Sarcocystis and Hepatozoon Infections in Tongues of Bobcats (Lynx rufus) in Oklahoma, USA
1
Beltsville Agricultural Research Center, Animal Parasitic Diseases Laboratory, Agricultural Research Service, United States Department of Agriculture, Beltsville, MD 20705, USA
2
Department of Veterinary Pathobiology, College of Veterinary Medicine, Oklahoma State University, 250 McElroy Hall, Stillwater, OK 74078, USA
*
Author to whom correspondence should be addressed.
Parasitologia 2025, 5(2), 24; https://doi.org/10.3390/parasitologia5020024
Submission received: 27 March 2025
/
Revised: 5 May 2025
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Accepted: 12 May 2025
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Published: 15 May 2025
Abstract
:Archived, frozen tongues of 56 bobcats (Lynx rufus) from Oklahoma, USA, were tested for parasites. Testing for Sarcocystis infections included microscopic examination of unstained muscle squashes, histological sections of paraffin-embedded tissues, and molecular characterization. By a combination of both compression methods and histology, sarcocysts were found in 28 (50.0%) of 56 bobcats. In compression preparations, the sarcocyst wall appeared thin and protrusions were not clear. Histologically, tissues were degraded and, in most tongues, only a few sarcocysts were found, except bobcat #35. Multilocus genotyping utilizing 18S rRNA, 28S rRNA, and cox1 genes yielded sequences exhibiting 98–100% identity with both Sarcocystis arctica and Sarcocystis caninum isolates available in the NCBI database. Hepatozoon rufi-like meronts were found in two tongues and is the first report of Hepatozoon infections in bobcats in Oklahoma.
1. Introduction
Protozoa in the genus Sarcocystis parasitize virtually all endotherms, and a few species also occur in ectotherms [1]. Sarcocystis species have an obligatory two-host life cycle, alternating between an herbivore and a carnivore or omnivore. The sexual cycle is restricted to the intestines of carnivores; the asexual cycle occurs in the extraintestinal tissues of the herbivore host after it ingests water or vegetation contaminated with sporocysts. A carnivore consuming tissues contaminated with sarcocysts releases bradyzoites, which then transform into gamonts in the intestine of the carnivore. Fertilization produces oocysts in the lamina propria, where they sporulate in situ and often rupture, releasing sporocysts in the feces. Once an intermediate host ingests these sporocysts, parasites multiply in blood vessels and finally become encysted as sarcocysts, often in muscles. Sarcocystis species are generally host-specific, especially for the intermediate host (herbivore). In some carnivore or omnivore hosts, sarcocysts occur in extraintestinal muscles; the life cycles of these uncommon sarcocysts remain incomplete. Until recently, the prevailing view was that sarcocysts were rare in carnivores [1]. Information on recent reports of muscular Sarcocystis in canids and mustelids was recently reviewed [2,3,4].
Sarcocysts appear to be common in wild felids [5]. In the USA, muscular sarocysts were found in more than 50% of bobcats from Florida and Mississippi, USA [6,7,8]. Here, we report Sarcocystis infections in bobcats from Oklahoma, USA, for the first time.
Infections with species in the protozoan genus Hepatozoon are worldwide, and they parasitize both ectotherms and endotherms [9]. Hepatozoon species have two-host life cycles with sexual stags in arthropod vectors and asexual stages in many vertebrate hosts. Hepatozoon canis and H. americanum are the two important species that are transmitted by ticks, and they can cause serious disease in dogs [10]. Hepatozoon spp. also infect both domestic cats and wild felids, but their arthropod vectors are unknown [11,12,13]. Little is known of Hepatozoon infections in domestic cats in the USA [10]. Hepatozoon spp. gamonts were reported previously in bobcats from Texas and California [14,15]. Recently, merogonic stages of Hepatozoon were found in the tissues of bobcats from Mississippi and were designated as a new species, Hepatozoon rufi [16]. Here, we confirm the meronts of H. rufi and report Hepatozoon infections in bobcats from Oklahoma, USA, for the first time.
