminal kinetoplast, round terminal kinetoplast, two nuclei, absence or indistinct kinetoplast, and vacuole-like structure posterior to the kinetoplast (Fig. 2A–F).

Table 2. Detection rate of Trypanosoma species according to Giemsa-stained blood smear examination, ITS-PCR, and RoTat 1.2 VSG-PCR.

Sampling area	Giemsa-stained blood smears a	ITS-PCR	RoTat 1.2 VSG-PCR b
El-Basateen Abattoir	0/19 (0%)	3/19 (15.8%)	0/3 (0%)
El-Waraq Abattoir	1/32 (3.1%)	6/32 (18.8%)	5/6 (83.3%)
Berkash Market	14/130 (10.8%)	34/130 (26.2%)	33/34 (97.1%)
Total	15/181 (8.3%)	43/181 (23.8%)	38/43 (88.4%)

^aTrypanosoma species is classified as belonging to the Trypanozoon subgenus based on their morphology. ^b RoTat 1.2 VSG-PCR was performed on ITS-PCR positive samples (n=43).

Fig. 2.

Morphological features of Trypanosoma species observed in Giemsa-stained blood smears from camel blood samples under light microscopy. (A) Double kinetoplast. (B) Elongated subterminal kinetoplast. (C) Round terminal kinetoplast. (D) Two nuclei. (E) Absence or indistinct kinetoplast. (F) Vacuole-like structure posterior to the kinetoplast. K, kinetoplast; N, nucleus; VC, vacuole-like structure.

ITS-PCR analysis identified Trypanosoma species in 23.8% (43/181) of the camels surveyed, with most cases from the Berkash market (Table 2). PCR-positive samples showed a 480 bp band (Supplementary Fig. 1A), characteristic of subgenus Trypanozoon [34]. A total of 15 blood samples were positive by microscopy and ITS-PCR, while 28 were negative by microscopy but positive by ITS-PCR (Supplementary Table 1). The ITS-PCR showed higher sensitivity compared to blood smear examination with a moderate agreement level (k=0.4496). All 43 ITS-PCR-positive samples underwent further typing using RoTat 1.2 VSG PCR, a specific marker for T. evansi. RoTat 1.2 VSG PCR identified 88.4% (38/43) as RoTat 1.2 T. evansi, showing a 257 bp band, while 11.6% (5/43) were non-RoTat 1.2 T. evansi (Supplementary Fig. 1B).

The logistic regression model (Table 3), based on ITS-PCR assay results, significantly improves prediction compared to the null model, with a Chi-square value of 19.267 (P=0.002). The model accounts for approximately 15.2% of the variance in disease occurrence, as indicated by the Nagelkerke R² value. Sampling month significantly affects the outcome (P=0.001), with higher odds of disease occurrence in February-March (OR=10.062) and October (OR=5.412) compared to December. Sex did not significantly influence disease occurrence, with males having an odds ratio of 0.603 relative to females. No significant effect of location on disease occurrence was found in this analysis. Detailed cross-tabulation of trypanosome occurrence by month, location, and sex is provided in Supplementary Table 2.

Table 3. Logistic regression of factors associated with Trypanosoma infection in camels based on ITS-PCR assay results.

	Independent Variables	B	SE	W	df	P-value*	OR (95% CI)
Sex
	Male	–0.506	0.942	0.288	1	0.591	0.603 (0.095–3.823)
Sampling month (overall effect)				13.211	2	0.001*
	Oct 2021	1.689	0.603	7.841	1	0.005*	5.412 (1.660–17.646)
	Feb–Mar 2022	2.309	0.637	13.140	1	0.000*	10.062 (2.888–35.063)
Location (overall effect)				2.988	2	0.225
	El-Basateen Abattoir	–0.951	0.702	1.836	1	0.175	0.386 (0.098–1.529)
	El-Waraq Abattoir	–0.837	0.604	1.920	1	0.166	0.433 (0.133–1.415)

*P-value was considered significant at <0.05, B: regression coefficient, SE: standard error, W: wald coefficient, df: degree of freedom, OR: odds ratio (95% confidence interval).

