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98 results for “parasite prevalence”
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>
Seasonal Effects in Gastrointestinal Parasite Prevalence, Richness and Intensity in Vervet Monkeys Living in a Semi-Arid Environment
<p>Data and R Notebook for Seasonal Effects in Gastrointestinal Parasite Prevalence, Richness and Intensity in Vervet Monkeys Living in a Semi-Arid Environment</p>
Fig. 3 in Prevalence and genetic diversity of haematozoa in South American waterfowl and evidence for intercontinental redistribution of parasites by migratory birds
Fig. 3. Bayesian phylogenetic tree of haematozoa mitochondrial DNA cytochrome b haplotypes obtained from infected waterfowl. Trees were rooted with mammalian Plasmodium outgroups. Node tips are labeled with parasite genus (Haem = Haemoproteus, Leuc = Leucocytozoon, and Plas = Plasmodium), followed by the lineage name, GenBank accession number for each sequence, host order (passerine/waterfowl), and the country/state from which the samples were collected. All haplotypes identified in this study are highlighted in red. Numbers on branches represent posterior probabilities from the analysis. Asterisks after node tip labels indicate sequences from our study that were identical to lineages previously found in non-waterfowl hosts. All reference sequences were obtained from the National Center for Biotechnology Information website.
Fig. 2. Minimum spanning network for haematozoa mitochondrial DNA cytochrome b in Prevalence and genetic diversity of haematozoa in South American waterfowl and evidence for intercontinental redistribution of parasites by migratory birds
Fig. 2. Minimum spanning network for haematozoa mitochondrial DNA cytochrome b haplotypes detected in South American waterfowl. Shaded circles represent unsampled nodes. All circles are drawn proportional to the frequency at which haplotypes were observed. Lines separating nodes are drawn to scale based on the number of nucleotide mutations, unless otherwise indicated by hash marks. Only haplotypes with a length of 358 bp or greater were included. Haplotype name abbreviations are as follows: Haem = Haemoproteus, Leuc = Leucocytozoon, and Plas = Plasmodium.
Fig. 1 in Prevalence and genetic diversity of haematozoa in South American waterfowl and evidence for intercontinental redistribution of parasites by migratory birds
Fig. 1. Map of sampling locations in Peru and Argentina. The number of waterfowl blood samples collected at each site is provided in parentheses.
Fig. 1 in High prevalence of haemosporidian parasites in Eurasian jays
Fig. 1 Median-joining network of mitochondrial cytochrome b lineages (476 bp, n = 60 sequences) of haemosporidian parasites found in Eurasian jays Garrulus glandarius. Circle size is proportional to the lineage frequency. Lineage names are noted at the associated circles together with one exemplary GenBank association number in parentheses. One hatch mark represents one mutation. Sample origins are represented by different colours, 'MalAvi' referring to sequences from other studies deposited in the MalAvi database (MalAvi 2023). Morphospecies names for GAGLA07 (H. homopicae) and TURDUS2 (H. minutus) are not provided within the figure
Figure 4 in Prevalence of Blastocystis sp. in Morocco: Comparative assessment of three diagnostic methods and characterization of parasite forms in Jones' culture medium
Figure 4. Observation under the light microscope (×400) of different forms of cultured Blastocystis sp. in Jones' medium. Panel A: Different sizes of the vacuolar form. Panel B: Granular form (blue arrow) and cystic form (black arrow). Panel C: Granular form (blue arrow) and vacuolar form (red arrow). Panel D: Granular form (black arrow) and vacuolar form (red arrow) stained with methylene blue. Panel E: Illustrates the process of transformation of vacuolar cells into multivacuolar forms in culture, showing the division of the central vacuole into smaller vacuoles. Panel D: Different aspects of the amoeboid form with the presence of a single or several pseudopodia (red arrow).
Figure 5 in Prevalence of Blastocystis sp. in Morocco: Comparative assessment of three diagnostic methods and characterization of parasite forms in Jones' culture medium
Figure 5. Representative gel image of PCR products from Blastocystis isolates. Lanes 1 to 13: Blastocystis isolates; lane NC: negative control; lane PC: positive control; DNA ladder – 50 bp.
Figure 3 in Prevalence of Blastocystis sp. in Morocco: Comparative assessment of three diagnostic methods and characterization of parasite forms in Jones' culture medium
Figure 3. Various forms of Blastocystis sp. were observed under the light microscope during the direct examination of stool specimens. Panel A: Vacuolar form (red arrow) and cyst form (black arrow) of Blastocystis in an unstained wet mount. N: Nuclei situated at the periphery of the organism. C. b: Central body. Panels B, C, and D: Vacuolar form (red arrow), Granular form (blue arrow), and cyst form (black arrow) stained with Lugol's iodine (×400).
Figure 2 in Prevalence of Blastocystis sp. in Morocco: Comparative assessment of three diagnostic methods and characterization of parasite forms in Jones' culture medium
Figure 2. Occurrence of Blastocystis sp. infection on its own or in conjunction with other protozoan species.
Fig. 5 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. 5. Scanning electron microscopic images of E. horridus attached to seal skin and fur. A: E. horridus utilizing a hair follicle of harbour seal skin. B: E. horridus attached to seal hair with the head pointing towards seal skin. C: E. horridus with six claws attached to seal fur. D: Close up of an unattached claw of E. horridus. Asterisks positioned on nits of E. horridus. Sale bars: A 200 μm, B 400 μm, C 400 μm, D 100 μm.
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.
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. 1. A in Seasonality, richness and prevalence of intestinal parasites of three neotropical primates (Alouatta seniculus, Ateles hybridus and Cebus versicolor) in a fragmented forest in Colombia
Fig. 1. A. Trichuris sp., B. Oxyuridae, C. Ancylostomatidae, D. Strongyloides sp. (larva), E. Ascarididae, F. Gnathostomatidae, G. Trichostrongylidae, H-I. Trematodes, J. Entamoeba sp. (cyst), K. Acanthocephala, L. Balantidiidae.
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Allen Brain Atlas
Allen Brain Atlas is an Allen Institute collection of brain map atlases, datasets, APIs, and analysis tools covering mouse, human, and non-human primate brain resources.
Annotated Behaviour and Observability Dataset (ABODe)
ABODe is a University of Edinburgh DataShare dataset for behavior classification in group-housed mice using home-cage video, identities, bounding boxes, ground-plate positions, and annotator labels.
DANDI Archive for NWB datasets
DANDI is a BRAIN Initiative archive for publishing and sharing neurophysiology data, including electrophysiology, optophysiology, and behavioral data packaged as NWB and related standards.
International Brain Laboratory public data
The International Brain Laboratory public data releases expose standardized mouse decision-making experiments, including Neuropixels recordings, widefield calcium imaging, behavior, and session metadata accessed through the ONE API.
OpenNeuro
OpenNeuro is a free, open platform for sharing neuroimaging datasets, with public search, dataset pages, and download paths for web, S3, DataLad, and the OpenNeuro CLI.