2. Materials and Methods
2.1. Samples from Bobcats
The bobcats tested here were those examined for Trichinella infections [17]. A total of 306 bobcats from 41 counties of Oklahoma were legally trapped in 2018–2019. Of these, enough tissues were available from 56 bobcats for the present study (Table 1). During interim, samples were stored frozen at −20 °C. Frozen samples were transported to the Animal Parasitic Diseases Laboratory (APDL), United States Department of Agriculture, Beltsville, Maryland, for further testing.
2.2. Cytological and Histological Examination
Muscle squashes of tongues from all 56 bobcats were examined by compression between a glass slide and a cover slip. Tongue tissues were often dehydrated and difficult to squash. For histological preparations, portions of muscles were fixed in 10% buffered formalin, processed routinely for paraffin-embedded histological sections, and examined after staining with hematoxylin and eosin (HE) or periodic acid–Schiff (PAS) reaction, counterstained with hematoxylin. Sarcocysts were enumerated in histological sections (enough tissue to fill a 4 × 2 cm area on a glass slide) and photographed using an Olympus AX-70 microscope with a DP-73 digital camera (Olympus Optical Ltd., Tokyo, Japan).
2.3. DNA Isolation and Amplification
Sarcocystis genomic DNA was extracted from infected tissues (presumed to be a single sarcocyst) of bobcat #35. The extraction was carried out using the DNeasy® Blood and Tissue Kit (Qiagen, Hilden, Germany), following the manufacturer’s protocol, with DNA stored at −20 °C. The purity and concentration of the extracted DNA were measured using a Nanodrop spectrophotometer (ThermoFisher Scientific, Waltham, MA, USA). The same procedure was used to extract Hepatozoon DNA from the tongues of bobcats #38 and #40.
Multilocus genotyping was performed for the 18S rRNA, 28S rRNA, and cox1 genes using already-published primers specific to Sarcocystis (Table 2). The PCR mixture (14.5 µL total volume) contained 2 µL of DNA template, 6.25 µL of Platinum Hot Start PCR Master Mix (Invitrogen, Waltham, MA, USA), 1 µL of 10 pmol/µL of each primer (Integrated DNA Technologies, Coralville, IA, USA), and 4.25 µL of molecular-grade water. The thermal cycling conditions included an initial denaturation step at 94 °C for 3 min, followed by 35 cycles of denaturation at 94 °C for 30 s, annealing at 60 °C for 30 s, and extension at 68 °C for 20 min, with a final elongation at 68 °C for 5 min.
The PCR products were visualized on 1.5% agarose gels. The positive samples were purified with ExoSAP-IT (Affymetrix, Santa Clara, CA, USA), and bi-directional Sanger sequencing was performed at Psomagen, Inc. (Rockville, MD, USA), on an ABI 3500xl Genetic Analyzer (Applied Biosystems™, Waltham, MA, USA). Sequences identified during the study were deposited to GenBank (accession numbers PV288307 for 18S rRNA, PV288308 for 28S rRNA, and PV294934 for cox1).
2.4. Phylogenetic Analysis
Phylogenetic analysis was performed on the nucleotide sequence of the 18S rRNA gene sequence. The analysis was performed on a selection of sequences that comprised new sequences (generated during the study) and additional sequences from closely related ones showing 98–100% homogeneity in NCBI GenBank database using BLASTn. Sequences were aligned using the ClustalW 2.1 alignment tool [20] implemented within Geneious Prime® 2024.0.5. All sequences were truncated slightly at both ends to preserve the homologous nucleotide positions for further analysis.
A model test was performed using jModelTest 2.1.7 [21] to determine the most suitable nucleotide substitution model, according to the Bayesian information criterion (BIC), and was applied to generate a Maximum Likelihood phylogenetic tree using MrBayes3.2 [22], with 1000 bootstrap values and branch lengths proportional to the number of substitutions per site. A total of 100,000 generations were taken for the phylogenetic tree. All positions containing gaps and missing data were eliminated. The final dataset included 21 taxa and 708 positions for the 18S rRNA gene. Sarcocystis myodes (accession number OM523016) was chosen as the out-group.