We sequenced a total of eighteen samples. BLASTn analysis indicated that all 10 samples positive for ITS-PCR showed a 100% match with sequences of Trypanosoma species identified in GenBank as T. evansi. Similarly, samples positive for RoTat 1.2 VSG-PCR (n=8) also exhibited a 100% match with T. evansi sequences. Accession numbers of sequences and closest matches are detailed in Supplementary Table 3. Phylogenetic analysis revealed clustering of ITS-PCR-positive samples into a monophyletic clade with T. evansi isolates from various geographical locations (Fig. 3A), while RoTat 1.2 VSG-PCR-positive samples clustered together with T. evansi isolates from Palestine and India (Fig. 3B).

Fig. 3.

(A) Phylogenetic analysis of Trypanosoma species based on ITS-1 gene, using the Maximum Likelihood method and Jukes-Cantor model. The tree with the highest log likelihood (−1,196.79) is shown, with Leishmania donovani (MK404717) used as an outgroup. (B) Phylogenetic analysis of Trypanosoma evansi based on RoTat 1.2 VSG gene, using the Maximum Likelihood method and Kimura 2-parameter model. The tree with the highest log likelihood (−464.78) is shown, with Trypanosoma brucei (M21448) used as an outgroup. All analyses were conducted with 1,000 bootstrap replications. Sequences obtained in this study are highlighted in red boldface.

DISCUSSION

The overall detection rate of Trypanosoma species in Giemsa-stained blood smear was 8.3% (15/181), lower than that previously reported from Egypt [17, 43, 46] but higher than those from Sudan [31], Kenya [19], Nigeria [29], India [45], Oman [4, 25], Saudi Arabia [1, 2], Algeria [11], and Pakistan [48]. Using ITS-PCR, 23.8% (43/181) of camels tested positive, a higher rate than the previous reports from Egypt [17, 40], Iran [24], and Somalia [22], but lower than others from Egypt [7], Saudi Arabia [30], Kenya [19], Oman [4], and Iran [6]. Differences in detection rates could be due to the differences in animal husbandry practices, sample size, seasonal influences, and vector availability. In Kenya, Getahun and colleagues identified T. vivax, T. evansi, T. brucei, and T. congolense DNA in various Diptera flies (Hippobosca camelina, Stomoxys calcitrans, Tabanus spp., Pangonia rueppellii, and Glossina pallidipes), with H. camelina being more prevalent on camels compared to other species [19]. Another finding from Kenya, reported by Kidambasi and colleagues, detected T. vivax, T. evansi, and a Trypanosoma species closely related to T. melophagium in H. camelina collected from camels [26]. In Brazil, Ramos and colleagues detected T. evansi DNA in the feeding parts of Dichelacera flies (D. alcicornis and D. januarii) using a PCR assay [38]. A rare T. evansi infection in an Indian German Shepherd, likely from attacking wild animals as well as consuming wild rabbits, was confirmed through blood smear and PCR analysis [41]. In Egypt, a study suggested that hard ticks and Stomoxys flies might transmit T. evansi among camels in the northwest coast [46], though detailed information on the diversity, abundance, and seasonal dynamics of fly vectors in Egypt is lacking.

The discrepancy between ITS-PCR and Giemsa-stained blood smear results aligns with previous reports [4, 17, 19, 30, 31, 43] and is likely due to the low sensitivity of blood smear examination, especially in chronic infections with low parasitaemia [14]. Animal tested positive by ITS-PCR but negative by microscopy may have subclinical infections. Sub clinically infected animals can serve as reservoirs for pathogen transmission to healthy animals. Animals tested negative by blood smear examination and PCR might still be infected. Capewell and colleagues [12] found substantial quantities of trypanosomes in the skin of experimentally infected mice without detectable parasitaemia, which could be transmitted to tsetse vectors. This suggests that the skin of PCR-negative animals may act as a silent source of infection, indicating a need to re-evaluate current trypanosomiasis diagnostic methods.

The morphological features of trypanosomes in Giemsa-stained blood smears matched descriptions by [25, 47], identifying them as T. evansi. Trypanosoma evansi, a monomorphic parasite, primarily exhibits slender forms with a long free flagellum and a subterminal small kinetoplast, along with some intermediate forms that have a shorter free flagellum and an almost terminal kinetoplast [16]. The size and shape of T. evansi blood forms are influenced by growing conditions and the host’s immune response [49]. Some trypanosomes lacked a visible kinetoplast, a phenomenon known as dyskinetoplastic explained by [23] as changes in the ultrastructure of kinetoplast making it unstainable. The presence of cytoplasmic vacuoles likely linked to changes in osmotic pressure [42].

