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opencc-by-4.0Aug 2014View details →

Fig. 1 in Nematode-coccidia parasite co-infections in African buffalo: Epidemiology and associations with host condition and pregnancy

Fig. 1. Age, sex and seasonal patterns of infection. Both parasites had the highest (a) prevalence (sample size for calf, juvenile, subadult, adult, and senescent respectively: N = 91, 555, 221, 326, 182) and (b) mean intensity in calves and juveniles (nematode N = 78, 448, 129, 208, 100; coccidia N = 58, 237, 55, 60, 14). (c) Males had lower estimated nematode prevalence and (d) higher estimated coccidia intensity compared to female buffalo. (e) The estimated nematode prevalence, coccidia prevalence, and (f) mean coccidia intensity were all increased in the early wet season compared to the late wet season. ‡Indicates significant differences at p <0.05.

opencc-by-4.0Aug 2014View details →

Fig. 1 in Global diversity of fish parasitic isopod crustaceans of the family Cymothoidae

Fig. 1. Absolute numbers and cumulative percentage of species of Cymothoidae (373) published per decade since Linnaeus (1758). Data from the World List of Marine, Freshwater and Terrestrial Isopod Crustaceans hosted by the Smithsonian and at the WoRMS database (Schotte et al., 1995 onwards).

Fig. 4 in Heartworm and seal louse: Trends in prevalence, characterisation of impact and transmission pathways in a unique parasite assembly on seals in the North and Baltic Sea

Fig. 4. Histological sections and staining of E. horridus revealing filarial stages in E. horridus. A: Filarial stages (arrowheads) in the pharynx. bar = 20 μm. B: Filarial stage (arrow) in the mouth region. bar = 40 μm. C: Filarial stage (arrowhead) in the intestine (in) surrounded by erythrocytes (e). bar = 15 μm. D: Filarial stage in the haemocoel (hc) of the abdomen of E. horridus (square). E: Close up of filarial stage. mp = mouthparts, mo = mouth, cu = cuticula. A–C: Haematoxylin - Eosin stain, D: Giemsa stain.

Fig. 3. A in Heartworm and seal louse: Trends in prevalence, characterisation of impact and transmission pathways in a unique parasite assembly on seals in the North and Baltic Sea

Fig. 3. A: Prevalence of A. spirocauda and E. horridus in harbour seals in the North and Baltic Sea from 1996 to 2021, data from 1996 to 2013 according to Lehnert et al. (2016). B: Prevalence of A. spirocauda and E. horridus in harbour seals during the seasons in the North and Baltic Sea from 2014 to 2021.

Fig. 2 in Heartworm and seal louse: Trends in prevalence, characterisation of impact and transmission pathways in a unique parasite assembly on seals in the North and Baltic Sea

Fig. 2. Sampling routine of E. horridus infected seal skin for histological and bacteriological examinations. A: Mild E. horridus infection of a harbour seal yearling, asterisk pointing at E. horridus. B: Close up of E. horridus C: Removing of E. horridus D: Cutting and removing of the infected skin with a sterile forceps for further investigations. Scale bars: A-D 1 cm.

Fig. 1 in Heartworm and seal louse: Trends in prevalence, characterisation of impact and transmission pathways in a unique parasite assembly on seals in the North and Baltic Sea

Fig. 1. Levels of infection with E. horridus in P. vitulina. A: Mild E. horridus infection of a harbour seal yearling, asterisk pointing at E. horridus B: Close up of E. horridus in the head area of a harbour seal C: Severe E. horridus infection of a harbour seal D: Close up of severe E. horridus infection. Scale bars: A-D 1 cm.

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Fig. 4 in Confirmation of a unique species of Giardia, parasitic in the quenda (Isoodon obesulus)

Fig. 4. Phylogenetic relationships of Giardia peramelis isolates obtained in this study (quenda QBN13, QM22, QBY95) with published reference material available at the ITS1-5.8SITS2 locus. Evolutionary history inferred using the neighbour-joining method supported with bootstrap test of 1000 replicates (values> 50% shown). G. muris is used as the out group.

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Fig. 3 in Confirmation of a unique species of Giardia, parasitic in the quenda (Isoodon obesulus)

Fig. 3. Phylogenetic relationships of Giardia peramelis isolates obtained in this study (quenda QBN13, QM22, QBY95) with published reference material available at the 18s rRNA locus. Evolutionary history inferred using the neighbour-joining method supported with bootstrap test of 1000 replicates (values> 50% shown). G. muris is used as the out group.