3. Results
3.1. Sarcocystis Infections
By a combination of both compression methods and histology, sarcocysts were found in 28 (50.0%) of 56 bobcats (Table 1). In compression preparations, the sarcocyst wall appeared thin and protrusions were not clear (Figure 1). Histologically, tissues were degraded. In most tongues, only a few sarcocysts were found, except bobcat #35 (Figure 2). Numerous sarcocysts were found in bobcat #35 (Figure 2A). All sarcocysts were mature. Sarcocysts were up to 216 µm wide; in these wide sarcocysts, the central of cysts had degenerated (Figure 2C). One slender sarcocyst was 1506 × 25 µm (Figure 2B). The sarcocyst wall was around 1 µm thick and protrusions were not clear (Figure 2D).
3.2. Hepatozoon Infections
Hepatozoon spp. meronts were found in the tongues of two bobcats (#38, #40). In bobcat #38, two meronts were seen in sections stained with HE; one was encapsulated and one was not encapsulated (Figure 3). An additional meront was seen in a section of tongue stained with PAS; this meront was encapsulated, and it contained peripherally located nuclei around a large PAS-positive residual body (Figure 3). In the HE-stained slide of tongue #40, an encapsulated meront was detected. The encapsulated meronts were 26–29 × 16–19 µm and were similar to type 2 meronts of H. rufi. The uncapsulated meront was 20 × 19 µm.
Attempts to obtain Hepatozoon DNA were unsuccessful, probably due to the low level of infection.
3.3. Molecular Analysis
The 18S rRNA sequences showed absolute identity to isolates of S. arctica (accession numbers MF596222-23, OQ689797, KX022100, KY947306, and MZ329343) and isolates of S. caninum (accession numbers KM362427 and MH469238). For the 28S rRNA gene, high sequence similarity (>98%) was observed with both S. arctica isolates (accession numbers MF596255, MF596253, and MF596057) and S. caninum (accession number MH469239) available in GenBank, and the mitochondrial cox1 sequences also displayed >99% identity to Sarcocystis caninum (accession number LC772898), isolates of S. lutrae (accession numbers MG273670 and MG273662), and isolates of S. arctica (accession numbers KY609324 and MF596290), confirming the species-level identification. The molecular analysis revealed that the Sarcocystis detected in bobcat tongue samples is either S. arctica or S. caninum, with all three gene regions (18S rRNA, 28S rRNA, and cox1) showing 98–100% sequence identity to S. arctica and S. caninum sequences. Moreover, this was consistent across nuclear and mitochondrial markers, strengthening the reliability of species identification (Table 3).
The phylogenetic tree based on the 18S rRNA gene showed that the bobcat-derived sequences (accession numbers PV288307, KF601301, KM362427, KX022101, KY947306, PP814746, KX022100-3, MH469238, and PP077100) clustered tightly within the S. arctica and S. caninum clade, with strong bootstrap support (95). We call this cluster the “S. arctica/S. caninum” clade.
Furthermore, the phylogenetic analysis reveals distinct host-specific clustering, with species primarily grouping according to their host families. The S. arctica/S. caninum clade, which includes sequences from Canis lupus, Canis lupus familiaris, Canis familiaris, and Vulpes lagopus, forms a well-supported cluster, indicating a close evolutionary relationship among Sarcocystis species infecting canids. Similarly, Sarcocystis lutrae isolates from Lutra lutra, Meles meles, and Vulpes vulpes form a separate clade, suggesting a shared evolutionary origin among carnivora-infecting species, with potential cross-host transmission to canids. Avian Sarcocystis species, including S. columbae from Columba palumbus and S. halieti from Sturnus vulgaris, group separately from mammalian species, highlighting distinct evolutionary pathways (Figure 4). These findings provide molecular evidence for the presence of Sarcocystis species in bobcats and contribute to a broader understanding of its evolutionary relationships within Sarcocystis species that infect carnivorous hosts. Moreover, the molecular analysis and the phylogenetic tree reconstructed suggest that S. arctica and S. caninum are closely related, cannot be genetically differentiated, and might be synonyms.
