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6,859 results for “parasitism”
Figure 8. Eutarsopolipus pulcher n in Sheltered life beneath elytra: three new species of Eutarsopolipus (Acari, Heterostigmatina, Podapolipidae) parasitizing Australian ground beetles
Figure 8. Eutarsopolipus pulcher n. sp. (larval female). (a) Body dorsum; (b) body venter; (c) right leg I; (d) right leg II; (e) right leg III. All legs in dorsal view.
Figure 2. Eutarsopolipus paryavae n in Sheltered life beneath elytra: three new species of Eutarsopolipus (Acari, Heterostigmatina, Podapolipidae) parasitizing Australian ground beetles
Figure 2. Eutarsopolipus paryavae n. sp. (male). (a) Body dorsum; (b) body venter; (c) right leg I; (d) right leg II; (e) right leg III. All legs in dorsal view.
Figure 5. Eutarsopolipus pulcher n in Sheltered life beneath elytra: three new species of Eutarsopolipus (Acari, Heterostigmatina, Podapolipidae) parasitizing Australian ground beetles
Figure 5. Eutarsopolipus pulcher n. sp. (adult female). (a) Body dorsum; (b) body venter; (c) right leg I; (d) ventral view of tarsus I; (e) right leg II; (f) right leg III. All legs in dorsal view.
Figure 4 in Sheltered life beneath elytra: three new species of Eutarsopolipus (Acari, Heterostigmatina, Podapolipidae) parasitizing Australian ground beetles
Figure 4. Key to the world species of Eutarsopolipus in the pterostichi group (based on adult females).
Figure 1. Eutarsopolipus paryavae n in Sheltered life beneath elytra: three new species of Eutarsopolipus (Acari, Heterostigmatina, Podapolipidae) parasitizing Australian ground beetles
Figure 1. Eutarsopolipus paryavae n. sp. (adult female). (a) Body dorsum; (b) body venter; (c) right leg I; (d) ventral view of tarsus I; (e) right leg II; (f) right leg III. All legs in dorsal view.
Figure 1 in Neohexostoma gymnosardae n. sp. (Monogenea, Hexostomatidae), a gill parasite of Gymnosarda unicolor (Valenciennes) (Teleostei, Scombridae) in the South China Sea
Figure 1. Maximum likelihood tree based on an analysis of 28S rDNA sequences. Bootstrap percentages with 1000 replicates.
Figure 7. Eutarsopolipus pulcher n in Sheltered life beneath elytra: three new species of Eutarsopolipus (Acari, Heterostigmatina, Podapolipidae) parasitizing Australian ground beetles
Figure 7. Eutarsopolipus pulcher n. sp. (male). (a) Body dorsum; (b) body venter; (c) right leg I; (d) right leg II; (e) right leg III. All legs in dorsal view.
Figure 3 in Neohexostoma gymnosardae n. sp. (Monogenea, Hexostomatidae), a gill parasite of Gymnosarda unicolor (Valenciennes) (Teleostei, Scombridae) in the South China Sea
Figure 3. Photographs of Neohexostoma gymnosardae n. sp. (A) Holotype, whole worm (ventral view); (B) Male copulatory organ; (C) Vaginal spines; (D) Haptor; (E)–(F) Eggs; (G) Clamp with sclerites in lateral view. (B)–(G) are paratypes.
Figure 3. Eutarsopolipus paryavae n in Sheltered life beneath elytra: three new species of Eutarsopolipus (Acari, Heterostigmatina, Podapolipidae) parasitizing Australian ground beetles
Figure 3. Eutarsopolipus paryavae n. sp. (larval female). (a) Body dorsum; (b) body venter; (c) right leg I; (d) right leg II; (e) right leg III. All legs in dorsal view.
Figure 6 in Sheltered life beneath elytra: three new species of Eutarsopolipus (Acari, Heterostigmatina, Podapolipidae) parasitizing Australian ground beetles
Figure 6. Phase-contrast micrograph of tarsus III in Eutarsopolipus pulcher n. sp. (adult female) representing modified trifurcate seta uļ and spur-like seta tcļļ.
Figure 10. Eutarsopolipus chlaenii n in Sheltered life beneath elytra: three new species of Eutarsopolipus (Acari, Heterostigmatina, Podapolipidae) parasitizing Australian ground beetles
Figure 10. Eutarsopolipus chlaenii n. sp. (adult female). (a) Body dorsum; (b) body venter; (c) right leg I; (d) ventral view of tarsus I; (e) right leg II; (f) right leg III. All legs in dorsal view.
Figure 12 in Sheltered life beneath elytra: three new species of Eutarsopolipus (Acari, Heterostigmatina, Podapolipidae) parasitizing Australian ground beetles
Figure 12. Key to closely related species of myzus group (based on adult females) possessing similar characters including short cheliceral stylets (<35 µm long).
Figure 3 in Taxonomic revision of the Nippostrongylinae (Nematoda, Heligmonellidae) parasites of Muridae from the Australasian region. The genus Odilia Durette-Desset, 1973
Figure 3. Synlophes of the genera treated in this work. Other species. (A) within anterior part of body, Chisholmia mawsonae n. comb., male. (B–H) at mid-body: (B) Hasegawanema mallomyos n. comb., female. (C) Hasegawanema maxomyos n. comb., male. (D) Hasegawanema moatense n. comb., female. (E) Hasegawanema sulawesiense n. comb., female. (F) Hughjonestrongylus implexus n. comb., female. (G) Parasabanema praeputiale n. comb., female. (H) Equilophos similis n. comb., male. Abbreviations: 1, 1', 2', 3': ridges 1, 1', 2', 3'; c: careen; G: gap; lr: left ridge; rr: right ridge. A, modified from [5]. B, modified from [12]. C–E, modified from [16]. F, modified from [21]. G, modified from [11]. H, modified from [22]. Scale-bars: 50 mm.
