This cross-sectional study was conducted in four districts across southern Egypt: Deshna (26°7′18.595"N, 32°28′17.511"E) and Naja' Hammadi (26°2′55.6"N, 32°14′25.12"E) in the Qena governorate, Girga (26°20′13.96"N, 31°53′34.6"E) in the Sohag governorate, and Esna in the Luxor governorate (25°17′24.3"N, 32°33′23"E). These areas were selected based on their high livestock densities and documented history of tick infestations and tick-borne diseases, making them epidemiologically relevant for studying tick-pathogen dynamics. The hot and arid climate of southern Egypt, characterized by minimal annual rainfall and elevated temperatures, provides favorable conditions for tick survival and pathogen transmission. The geographical positions of the study areas within southern Egypt are shown in Figure 1.

The current study was conducted to investigate the presence and molecular diversity of tick species and associated piroplasmids in southern Egypt. The aim was to provide baseline data on tick infestation patterns and tick-borne protozoa with a focus on identifying emerging or underreported tick-borne pathogens. Collected ticks were morphologically identified and pooled according to species, developmental stage, and geographic origin for molecular analysis targeting piroplasmids and related protozoan parasites. All laboratory procedures, including DNA extraction, PCR amplification, and sequencing, were performed using standardized protocols to ensure reproducibility and accuracy.

A total of 110 cattle were selected for tick collection during routine veterinary inspections conducted between January and July 2019. The sample size was determined based on accessibility and logistical constraints, with the aim of generating preliminary baseline data on tick species composition and associated pathogens in the study area. We attempted to collect a uniform number (2, 3) of ticks from each bovine host. All ticks were removed manually from the medial aspect of the thighs, udder, dewlap, or axilla, taking care to minimize any discomfort to the animal. Sampling was not randomized or stratified due to logistical constraints and the observational nature of the study; however, efforts were made to include animals from multiple locations within each location to attain variability in tick populations. The owner of each cattle provided informed consent for the collection of ticks from their cattle, and the use of the samples in this study. The number of collected ticks was recorded for each cattle at each location. After collection, the ticks were preserved in tubes containing 70% ethyl alcohol. Each tick was subjected to a cleaning process before species identification by microscopy. The procedure involved placing the tick in a fine mesh and subjecting it to a gentle stream of tap water to eliminate surface dirt and debris. Subsequently, the ticks were immersed in 70% ethanol for 2 min and then rinsed twice in sterile Milli-Q water. Ticks were cleaned thoroughly to enhance the visibility of their taxonomic characteristics. Tick species were identified using standard identification keys (21–24).

Ticks underwent an initial cleansing process with 70% ethanol and were washed with sterile Milli-Q water before air-drying. Each tick was then placed in a 2-ml tube also containing a stainless-steel bead for subsequent crushing, following overnight freezing at −80°C. The crushing procedure utilized an Automill crusher (Tokken. Inc, Japan) for three cycles, each lasting 30 s at 2,000 rpm. Following crushing, 200 μl of 1 M Tris-HCl (pH = 7.5) was added to each sample tube, and the tube was well shaken for 15 min, to ensure thorough mixing of the contents. After removing the stainless-steel beads, the tubes were centrifuged at 14,000 g for 5 min at 4°C. A 200-μl amount of tick homogenate was then carefully collected for DNA isolation.

The tick samples collected from each animal were pooled according to species, life cycle stage, and sampling area before DNA extraction. One hundred seventy-seven pools were generated based on sample size (1– 3 ticks/pool). The DNA isolation process employed a fully automated nucleic acid extractor, (magLEAD 6gc, Precision System Science Co., Ltd., Chiba, Japan), employing MagDEA^® Dx SV (Precision System Science Co., Ltd., Chiba, Japan), and was conducted following the manufacturer's instructions. This method was chosen over manual extraction due to its higher efficiency, reduced contamination risk, and improved reproducibility, ensuring standardized DNA yields. The DNA concentration was determined using a Nanodrop^TM 2,000 spectrophotometer (Thermo Fisher Scientific, Waltham, MA, USA). The DNA samples were stored at −30°C for subsequent analysis.

Samples were then subjected to a conventional PCR assay to confirm the existence of tick genomic DNA and as assure the absence of PCR inhibitors. In this assay, we targeted a specific partial fragment of the 16S rRNA gene unique to tick species (25) for amplification. The PCR mixture was prepared at a total volume of 10 μl, and comprised 5 μl of 2 × Gflex PCR buffer (containing Mg2⁺, dNTP plus; Takara Bio Inc., Kusatsu, Shiga, Japan), two 0.5 μl primer solutions (one each for the forward and reverse primers) with a concentration of 10 μm, 0.5 μl of Tks Gflex^TM DNA polymerase (1.25 U/μl; Takara Bio Inc.), 3.2 μl of Milli-Q water, and 0.5 μl of the extracted DNA template, ranging in concentration from 10 to 30 ng/μl. The sequences and annealing temperatures of the used primers are stated in Table 1. Nuclease-free water was utilized as a negative control and template for contamination monitoring. A confirmed NA sample for Haemaphysalis longicornis that had been stored in our laboratory and subject to confirmatory sequencing was utilized as the positive control, and exhibited a DNA concentration of 10 ng/μl.