4. Discussion
Sarcocystis spp. were reported previously from Florida bobcats (Felis rufus floridanus), considered a subspecies of Lynx rufus [6]. In the same year, sarcocysts were found in four of six bobcats from Arkansas and named as a new species, Sarcocystis felis [23]. Only one morphologic type of sarcocyst was found in each report. Those sarcocysts were up to 2.1 mm long and had a thin (around 1 µm) wall with small hobnail-type projections on the cyst wall. Verma et al. [7] found sarcocysts in the histological muscle sections of 26 (74.2%) of 35 bobcats from Mississippi; they, too, were considered S. felis. Additionally, in two of these bobcats, sarcocysts with thicker walls bearing finger-like villi were detected; these resembled sarcocysts of S. neurona sarcocyst, a diagnosis confirmed by characterizing ITS1 [7]. Dubey et al. [8] found sarcocysts in 21 (84%) of 25 bobcats from Mississippi and morphologically documented more than three morphological types of sarcocysts in these bobcats.
The results of the present study support the view that sarcocysts are common in the muscles of bobcats and are not incidental findings. The tissues in the present study were degraded to allow for optimal morphological description of sarcocysts. Further studies using transmission electron microscopic examination and molecular characterization using more discriminatory markers will be needed to clarify how many discrete entities occur, and further fieldwork and experimental transmission studies will be needed to clarify their epizootiology.
The sequences from S. arctica and S. caninum form a distinct cluster, reflecting their close genetic relationship. The short branch lengths within this group indicate limited divergence among the isolates, potentially due to their association with a common host, such as canids. This pattern may suggest either a recent divergence from a shared ancestor or a slower evolutionary rate within this lineage. Additionally, the high bootstrap support values at the node reinforces strong statistical confidence in the robustness of this grouping [7,24,25].
Molecular markers play a crucial role in the detection and identification of closely related Sarcocystis species (i.e., S. arctica, S. caninum, and S. lutrae) in carnivores, since the classification is based solely on the cyst wall [1]). These markers, particularly those targeting ribosomal DNA regions such as the 18S rRNA and 28S rRNA genes and mitochondrial regions such as cox1, provide high specificity and sensitivity in distinguishing these closely related Sarcocystis species, which could be synonyms, thus facilitating a better understanding of the parasite’s epidemiology and host specificity [1,2,26,27,28,29]. Recently, Liao et al. [30] found S. arctica-like sarcocysts in two of six captive cheetahs (Acinonyx jubatus) from China. Based on 18S rRNA, cox1, and ITS1, sarcocysts closely resembled S. arctica. Further studies are needed to determine the number of Sarcocystis species in wild felids and their host specificity.
Hepatozoon rufi was recently described in bobcats from Mississippi [16]. Meronts were detected in 11 of 25 bobcats: in the myocardium of all 11, the tongues of 4, and the limb muscles of 5. Three types of meronts were reported. In the present study, only tongues were examined, and, thus, prevalence might have been higher if hearts were examined. The two types of meronts (with and without capsules) found here were structurally similar to those from bobcats from Mississippi; attempts to amplify Hepatozoon DNA were unsuccessful. The vector for H. rufi is unknown. The results of the present study indicate that the vector for H. rufi is present in Oklahoma.
5. Conclusions
The results demonstrate that Sarcocystis infections are common in the tongues of bobcats in Oklahoma, but species-level identification was not possible using currently available molecular markers. The presence of Hepatozoon sp. in bobcats in the USA was confirmed, but unlike Sarcocystis, infections were rare.
Author Contributions
The work described in this manuscript has not been published previously and is not under consideration for publication elsewhere. All authors have approved this submission. Material preparation and data collection were carried out by A.G. and M.R., J.P.D., A.G., B.M.R. and M.R. were involved in data analysis and manuscript write-up. J.P.D. conceptualized this study and prepared the first draft of this manuscript. All authors have read and agreed to the published version of the manuscript.
Funding
This research received no external funding.
Institutional Review Board Statement
No animal experiments were performed. All tissues were provided by hunters and trappers who legally harvested the bobcats. No permit was necessary.
Informed Consent Statement
Not applicable.
Data Availability Statement
Sequences were submitted to GenBank, and accession numbers were obtained (details provided in Table 2).
Acknowledgments
This research was supported in part by an appointment of Aditya Gupta by the Oak Ridge Institute for Science and Education (ORISE).
Conflicts of Interest
The authors declare no conflicts of interest.