The virulence of T. evansi strains plays a significant role in determining the severity of infection. A study by Perrone and colleagues [37] examined two T. evansi strains: TeAp-ElFrio01 from the capybara and TeGu-Terecay323 from the donkey. The TeAp-ElFrio01 strain, considered moderately virulent, caused 100% mortality in immunosuppressed NMRI mice within 5 days and resulted in a 15% reduction in haematocrit levels. In contrast, the low virulence TeGu-Terecay323 strain led to 80% survival over 8 days, with only a 3% decrease in haematocrit. The mice infected with the TeAp-ElFrio01 strain also experienced body weight loss, while those infected with the TeGu-Terecay323 strain showed no significant weight changes. Additionally, this study observed fluctuating parasitaemia, which may explain why T. evansi could be observed under the microscope without obvious clinical manifestations. This suggests that the clinical manifestations of T. evansi infection can vary depending on the strain’s virulence.

The ITS-PCR showed a 480 bp band in all positive samples, characteristic of the Trypanozoon subgenus [34]. Although ITS-1 PCR can detect multiple trypanosome species, sequencing did not definitively identify the Trypanozoon species. Based on the blood smear examination results, ITS-PCR, and the absence of tsetse flies in the study area, we hypothesized that the Trypanozoon species was likely T. evansi. Using RoTat 1.2 VSG gene-based PCR, a specific marker for T. evansi [15], 88.4% (38/43) of ITS-PCR-positive samples were RoTat 1.2 T. evansi with a 257 bp band on the agarose gel. Sequencing further supported the hypothesis, aligning our isolates with T. evansi from Palestine and India.

In this study, 11.6% of ITS-PCR-positive samples (5/43) did not show positivity in the RoTat 1.2 VSG gene-based PCR assay, classifying them as non-RoTat 1.2 T. evansi. It was presumed that most RoTat 1.2 positive isolates likely harbour T. evansi type A minicircles or are dyskinetoplastic. Another subset lacks the RoTat 1.2 VSG but possesses the JN 2118Hu VSG. These isolates possess a rare minicircle type B, are referred to as non-RoTat 1.2 T. evansi [35]. The term “non-RoTat 1.2” should be used cautiously, as analysis of maxicircle and minicircle genes in trypanosome stocks negative for RoTat 1.2 VSG revealed the presence of T. evansi type A without RoTat 1.2, T. evansi type B, and T. b. brucei [36]. The distribution of non-RoTat T. evansi is uncertain but has been reported in Kenya, Sudan, and Ethiopia [8, 31, 32, 39]. This study is the second to report non-RoTat 1.2 T. evansi in Egyptian camels, following an earlier study that found these as single (27.6%) or mixed infections with RoTat 1.2 T. evansi (42.6%) [7]. However, no distinct symptoms differentiate RoTat from non-RoTat T. evansi infections. The detection of non-RoTat T. evansi beyond Kenya suggests its broader distribution than previously thought, underscoring the need to adapt molecular diagnostic techniques to comprehensively understand the epidemiology of different T. evansi types.

The detection rate of trypanosomes using parasitological and molecular methods showed no significant variation by sex. With advancing age, both male and female camels become more susceptible to infection due to increased exposure to vectors, as reported previously [11, 43, 46, 48]. The higher detection rates of trypanosomes during February to March compared to October to December suggest a seasonal influence, with vector density influenced by rising temperatures, particularly in spring and summer in Egypt [43, 46]. Hamed and colleagues [20] highlighted Egypt’s susceptibility to climate change, projecting increased hot extremes and reduced cold extremes, likely impacting vector density year-round.

This study faced challenges in obtaining sufficient blood samples from female camels and lacked comprehensive data on management practices. Future research should expand its scope with larger sample sizes from diverse geographical regions, including both male and female camels managed under varied conditions, to address these gaps and enhance understanding of non-RoTat 1.2 T. evansi dynamics in Egyptian camels. Given the endemicity of trypanosomiasis in Egypt, there is a high risk of cross-species infections among livestock, exacerbated by poorly studied fly vectors. Inadequate research on human trypanosomiasis also poses potential health threats, emphasizing the urgent need for a ‘One Health’ approach that integrates veterinary and human health efforts to protect both animals and humans.

CONFLICT OF INTEREST

The authors declare that they have no known competing financial interests or personal relationships that could have appeared to influence the work reported in this paper.