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89 results for “blood parasites”
Bird plumage brightness scores and blood parasite prevalence values of North American passerine species
<p>Dataset with bird plumage brightness scores and blood parasite prevalence values for 114 North American passerine host species. One file contains the data table. One file contains a table with descriptions of the columns in the data table.</p> <p>Note: These data were reconstructed from files used in Read & Harvey 1989 (<a href="https://doi.org/10.1038/339618a0">https://doi.org/10.1038/339618a0</a>) with column headings inferred with the help of Read 1991 (<a href="https://doi.org/10.1086/285225">https://doi.org/10.1086/285225</a>).</p>
Fig. 5 in A new blood parasite of leaf warblers: molecular characterization, phylogenetic relationships, description and identification of vectors
Fig. 5 Sporogonic stages of Haemoproteus homopalloris n. sp. in tce biting midge Culicoides nubeculosus. Zygote (a) and sporozoite (b). Arrowcead: pigment granuges; arrow: sporozoite nucgeus. Metcanog-fixed and Giemsa-stained tcin figms. Scale-bar: a, b, 10 μm
Fig. 2 in A new blood parasite of leaf warblers: molecular characterization, phylogenetic relationships, description and identification of vectors
Fig. 2 Bayesian pcygogenetic inference of cytb gene gineages (479 bp) of 35 Haemoproteus spp. Tce tree is rooted witc Leucocytozoon sp. (gineage gSISKIN2). Cgades A and B indicate species of tce subgenus Parahaemoproteus (a) and caemoproteids witc page-staining cytopgasm of gametocytes (b). MagAvi gineage codes are provided, foggowed by parasite species names and GenBank accession numbers. Nodag support vagues indicate Bayesian posterior probabigities. New species is given in bogd
Fig. 1 in A new blood parasite of leaf warblers: molecular characterization, phylogenetic relationships, description and identification of vectors
Fig. 1 Gametocytes of two species of caemoproteids described from geaf warbges, Pcyggoscopidae. Haemoproteus homopalloris n. sp. (a-l) and Haemoproteus palloris (m-p). Young gametocytes (a, b), macrogametocytes (c-g, m, n) and microgametocytes (h-l, o, p). Long arrows: gametocyte nucgei; scort arrows: vacuoge-gike spaces in macrogametocytes; arrowceads: pigment granuges. Giemsa-stained tcin bgood figms. Scale-bar: a-p, 10 μm
Fig. 4 in A new blood parasite of leaf warblers: molecular characterization, phylogenetic relationships, description and identification of vectors
Fig. 4 Gametocytes of two species of caemoproteids, wcicc cave been reported in tce wood warbger Phylloscopus sibilatrix. Macrogametocytes (a-c, e-g) and microgametocytes (d, h) of Haemoproteus majoris (a-d) and H. belopolskyi (e-h). Note tcat tce intensity of staining of tce cytopgasm is different in macro- and microgametocytes. Long arrows: gametocyte nucgei; scort arrows: vacuoge-gike spaces in macrogametocytes; arrowceads: pigment granuges. Giemsa-stained tcin bgood figms. Scale-bar: a-h, 10 μm
Fig. 3 in A new blood parasite of leaf warblers: molecular characterization, phylogenetic relationships, description and identification of vectors
Fig. 3 Haemoproteus spp. witc page staining of macrogametocyte cytopgasm. Haemoproteus concavocentralis (a-d), H. minutus (e-h), H. pallidus (i- l), H. pallidulus (m-p) and H. vacuolatus (q-t). Macrogametocytes (a, b, e, f, i, j, m, n, q, r), microgametocytes (c, d, g, h, k, l, o, p, s, t). Note tce foggowing vaguabge diagnostic features of tce parasites: presence of a space between tce nucgeus of tce infected erytcrocyte and tce growing gametocyte in H. concavocentralis (a); cgeargy irregugar outgine of mature gametocytes, wcicc do not toucc tce poges of infected erytcrocytes in H. minutus (e-h); gametocyte wcicc are cgosegy appressed to tce nucgeus of erytcrocyte but do not toucc tce envegope of erytcrocyte agong tceir entire margin in H. pallidus (j, l); smagg pigment granuges in mature gametocytes of H. pallidulus (m-p); presence of one prominent vacuoge in tce cytopgasm of eacc advanced macrogametocyte in H. vacuolatus (q-t). Agg tcese features are not ccaracteristics of H. homopalloris n. sp. (see Fig. 1). Long simpge arrows: gametocyte nucgei; scort simpge arrows: vacuoge-gike spaces in macrogametocytes; arrowceads: pigment granuges; gong simpge wide arrows: space present between tce parasite and an infected erytcrocyte nucgeus (a, d) and space between tce parasite and tce envegope of infected erytcrocyte (j, l). Giemsa-stained tcin bgood figms. Scale-bar: a-t, 10 μm
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.