A nested PCR assay was used to detect the 1,550-bp fragment of the 18S rRNA gene specific to piroplasms (26). Moreover, we employed nested PCR assays to pinpoint gene fragments specific to pathogens: the 503-bp fragment of the spherical body protein-4 (SBP-4) gene for B. bovis (27), and the 412-bp fragment of the rhoptry-associated protein 1 (RAP1a) gene for Babesia bigemina (27). Additionally, a conventional PCR assay was conducted to detect the 721-bp fragment of the major merozoite surface antigen (Tams1) gene, which is specific to T. annulata (28). The PCR mixture was prepared at a total volume of 10 μl. It included 5 μl of 2 × Gflex PCR buffer (Mg²⁺, dNTP plus) (Takara Bio Inc.), two 0.5 μl primer solutions (one each for the forward and reverse primers) at a concentration of 10 μm, 0.5 μl of Tks Gflex^TM DNA polymerase (1.25 U/μl) (Takara Bio Inc.), 3.2 μl of Milli-Q water, and 0.5 μl of the extracted DNA template with a concentration ranging from 10 to 30 ng/μl. In the nested PCR assays, 0.5 μl of the first PCR product (diluted 25-fold) was used as the template in the second PCR reaction. The sequences and annealing temperatures of the used primers are provided in Table 1. All PCR assays included strict measures for contamination monitoring, and employed nuclease-free water as the negative control. Additionally, confirmed DNA samples for each pathogen (Piroplasma spp., B. bovis, B. bigemina, and T. annulata) that had been obtained from Egyptian cattle blood and stored in our laboratory were used as positive controls. These positive control samples had undergone confirmatory sequencing, and they demonstrated DNA concentrations ranging from 10 to 15 ng/μl.

The PCR products obtained from the assays above were subjected to electrophoresis on a 1.5% agarose gel in 1 × Tris-acetate-EDTA (TAE) buffer. Gel electrophoresis was performed using a Mupid electrophoresis device (Mupid Co., Ltd., Tokyo, Japan). After electrophoresis, the DNA bands were visualized using a gel documentation system with a UV device (WUV-M20; ATTO Co., Ltd., Tokyo, Japan). Bands were visualized after staining the gel with ethidium bromide at a 5 μg/ml concentration in 1 × TAE buffer.

The positive amplicons obtained from each gene detection were subjected to purification from the agarose gel, employing the NucleoSpin Gel and PCR Clean-up kit (Macherey-Nagel, Leicestershire, Duren, Germany), in accordance with the manufacturer's instructions. Subsequently, the purified DNA samples were sent to a commercial laboratory (Eurofins NSC Japan KK, Kanagawa, Japan) for sequence analysis using a 3730 × 1 DNA Analyzer (Thermo Fisher Scientific, Waltham, Massachusetts, USA). The bi-directional sequencing results were first evaluated using SnapGene Viewer software (GSL Biotech, LLC., Boston, USA) (https://www.snapgene.com/). The forward and reverse sequences were aligned and combined using MEGA 11 software to produce complete sequences for subsequent analysis.

To validate the identification of the tick-borne protozoa, the obtained sequences were compared with reference sequences in the GenBank database utilizing the BLASTn tool. Obtained sequences were aligned with reference sequences from the GenBank database, using the ClustalW algorithm in MEGA 11 software. After alignment, the sequences were trimmed, and a model test was conducted in MEGA 11 to identify the most suitable evolutionary model for the data. Subsequently, phylogenetic trees were constructed using the Maximum Likelihood method, with the Tamura 3-parameter model applied for identifying tick species, B. bovis SBP4 gene, and T. annulata Tams-1 gene. For the detection of Protozoal spp. 18S rRNA genes, the Tamura-Nei model was used as the evolutionary model. Tree reliability was evaluated by bootstrap analysis with 1,000 replicates (29).

The minimum infection rate (MIR) was calculated as the number of positive pools divided by the total number of the tested ticks within each geographic location and in relation to the examined corresponding tick spp., multiplied by 100. This metric was used to estimate the lower bound of infection at the individual tick level, particularly because samples were analyzed in pools. As prevalence could not be directly determined from pooled testing due to the inability to identify the exact number of infected individuals, MIR was selected as the primary measure of infection frequency in this study.