Fig. 4. Maximum Likelihood phylogenetic tree generated using N in The African buffalo parasite Theileria. sp. (buffalo) can infect and immortalize cattle leukocytes and encodes divergent orthologues of Theileria parva antigen genes

Fig. 4. Maximum Likelihood phylogenetic tree generated using N-terminal sequences of T. sp. (buffalo) and T. parva PIM antigen genes. Maximum composite likelihood trees were constructed using 1000 bootstrap replicates as implemented in MEGA5; the optimal nucleotide substitution model was identified using data monkey. The tree constructed with RAxML (Stamatakis et al., 2014) using a GTR/G/I model with 100 bootstrap iterations.

Fig. 5 in The African buffalo parasite Theileria. sp. (buffalo) can infect and immortalize cattle leukocytes and encodes divergent orthologues of Theileria parva antigen genes

Fig. 5. Maximum Likelihood Phylogenetic trees illustrating the genetic relationships of T. parva CD8 T target antigen gene orthologues from T. sp. (buffalo). Panel (A) Tp6; Panel B Tp7: Panel C Tp8. Sequences were aligned and used to construct a maximum likelihood tree, at which the nodes were confirmed using 1000 bootstrap replications. The bootstrap values indicating the degree of support for each node are shown and also the GenBank accession numbers of the sequences. For Tp6, the tree was rooted using the prohibitin gene sequences present in Babesia bovis (XM001609045) and Theileria orientalis (AB161472). For Tp7, the tree was rooted using the putative Heat shock protein 90 gene sequences from Toxoplasma gondii (AY344115), Babesia bovis (AK442026) and Theileria annulata (XM_947380). For Tp8, the tree was rooted using an orthologue of Tp8 found in Theileria equi (CP001669).

Fig. 3 in The African buffalo parasite Theileria. sp. (buffalo) can infect and immortalize cattle leukocytes and encodes divergent orthologues of Theileria parva antigen genes

Fig. 3. PCR amplification of genes encoding Theileria parva antigens from Marula schizont-infected leukocyte cultures. Panel A, p104 primers; Panel B PIM, primers; Panel C p67 primers. The order of the schizont-infected lymphocyte samples is (1) N6; (2). N13; (3). N18; (4). N20; (5). N33; (6). N36; (7). N38; (8). N43; (9). N50; (10). N55; (11). N69; (12). N76; (13). N77, (14). N79; (15). N86, (16). N88; (17). N99; (18). N100; (19). N102; (20). N103; (21). N106; (22). N107.

Fig. 2 in The African buffalo parasite Theileria. sp. (buffalo) can infect and immortalize cattle leukocytes and encodes divergent orthologues of Theileria parva antigen genes

Fig. 2. Results of a semi-nested PCR assay used to amplify 18S ribosomal subunit DNA using primers specific for T. parva and T. sp. (buffalo). Samples are as follows: 1)N13 2)N18 3) N20 4)N33 5)N36 6) N43 7)N50 8)N55 9) N69 10)N76 11) N79 12) N86 13) N88 14) N99 15)N100 16) N102 17) N103 18)N107 19—21) T. parva clones 22—24) T. sp. (buffalo) clones (documented in Table 2).

Fig. 1 in The African buffalo parasite Theileria. sp. (buffalo) can infect and immortalize cattle leukocytes and encodes divergent orthologues of Theileria parva antigen genes

Fig. 1. Reverse line blot analysis of schizont cultures containing parasites isolated from Marula farm. The following species-specific oligonucleotide probes were used (a) T. annulata, (b) T. parva, (c) T. mutans, (d) T. velifera, (e) T. taurotragi, (f) T. buffeli, (g) T. sp. (buffalo). (h) B. bigemina, (i) B. bovis. The order of the experimental samples hybridized is DNA from cell culture isolates in lanes 1—22 was lane 1; (1) N6, (2) N13, (3) N18, (4) N20, (5) N33, (6) N36, (7) N38 (8) N43, (9) N50 (10) N55, (11) N69, (12) N76, (13) N77, (14) N79, (15) N88, (16) N99, (17) N100 (18) N103, (19) N106, (20) N107, (21) N86, (22) N102 and DNA extracted from whole cattle blood (23) N106 (24) N69 (25) N86.