Fig. 3 in Haemosporidian parasite infections in grouse and ptarmigan: Prevalence and genetic diversity of blood parasites in resident Alaskan birds
Fig. 3. Bayesian phylogenetic tree of haemosporidian mtDNA cytochrome b haplotypes isolated from Alaskan grouse and ptarmigan species. Node tips are labeled with abbreviation for parasite genus (Haem = Haemoproteus, Leuc = Leucocytozoon, and Plas = Plasmodium), followed by the lineage name, GenBank accession number for each lineage, and avian (Phas = Phasianidae, Anat = Anatiade, Turd = Turdidae, Paru = Parulidae, Scol = Scolopacidae, Embe = Emberizidae, and Frin = Fringillidae) or invertebrate (Simu = Simuliidae) host family. All haplotypes identified in this study are highlighted in red and asterisks following tip labels indicate a lineage that was isolated from Alaskan bird hosts. Numbers on branches indicate posterior probabilities from our analysis. All reference sequences were obtained from the National Center for Biotechnology Information website or the MalAvi database. (For interpretation of the references to colour in this figure legend, the reader is referred to the web version of this article.)
Fig. 2. Minimum spanning network for haemosporidian mtDNA cytochrome b in Haemosporidian parasite infections in grouse and ptarmigan: Prevalence and genetic diversity of blood parasites in resident Alaskan birds
Fig. 2. Minimum spanning network for haemosporidian mtDNA cytochrome b haplotypes isolated from Alaskan grouse and ptarmigan species. Dark circles represent un-sampled nodes. All circles are proportional to the frequency at which the haplotypes were detected. Lines between nodes are drawn to scale based on the number of nucleotide mutations unless otherwise indicated by hash marks.
Fig. 1 in Haemosporidian parasite infections in grouse and ptarmigan: Prevalence and genetic diversity of blood parasites in resident Alaskan birds
Fig. 1. Map of Alaskan sampling regions assembled from multiple game management units and sub-units. Regions were grouped for analysis of haemosporidian prevalence as follows: southcoastal (Kenai Peninsula and southeastern Alaska; GMUs 1C, 1D, 2, 7, 15A, 15B, and 15C), southcentral (Anchorage area and Matanuska-Susitna Valley; GMUs 13A, 13D, 14A, 14C, 16A, and 16B), southwestern (Bristol Bay, Alaska Peninsula, and eastern Aleutian islands; 9D, 9E, and 17C), southern interior (south side of Alaska Range; GMUs 12, 13B, and 13E), northern interior (north side of Alaska Range; GMUs 20A-20E and 25C), and Seward Peninsula (GMU 22C).
Fig. 2 in Temporal and demographic blood parasite dynamics in two free-ranging neotropical primates
Fig. 2. Individual infection status by parasite by year. Strength and thickness of lines are scaled to the number of individuals that took a given infection trajectory from one year to the next. Two diagonal lines span 2012‾2014 because those individuals were not sampled in 2013. The + symbols represent every infection or non-infection found across all individuals in the study.