We collected 258 ticks from 110 cattle across the study locations. In initial morphological characterization. Specifically, in the Qena governorate, 79 ticks were retrieved from 36 cattle. Broken down by location, we collected 35 ticks from 17 cattle at Deshna (R. annulatus: 23 adult females, 11 nymphs; H. dromedarii: one adult female), 44 ticks from 19 cattle at Naja' Hammadi (R. annulatus: 35 adult females, 8 nymphs; H. dromedari: one adult male). At the Girga location in the Sohag governorate, we collected 82 ticks from 34 cattle (R. annulatus: 54 adult females, one adult male; H. dromedarii: 12 adult males, 15 adult females). At the Esna location in the Luxor governate, we collected 96 ticks from 40 cattle (R. annulatus: 68 adult females, 23 nymphs; H. dromedarii: one adult male; H. marginatum: four adult females). For each species, and at each geographical location, adult females accounted for the largest proportion of collected ticks. Representative examples displaying the morphological characteristics of each collected species are shown in Figures 2, 3.

We then targeted 177 pooled tick samples for DNA extraction and PCR analysis. Each pooled sample exhibited the expected 455-bp band of the 16S rRNA gene specific to tick species, thus confirming successful DNA extraction. Twenty PCR products representative for each morphologically identified tick species were then subjected to sequencing and phylogenetic analysis as shown in Figure 4. The H. dromedarii sequences identified in this study exhibited 99.8% similarity with H. dromedarii sequences (accession numbers MG757400 and MN960580) previously obtained from Egypt and Tunisia, respectively. Meanwhile, the H. marginatum sequences we obtained displayed a 99.3% similarity with H. marginatum sequences from Tunisia and Turkey (accession numbers OQ269610 and OQ975265, respectively). Additionally, the R. annulatus sequences we identified exhibited a 99.6% similarity with an R. annulatus sequence from Egypt (accession number: KY945491). The obtained sequences for tick species identification were deposited in the GenBank database with accession numbers PP937570, PP937571, and PP937573 for R. annulatus, PP937568 for H. marginatum, and PP937569, PP937572, and PP937574 for H. dromedarii.

In this study, 177 pooled tick DNA samples were examined using nested PCR to detect the 18S rRNA gene, which is indicative of piroplasmid species. Seventeen of these DNA samples tested positive, representing a MIR of 9.6%, with each positive finding representing a single bovine host. These 17 positive samples then underwent further PCR testing to determine the MIR of specific piroplasmid pathogens (B. bovis, B. bigemina, and T. annulata) employing primers specific for each species. A nested PCR assay targeting the SBP-4 gene of B. bovis revealed the presence of this piroplasmid pathogen in 9/17 pooled samples. Contrastingly, B. bigemina, was not identified in any sample in an assay targeting its RAP1a gene. A conventional PCR assay targeting the Tams1 gene of T. annulata revealed this pathogen was present in 12/17 pooled samples. The MIR for these tick-borne protozoans are displayed together with the absolute numbers of samples in which they were detected, in Table 2.

In Hyalomma ticks, MIR was 5.9% for T. annulata, 2.9% for B. bovis, and 0% (undetected) for T. orientalis and Colpodella spp. In R. annulatus ticks, the MIR was 4.4% for T. annulata, 3.3% for B. bovis, 0.5% for T. orientalis, and 2.7% for Colpodella spp. The MIRs for all pathogens in different tick species by sampling location are presented in Table 3. In addition, multiple pathogens were detected within single tick pools. Table 4 provides detailed information on the co-infection rates for the detected protozoa, categorized by the tick collection sites and the specific tick species.

We then submitted the 17 pooled piroplasm (18S rRNA gene)-positive samples described above for direct sequencing. Among the obtained sequences, the numbers exhibiting high similarity with existing sequences in the GenBank database were six for Colpodella spp., six for T. annulata, four for B. bovis, and one for T. orientalis. The obtained sequences for the 18S rRNA gene were submitted to the GenBank database with accession numbers PP937594, PP937595 and PP937596 for Colpodella spp., PP937591 for T. orientalis, PP937589 and PP937590 for T. annulata, and PP937592 and PP937593 for B. bovis. The phylogenetic tree depicting the evolutionary relationship between the detected protozoa and other apicomplexans is shown in Figure 5. Positive PCR products obtained with B. bovis or T. annulata-specific primers were selected for sequencing. For B. bovis (n = 1), the obtained sequence showed approximately 99% similarity with B. bovis sequences previously reported in South Africa (accession number KF626637) and Egypt (accession number LC775376). Furthermore, the obtained sequences for T. annulata (n = 2) showed 98.6% identity with T. annulata sequences previously reported in Tunisia (accession number AF214904) and China (accession numbers MH538103 and MF116148). We thus confirmed the presence of B. bovis and T. annulata in the samples analyzed. The obtained sequences were deposited in the GenBank database with accession numbers PP941969 (for B. bovis SBP-4 gene), or PP941967 or PP941968 (for T. annulata Tams1 gene). The phylogenetic trees for B. bovis and T. annulata, which illustrate the relationships and evol.utionary connections between different strains of B. bovis and T. annulata, are shown in Figures 6, 7, respectively.