Fig. 3 in Seasonal variation in the abundance and distribution of ticks that parasitize Microcebus griseorufus at the BezàMahafaly Special Reserve, Madagascar

Fig. 3. Possible life cycle of H. lemuris. Peak activity for larvae occurs in May, but larvae may be found feeding into June and October. Larvae attach to Microcebus hosts and after a blood meal, fall off and molt into nymphs. Nymphs are active and feed on Microcebus throughout the dry season and likely feed on other lemurs during part of the wet season. Adult-stage ticks remain active during the wet season, feeding on larger-bodied lemurs, such as L. catta, and P. verreauxi. Engorged females fall off and lay eggs in leaf litter. It is possible that all four stages can diapause if no suitable hosts or conditions are found (gray dotted line). Mice or rats may also serve as hosts to larvae during the dry season.

Fig. 1. Monthly averages for A in Seasonal variation in the abundance and distribution of ticks that parasitize Microcebus griseorufus at the BezàMahafaly Special Reserve, Madagascar

Fig. 1. Monthly averages for A) tick intensity on mouse lemurs as it compares to B) rainfall and C) temperature, during the year-long study season. Shaded area indicates months included in the dry season. Environmental data were collected daily.

Fig. 1. Circular Bayesian tree inferred from mtDNA cox-2 in Temporal stability of parasite distribution and genetic variability values of Contracaecum osculatum sp. D and C. osculatum sp. E (Nematoda: Anisakidae) from fish of the Ross Sea (Antarctica)

Fig. 1. Circular Bayesian tree inferred from mtDNA cox-2 sequences obtained from specimens of C. osculatum sp. D and C. osculatum sp. E analysed in the present study, based on Bayesian Inference (BI) method using MrBayes v3.2.2 (Ronquist et al., 2012). Evolutionary distance was estimated using the TrN + G (G = 0.60) substitution model as implemented in jModeltest (Posada, 2008), with the AIC approach (Posada and Buckley, 2004). Posterior probability values are the result of 1.000000 of runs and are reported at the nodes. The coloured icons correspond to the two species considered in this study (red = C. osculatum sp. D and blue = C. osculatum sp. E).

Fig. 2 in Temporal stability of parasite distribution and genetic variability values of Contracaecum osculatum sp. D and C. osculatum sp. E (Nematoda: Anisakidae) from fish of the Ross Sea (Antarctica)

Fig. 2. Schematic distribution of the fish species examined in the present study for larval of C. osculatum sp. D and C. osculatum sp. E, along the continental shelf of the Ross Sea coastal ecosystem. Arrows indicating preferred preys and the diet preference for each fish species are reported according to the literature (La Mesa et al., 2004). The represented pelagic organisms comprise species of euphausiids and fish juveniles, benthic and epibenthic organisms are polychaetes, amphipods, decapods and gastropods. A pie chart with the relative proportions of C. osculatum sp. D and C. osculatum sp. E is given for each fish species. Squares and circles represent the hypothetical distribution of C. osculatum sp. D and C. osculatum sp. E larvae in their intermediate hosts.

opencc-by-4.0Dec 2016View details →

Fig. 1 in Intestinal parasites of Tolypeutes matacus, the most frequently consumed armadillo in the Chaco region

Fig. 1. Distribution of helminths in the small intestine of Tolypeutes matacus from Chamical, La Rioja, Argentina. MOE: Moennigia virilis, ASP: Aspidodera spp., TRI: Trichohelix tuberculata, PTE: Pterygodermatites spp., DER: Delicata ransomi, CYC: Cyclobulura superinae, CES: Cestoda.

opencc-by-4.0Dec 2016View details →

Fig. 1 in Endoparasites of American marten (Martes americana): Review of the literature and parasite survey of reintroduced American marten in Michigan

Fig. 1. Hookworm egg from an American marten (Martes americana). There was no significant difference in the prevalence of hookworm eggs in faecal samples from American marten from the Upper Peninsula (n = 18) and Northern Lower Peninsula (n = 31) of Michigan, USA (prevalence 11.1% and 16.1%, respectively). American marten infected with hookworms were significantly more likely to be anaemic than noninfected American marten (P = 0.01) with an odds ratio of 8.75 (95% confidence interval: 1.4‾56.4).

Fig. 1 in Manifold habitat effects on the prevalence and diversity of avian blood parasites

Fig. 1. Diagram illustrating how conditions of the vector, parasite, host and habitat must all be permissive for pathogen transmission to occur. The outer layer depicts some factors that are currently causing rapid environmental change, which will affect host‾parasite dynamics.