Abbreviations
The following abbreviations are used in this manuscript:
| APDL | Animal Parasitic Diseases Laboratory |
| BIC | Bayesian information criterion |
| BLAST | Basic Local Alignment Search Tool |
| cox1 | Cytochrome c oxidase subunit 1 |
| HE | Hematoxylin and eosin |
| ITS1 | Internal transcribed spacer-1 |
| ML | Maximum Likelihood |
| PAS | Periodic acid–Schiff |
| NCBI | National Center for Biotechnology Information |
| USDA | United States Department of Agriculture |
| 18S rRNA | 18S ribosomal RNA |
| 28S rRNA | 28S ribosomal RNA |
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Figure 1.
Sarcocysts walls in compression smears of tongues of bobcat #35. Arrowheads point to the thickness of cyst walls. Unstained (A) thin cyst wall. (B) Thicker cyst wall. Note host cell (hc) juxtaposed with sarcocyst wall.
Figure 1.
Sarcocysts walls in compression smears of tongues of bobcat #35. Arrowheads point to the thickness of cyst walls. Unstained (A) thin cyst wall. (B) Thicker cyst wall. Note host cell (hc) juxtaposed with sarcocyst wall.
Figure 2.
Sarcocystis spp. infections in tongues of bobcats from Oklahoma. Hematoxylin and eosin stain. (A) Numerous sarcocysts (arrowheads) in bobcat #35. (B) A slender long sarcocyst (arrowheads) in bobcat #54. (C) A large sarcocyst in bobcat #15. Note sarcocyst wall (arrowhead) and central area with degenerated material (arrow). (D) Higher magnification of the sarcocyst wall of sarcocyst in Figure 1B. Note thin, smooth wall (opposing arrowheads).
Figure 2.
Sarcocystis spp. infections in tongues of bobcats from Oklahoma. Hematoxylin and eosin stain. (A) Numerous sarcocysts (arrowheads) in bobcat #35. (B) A slender long sarcocyst (arrowheads) in bobcat #54. (C) A large sarcocyst in bobcat #15. Note sarcocyst wall (arrowhead) and central area with degenerated material (arrow). (D) Higher magnification of the sarcocyst wall of sarcocyst in Figure 1B. Note thin, smooth wall (opposing arrowheads).
Figure 3.
Hepatozoon spp. meronts in sections of tongues of bobcats from Oklahoma, USA. (A–C), hematoxylin and eosin stain (HE); (D) = periodic acid–Schiff reaction (PAS) counterstained with HE. (A–C), bobcat # 38; (D) = bobcat #40. Arrowheads point to nuclei and the bar points to the capsule around the meront. (A) Type 1 meront. Note, peripherally located nuclei. (B) Type 1 meront with peripherally and centrally located nuclei. (C) Type 2 meront, apparently without any visible capsule. (D) Type 1 meront (arrow) with a large PAS-positive residual body and peripherally located nuclei.
Figure 3.
Hepatozoon spp. meronts in sections of tongues of bobcats from Oklahoma, USA. (A–C), hematoxylin and eosin stain (HE); (D) = periodic acid–Schiff reaction (PAS) counterstained with HE. (A–C), bobcat # 38; (D) = bobcat #40. Arrowheads point to nuclei and the bar points to the capsule around the meront. (A) Type 1 meront. Note, peripherally located nuclei. (B) Type 1 meront with peripherally and centrally located nuclei. (C) Type 2 meront, apparently without any visible capsule. (D) Type 1 meront (arrow) with a large PAS-positive residual body and peripherally located nuclei.
Figure 4.
Phylogenetic relationships of Sarcocystis sp. sequences isolated from the tongue of a bobcat (Lynx rufus) in Oklahoma, inferred through the 18S rRNA gene marker. Branch support values are displayed near the corresponding nodes. Species identified in this study are marked with an asterisk. The S. arctica/S. caninum clade has been highlighted with curly brackets.
Figure 4.
Phylogenetic relationships of Sarcocystis sp. sequences isolated from the tongue of a bobcat (Lynx rufus) in Oklahoma, inferred through the 18S rRNA gene marker. Branch support values are displayed near the corresponding nodes. Species identified in this study are marked with an asterisk. The S. arctica/S. caninum clade has been highlighted with curly brackets.