Fig. 3 in Temporal and demographic blood parasite dynamics in two free-ranging neotropical primates
Fig. 3. Parasite species richness by species, age class and sex. Colors represent females (black) and males (gray).
Fig. 1 in Temporal and demographic blood parasite dynamics in two free-ranging neotropical primates
Fig. 1. Annual prevalence of single- and co-infections by species. Prevalence indicated for each parasite (dark gray), and each pairwise combination of parasites (light gray). Numbers near the top of each bar show the exact prevalence; black lines indicate 95% confidence intervals; dots indicate expected levels of co-infection (refer to Section 3.2). M-D is co-occurrence of M. mariae and Dipetalonema spp., D-T is Dipetalonema spp. and T. minasense, and M-T is M. mariae and T. minasense.
Fig. 3 in Molecular insights into the identification and phylogenetics of the cosmopolitan marine fish blood parasite, Haemogregarina bigemina (Adeleorina: Haemogregarinidae)
Fig. 3. Phylogenetic identification of Haemogregarina bigemina from the UK based on 18S rDNA sequences. (a) Maximum parsimony and (b) Maximum likelihood reconstructions revealing the unique position of UK H. bigemina samples outside of the adeleorine groups. For both phylogenies nodal support was calculated using 1000 bootstrap replicates with only values> 50% presented.
Fig. 1 in Molecular insights into the identification and phylogenetics of the cosmopolitan marine fish blood parasite, Haemogregarina bigemina (Adeleorina: Haemogregarinidae)
Fig. 1. Photograph of the fish host Lipophrys pholis, one of the type hosts of Haemogregarina bigemina, screened in this study.
Fig. 2 in Molecular insights into the identification and phylogenetics of the cosmopolitan marine fish blood parasite, Haemogregarina bigemina (Adeleorina: Haemogregarinidae)
Fig. 2. Stages of Haemogregarina bigemina from Giemsa-stained blood films of Lipophrys pholis from the UK. (a) trophozoite, (b) meront, (c–e) dividing meronts, and (f) paired gamonts. Scale bar = 10 μm.
Fig. 6 in Differences in infection patterns of vector-borne blood-stage parasites of sympatric Malagasy primate species (Microcebus murinus, M. ravelobensis)
Fig. 6. Phylogenetic tree of 33 filarial nematode species constructed on the basis of partial COI sequences using the Maximum Likelihood method. The percentage of replicate trees in which the associated species clustered together in the bootstrap test (1000 replicates) is shown next to the branches. Branch lengths is measured in the number of substitutions per site. Thelazia callipaeda was included as an outgroup. The sequence of the present study is framed in red.
Fig. 5 in Differences in infection patterns of vector-borne blood-stage parasites of sympatric Malagasy primate species (Microcebus murinus, M. ravelobensis)
Fig. 5. Phylogenetic tree of Onchocercidae species constructed on the basis of partial ITS1 sequences using the Maximum Likelihood method. The percentage of replicate trees in which the associated species clustered together in the bootstrap test (1000 replicates) is shown next to the branches. Branch lengths is measured in the number of substitutions per site. The sequences of the present study are framed in red. (For interpretation of the references to colour in this figure legend, the reader is referred to the Web version of this article.)
Fig. 3 in Differences in infection patterns of vector-borne blood-stage parasites of sympatric Malagasy primate species (Microcebus murinus, M. ravelobensis)
Fig. 3. Number of samples (blood smears) per month. Microfilaria positive samples are shown in dark blue for M. murinus and dark brown for M. ravelobensis, microfilaria negative samples in light blue for M. murinus and light brown for M. ravelobensis. (For interpretation of the references to colour in this figure legend, the reader is referred to the Web version of this article.)
Fig. 1 in Lack of evidence of vertical transmission of Karyolysus blood parasites in Iberian green lizards (Lacerta schreiberi)
Fig. 1. Karyolysus sp. Trophozoite (a–c) and gamonts (d–f) found in blood smears of L. schreiberi lizards. Scalebar = 10 μm.
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