Figure 2 in Taxonomic revision of the Nippostrongylinae (Nematoda, Heligmonellidae) parasites of Muridae from the Australasian region. The genus Odilia Durette-Desset, 1973
Figure 2. Synlophes at mid-body of the genera treated in this work. Type-species. (A) Odilia (O. mackerrasae), male. (B) Hasegawanema n. gen. (Hasegawanema mamasaense n. comb), female. C, Hughjonestrongylus (H. ennisae), female. (D) Chisholmia n. gen. (Chisholmia bainae n. comb.), male. (E) Lesleyella n. gen. (Lesleyella wauensis n. comb.), female. (F) Sanduanensis n. gen. (Sanduanensis dividua n. comb.), female. (G) Parasabanema (P. szalayi), male. (H) Equilophos n. gen. (Equilophos polyrhabdote n. comb.), female. Abbreviations: 1, 1', 2', 3': ridges 1, 1', 2', 3'; AO: axis of orientation of the ridges; c: careen; G: gap; lr: left ridge; n: last dorsal ridge; n': last ventral ridge; rr: right ridge. A, H, modified from [5]. B, modified from [16]. C, G, modified from [26]. D, modified from [1]. E, modified from [20]. F, modified from [25]. Scale-bars: 50 mm.
Figure 1 in Taxonomic revision of the Nippostrongylinae (Nematoda, Heligmonellidae) parasites of Muridae from the Australasian region. The genus Odilia Durette-Desset, 1973
Figure 1. (A–E) Distribution of ridges around the body circumference. (A) According to the sagittal axis, ridges are named left ridges and right ridges. (B) According to the frontal axis, ridges are named dorsal ridges and ventral ridges. (C) Division into quadrants resulting from the intersection of sagittal and frontal axes. Ridges are named left-dorsal, right-dorsal, right-ventral, and left-ventral ridges. (D) Division into quadrants resulting from the intersection of the axes ''right-ventral/left-dorsal'' and ''left-ventral/right-dorsal''. Ridges are named mid-dorsal, mid-right, mid-ventral, and mid-left ridges. (E) Division into octants. Ridges are named dorsal-right-dorsal, right-right-dorsal, right-rightventral, ventral-right-ventral, ventral-left-ventral, left-left-ventral, left-left-dorsal, and dorsal-left-dorsal ridges. Abbreviations: D: dorsal side; FA: frontal axis; L: left side; LV/RD: left-ventral/right-dorsal axis; R: right side; RV/LD: right-ventral/left-dorsal axis; SA: sagittal axis; V: ventral side; dr: dorsal ridges; d-l-dr: dorsal-left-dorsal ridges; d-r-dr: dorsal-right-dorsal ridges; lr: left ridges; l-dr: left-dorsal ridges; l-vr: left-ventral ridges; l-l-dr: left-left-dorsal ridges; l-l-vr: left-left-ventral ridges; m-dr: mid-dorsal ridges; m-lr: mid-left ridges, m-rr: mid-right ridges, m-vr: mid-ventral ridges; r-dr: right-dorsal ridges; rr: right ridges; r-vr: right-ventral ridges; r-r-dr: right-right-dorsal ridges; r-r-vr: right-right-ventral ridges; vr: ventral ridges, v-l-vr: ventral-left-ventral ridges; v-r-vr: ventral-right-ventral ridges.
Figure 4 in Application of the NucliSENS easyMAG system for nucleic acid extraction: optimization of DNA extraction for molecular diagnosis of parasitic and fungal diseases
Figure 4. Identification of PCR inhibitors in 18 biological samples positive for Aspergillus. Graph A: Ct values obtained from pure and diluted DNA samples (dilution rate 1/20). Graph B: Ct values obtained with 20 copies of a plasmid DNA systematically added to the same biological samples (undiluted and diluted) and a negative control sample (NC).
Figure 2 in Application of the NucliSENS easyMAG system for nucleic acid extraction: optimization of DNA extraction for molecular diagnosis of parasitic and fungal diseases
Figure 2. Influence of proteinase K digestion (56 °C overnight) on DNA extraction. Graph A shows the Ct values obtained by quantifying THP1 cell DNA derived from direct extraction with the NucliSENS easyMAG system and extraction performed on the same quantity of cells following overnight (ON) digestion with Proteinase K. Graph B shows Leishmania quantification after extraction with the NucliSENS easyMAG system both with and without PK and quantification after extraction using a QIAamp DNA Mini kit after ON digestion with PK.
Figure 7 in Application of the NucliSENS easyMAG system for nucleic acid extraction: optimization of DNA extraction for molecular diagnosis of parasitic and fungal diseases
Figure 7. Variation of the ratio between kinetoplastic DNA and nuclear DNA extraction with various Leishmania quantities in the presence of 103 THP1 cells.
Figure 5 in Application of the NucliSENS easyMAG system for nucleic acid extraction: optimization of DNA extraction for molecular diagnosis of parasitic and fungal diseases
Figure 5. Yield of DNA extraction from Leishmania and THP1 cells using the NucliSENS easyMAG system.
Figure 3 in Application of the NucliSENS easyMAG system for nucleic acid extraction: optimization of DNA extraction for molecular diagnosis of parasitic and fungal diseases
Figure 3. Results of the extraction experiments performed on yeast (Candida albicans) and filamentous fungi (Aspergillus fumigatus). A presents the kinetics of the extraction process after vortexing and glass-bead treatment. B shows the differences in DNA quantity obtained from fungal cells using the FastPrep system (with) compared to the same process without grinding.
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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.