Table 1.
Details of the bobcat (Lynx rufus) from Oklahoma state, USA.
| APDL No. | Bobcat ID | County | Histology ID | Tissue | Compression Method | Histology | No. of Sarcocysts in Histological Sections | |
|---|---|---|---|---|---|---|---|---|
| Sarcocystis | Hepatozoon | |||||||
| 1. | 012-2019 | McCurtain | JP-14-1 | Tongue | + | + | − | 3 |
| 2. | 014-2019 | McCurtain | JP-14-2 | Tongue | − | + | − | 1 |
| 3. | 015-2019 | McCurtain | JP-14-3 | Tongue | + | + | − | 7 |
| 4. | 016-2019 | McCurtain | JP-14-4 | Tongue | − | + | − | 1 |
| 5. | 023-2019 | McCurtain | JP-14-5 | Tongue | − | − | − | 0 |
| 6. | 030-2019 | McCurtain | JP-14-6 | Tongue | − | − | − | 0 |
| 7. | 031-2019 | McCurtain | JP-14-7 | Tongue | − | + | − | 2 |
| 8. | 033-2019 | McCurtain | JP-14-8 | Tongue | − | + | − | 1 |
| 9. | 035-2019 | McCurtain | JP-14-9 | Tongue | − | − | − | 0 |
| 10. | 038-2019 | McCurtain | JP-14-10 | Tongue | − | + | − | 1 |
| 11. | 046-2019 | McCurtain | JP-14-11 | Tongue | − | − | − | 0 |
| 12. | 050-2019 | McCurtain | JP-14-12 | Tongue | − | − | − | 0 |
| 13. | 053-2019 | Atoka | JP-14-13 | Tongue | + | + | − | 2 |
| 14. | 058-2019 | Atoka | JP-14-14 | Tongue | − | − | − | 0 |
| 15. | 009-2019 | McCurtain | JP-14-15 | Tongue | + | + | − | 2 |
| 16. | 121-2019 | Carter | JP-14-16 | Tongue | − | + | − | 1 |
| 17. | 124-2019 | Carter | JP-14-17 | Tongue | − | + | − | 1 |
| 18. | 126-2019 | Carter | JP-14-18 | Tongue | − | + | − | 3 |
| 19. | 127-2019 | Carter | JP-14-19 | Tongue | − | − | − | 0 |
| 20. | 136-2019 | Choctaw | JP-14-20 | Tongue | − | − | − | 0 |
| 21. | 144-2019 | Choctaw | JP-14-21 | Tongue | + | − | − | 0 |
| 22. | 149-2019 | Creek | JP-14-22 | Tongue | − | − | − | 0 |
| 23. | 150-2019 | Creek | JP-14-23 | Tongue | + | + | − | 1 |
| 24. | 151-2019 | Creek | JP-14-24 | Tongue | + | − | − | 0 |
| 25. | 225-2019 | Johnston | JP-14-25 | Tongue | − | − | − | 0 |
| 26. | 228-2019 | Johnston | JP-14-26 | Tongue | − | − | − | 0 |
| 27. | 229-2019 | Johnston | JP-14-27 | Tongue | − | − | − | 0 |
| 28. | 230-2019 | Johnston | JP-14-28 | Tongue | + | + | − | 22 |
| 29. | 231-2019 | Johnston | JP-14-29 | Tongue | − | + | − | 2 |
| 30. | 236-2019 | Johnston | JP-14-30 | Tongue | − | − | − | 0 |
| 31. | 243-2019 | Le Flore | JP-14-31 | Tongue | − | + | − | 1 |
| 32. | 246-2019 | Le Flore | JP-14-32 | Tongue | − | − | − | 0 |
| 33. | 247-2019 | Le Flore | JP-14-33 a | Tongue | − | − | − | 0 |
| 34. | 250-2019 | Le Flore | JP-14-34 | Tongue | − | − | 0 | |
| 35. | 251-2019 | Le Flore | JP-14-35 b | Tongue | + | + | − | >40 |
| 36. | 252-2019 | Le Flore | JP-14-36 | Tongue | − | − | − | 0 |
| 37. | 255-2019 | Lincoln | JP-14-37 | Tongue | − | + | − | 1 |
| 38. | 256-2019 | Lincoln | JP-14-38 | Tongue | − | + | + | 1 |
| 39. | 257-2019 | Lincoln | JP-14-39 | Tongue | + | + | − | 3 |
| 40. | 258-2019 | Lincoln | JP-14-40 | Tongue | − | + | + | 3 |
| 41. | 260-2019 | Lincoln | JP-14-41 | Tongue | − | − | − | 0 |
| 42. | 264-2019 | Lincoln | JP-14-42 | Tongue | − | − | − | 0 |
| 43. | 265-2019 | Lincoln | JP-14-43 | Tongue | − | − | − | 0 |
| 44. | 266-2019 | Logan | JP-14-44 | Tongue | − | − | − | 0 |
| 45. | 267-2019 | Logan | JP-14-45 | Tongue | − | − | − | 0 |
| 46. | 270-2019 | Major | JP-14-46 a | Tongue | − | + | − | 1 |
| 47. | 275-2019 | Muskogee | JP-14-47 | Tongue | − | − | − | 0 |
| 48. | 276-2019 | Noble | JP-14-48 | Tongue | − | − | − | 0 |
| 49. | 281-2019 | Okmulgee | JP-14-49 | Tongue | − | − | − | 0 |
| 50. | 293-2019 | Pawnee | JP-14-50 | Tongue | − | − | − | 0 |
| 51. | 295-2019 | Pittsburg | JP-14-51 | Tongue | − | + | − | 1 |
| 52. | 296-2019 | Pittsburg | JP-14-52 a | Tongue | − | + | − | 1 |
| 53. | 339-2019 | Woods | JP-14-53 | Tongue | − | − | − | 0 |
| 54. | 340-2019 | Woods | JP-14-54 | Tongue | + | + | − | 1 |
| 55. | 345-2019 | Woods | JP-14-55 | Tongue | + | + | − | 1 |
| 56. | 347-2019 | Woods | JP-14-56 | Tongue | − | + | − | 2 |
a Positive for Trichinella. b Sarcocystis DNA amplified in the present study targeting several markers. + = present, − = not seen.
Table 2.
Primer and accession numbers of Sarcocystis sp. isolated from Lynx rufus from Oklahoma, USA.
Table 2.
Primer and accession numbers of Sarcocystis sp. isolated from Lynx rufus from Oklahoma, USA.
| Gene | Fragment Size | Bobcat ID | Cyst No. | PCR Amplification Primers (5′-3′) | Sequencing Primers (5′-3′) | GenBank Accession Numbers | Reference |
|---|---|---|---|---|---|---|---|
| 18S rRNA | 703 | #35 | 1 | 562F-TTCCCTCGTGGAAGGGTAGT 946R-TCACCGGAACACTCAATCGG | 320F-CTGGCATCCTCCTGATTGGT 807R-CGTGCAGCCCAGAACATCTA 946R-TCACCGGAACACTCAATCGG | PV288307 | [18] |
| 28S rRNA | 580 | #35 | 1 | 137F-TCAAGCCCCTGATTCTTTCACT 869R-CCACGTCTTCCTACTCATTGC | Same as PCR primers | PV288308 | [18] |
| cox1 | 776 | #35 | 1 | SF1-ATGGCGTACAACAATCATAAAGAA SR5-TAGGTATCATGTAACGCAATATCCAT | SF1-ATGGCGTACAACAATCATAAAGAA SR4-CCACACCTGTAGTACCCCC SR5-TAGGTATCATGTAACGCAATATCCAT | PV294934 | [19] |
Table 3.
Homogeneity table showing interspecific nucleotide sequence similarities between Sarcocystis sp. identified from Lynx rufus from Oklahoma, USA, and closely related taxa available in GenBank.
Table 3.
Homogeneity table showing interspecific nucleotide sequence similarities between Sarcocystis sp. identified from Lynx rufus from Oklahoma, USA, and closely related taxa available in GenBank.
| Gene | Bobcat ID | Cyst No. | Fragment Size (bp) | Accession Number | Identity of Closely Related Taxa Available in GenBank | Variations and Gaps in BLAST Search | ||
|---|---|---|---|---|---|---|---|---|
| Sarcocystis sp. (Accession Numbers) | % Query Coverage | % Identity (Average) | ||||||
| 18S rRNA | #35 | Cyst 1 | 703 | PV288307 | All isolates of S. arctica and S. caninum S. svanai (acc. no.’s KM362428 and OR921254) S. lutrae (acc. no.’s MG372102, MG372103, MG272288, KM657769 and MF596216) S. wenzeli (acc. no. MT756991) S. columbae (acc. no. GU253883) S. halieti (acc. no. MZ329690) | 60–100 | 100 | No gaps and no variations |
| 100 100 | 99.3–99.2 99.3–99.1 | One gap at the 375 position; T/C at the 461 position, and AA/TG at the 480-81 positions Did not study | ||||||
| 28S rRNA | #35 | Cyst 1 | 580 | PV288308 | S. arctica (acc. no.’s MZ329344, KX022104, MF596251, MF596242, OR921264, MF596260, MF596249, KY947309) S. caninum (acc. no.’s PP814748, MH469239) S. svanai (acc. no. OR921260) S. lutrae (acc. no.’s MG272279, MF596238, MG272276, ON796572, MT036249) S. columbae (acc. no.’s HM125053, GU253887) S. halieti (acc. no.’s OL672520, MZ329403) | 53–100 | 98.3–99.9 | Two gaps each at the 338 and 341 positions; G/A at the 345 position, G/T at the 539 position, and A/G at the 555 position in the present sequence |
| 53–100 | 95.9–96.1 | Seven gaps each at the 481, 484, 485, 486, 488, 516, and 517 positions; C/T at 73, A/G at 319, C/T at 325 and 339, C/A at 345, G/A at 348, AA/GG at 353 and 354, T/A at 360, C/T at 418, G/A at 435, T/C at 449, C/T at 454, and G/A at 497 | ||||||
| 53–100 53–100 | 96.6–96.9 96.6–96.7 | Did not study | ||||||
| cox1 | #35 | Cyst 1 | 776 | PV294934 | S. caninum (acc. no.’s LC772898, MH469240) S. arctica (acc. no.’s PQ243234, MF596290, MF596299, MF596297, KY609324, MF596305) S. svanai (acc. no. PP819578) S. lutrae (acc. no.’s PP078754, MG273668, MG273669) S. lari (acc. no.’s MF596283, MF946584) S. columbae (acc. no. MH138312) S. halieti (acc. no. MH138309) | 100 | 99.2–100 | No gaps/variations in LC772898; no gaps for MH469240, T/C at the 107 position, G/C at the 265 position, C/A at the 338 position, and A/T at the 386 position |
| 100 | 99.4 | Did not study | ||||||
| 100 | 99.4 | |||||||
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MDPI and ACS Style
Dubey, J.P.; Gupta, A.; Rosenthal, B.M.; Reichard, M. Sarcocystis and Hepatozoon Infections in Tongues of Bobcats (Lynx rufus) in Oklahoma, USA. Parasitologia 2025, 5, 24. https://doi.org/10.3390/parasitologia5020024
AMA Style
Dubey JP, Gupta A, Rosenthal BM, Reichard M. Sarcocystis and Hepatozoon Infections in Tongues of Bobcats (Lynx rufus) in Oklahoma, USA. Parasitologia. 2025; 5(2):24. https://doi.org/10.3390/parasitologia5020024
Chicago/Turabian StyleDubey, Jitender P., Aditya Gupta, Benjamin M. Rosenthal, and Mason Reichard. 2025. "Sarcocystis and Hepatozoon Infections in Tongues of Bobcats (Lynx rufus) in Oklahoma, USA" Parasitologia 5, no. 2: 24. https://doi.org/10.3390/parasitologia5020024
APA StyleDubey, J. P., Gupta, A., Rosenthal, B. M., & Reichard, M. (2025). Sarcocystis and Hepatozoon Infections in Tongues of Bobcats (Lynx rufus) in Oklahoma, USA. Parasitologia, 5(2), 24. https://doi.org/10.3390/parasitologia5020024
