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Fig. 18. The parasitic earwig Hemimerus talpoides Walker, 1871 in The Earwig Collection (Dermaptera) of the Manchester Museum, UK, with a complete type catalogue
Fig. 18. The parasitic earwig Hemimerus talpoides Walker, 1871, the Manchester Museum. Scale bar = 1 cm.
Figs 39–41. 39 in Seven new species of cestode parasites (Neodermata, Platyhelminthes) from Australian birds
Figs 39–41. 39. Dilepididae genus sp. Hooks. 40. Passerilepis sp. Hooks. 41. Cracticotaenia fieldingi (Maplestone & Southwell, 1923) Hooks. Scale bars: 39–41 = 20 µm.
Figs 23–28 in Seven new species of cestode parasites (Neodermata, Platyhelminthes) from Australian birds
Figs 23–28. Dictymetra longiuncinata sp. nov. 23. Scolex. 24. Hooks. 25. Mature proglottis. 26. Terminal genitalia. 27. Cirrus armament. 28. Eggs. Scale bars: 23 = 100 µm; 24, 26 = 50 µm; 25 = 200 µm; 27 = 10 µm; 28 = 20 µm.
Figs 7–11 in Seven new species of cestode parasites (Neodermata, Platyhelminthes) from Australian birds
Figs 7–11. Spiniglans beveridgei sp. nov. 7. Scolex. 8. Hooks. 9. Mature proglottis. 10. Terminal genitalia. 11. Pregravid proglottis with young reticular uterus. Scale bars: 7 = 100 µm; 8 = 20 µm; 9 = 200 µm; 10 = 50 µm; 11 = 500 µm.
Figs 1–6 in Seven new species of cestode parasites (Neodermata, Platyhelminthes) from Australian birds
Figs 1–6. Sobolevitaenia whittingtoni sp. nov. 1. Scolex. 2. Sucker armament. 3. Hooks. 4. Mature proglottis. 5. Terminal genitalia. 6. Egg. Scale bars: 1 = 100 µm; 2, 5 = 50 µm; 3, 6 = 20 µm; 4 = 250 µm.
Figs 34–38 in Seven new species of cestode parasites (Neodermata, Platyhelminthes) from Australian birds
Figs 34–38. Cracticotaenia adelaidae sp. nov. 34. Scolex. 35. Detail of hooks crowns. 36. Hooks: anterior (A) and posterior (B). 37. Mature proglottis. 38. Eggs. Scale bars: 34 = 100 µm; 35 = 10 µm; 19 = 500 µm; 36, 38 = 20 µm; 37 = 250 µm.
Figs 29–33 in Seven new species of cestode parasites (Neodermata, Platyhelminthes) from Australian birds
Figs 29–33. Notopentorchis musealis sp. nov. 29. Scolex. 30. Hooks: A, C. Anterior hooks, B. Posterior hook. 31. Mature proglottis. 32. Postmature proglottis. 33. Pregravid proglottis. Scale bars: 29, 31–33 = 100 µm; 30 = 20 µm.
Figs 12–16 in Seven new species of cestode parasites (Neodermata, Platyhelminthes) from Australian birds
Figs 12–16. Monopylidium australiense sp. nov. 12. Scolex. 13. Hooks. 14. Mature proglottis. 15. Terminal genitalia. 16. Eggs. Scale bars: 12, 14 = 100 µm; 13, 16 = 20 µm; 15 = 50 µm.
Figs 17–22 in Seven new species of cestode parasites (Neodermata, Platyhelminthes) from Australian birds
Figs 17–22. Dictymetra gerganae sp. nov. 17. Scolex. 18. Hooks. 19. Mature proglottis. 20. Terminal genitalia. 21. Terminal genitalia tuft. 22. Egg. Scale bars: 17 = 100 µm; 18, 20, 21 = 50 µm; 19 = 500 µm; 22 = 20 µm.
Fig. 1 in Are juveniles of the enigmatic deep-sea nematode Rhaptothyreus (Rhaptothyreida: Rhaptothyreidae) parasitic?
Fig. 1. Rhaptothyreus typicus Hope & Murphy, 1969. Line drawings. A. Lateral view of male head. B. Lateral view of moulting juvenile head, with details of surface striations on outer cuticle and outline of amphid under moulting cuticle. C. Anterior body region of male. D. Anterior body region of juvenile. E. Lateral view of male mid-body region. F. Posterior body region of male. G. Posterior body region of juvenile. Arrows show the position of the chord (c) and turgescent cells (t). Scale bar: A–B = 50 µm, C–D = 75 µm, E = 60 µm, F–G = 70 µm.
Fig. 4 in Are juveniles of the enigmatic deep-sea nematode Rhaptothyreus (Rhaptothyreida: Rhaptothyreidae) parasitic?
Fig. 4. Rhaptothyreus typicus Hope & Murphy, 1969. Scanning electron micrographs. A-B. Male head. C. Juvenile head. D. Male posterior body region. Scale bar: A = 20 µm, B = 8 µm, C = 12 µm, D = 16 µm.
Fig. 3 in Are juveniles of the enigmatic deep-sea nematode Rhaptothyreus (Rhaptothyreida: Rhaptothyreidae) parasitic?
Fig. 3. Rhaptothyreus typicus Hope & Murphy, 1969. Light micrographs (moulting juvenile). A. Lateral view of head, showing stylet-like structure. B. Sharp transition between anterior (left) and posterior trophosome (right). C. Lateral chord, mid-body region. D. Posterior body region. Scale bar: A = 20 µm, B = 40 µm, C = 28 µm, D = 25 µm.
Fig. 2 in Are juveniles of the enigmatic deep-sea nematode Rhaptothyreus (Rhaptothyreida: Rhaptothyreidae) parasitic?
Fig. 2. Rhaptothyreus typicus Hope & Murphy, 1969. Light micrographs (♂). A. Lateral view of head showing amphid. B. Mid-body region showing cuticle, turgescent cells and portion of anterior trophosome with rod-shaped structures. C. Cells of lateral chord with clear, round inclusions. D. Posterior body region. Arrows show the position of the two small ducts apparently joining just prior to the cloacal opening. Scale bar: A, C = 20 µm, B = 18 µm, C = 30 µm.
Figs 27–30 in Cestode parasites (Neodermata, Platyhelminthes) from Malaysian birds, with description of five new species
Figs 27–30. Raillietina mahnerti sp. nov. 27. a) Scolex. b) Sucker hooklets. c) Rostellar hook. 28. Mature proglottis, ventral view. 29. Copulatory organs. 30. Gravid proglottis with egg capsules. Scale bars: 27a, 28, 30 = 100 µm; 27b = 20 µm; 27c = 10 µm; 29 = 50 µm.
Figs 9–13 in Gallancyra gen. nov. (Phthiraptera: Ischnocera), with an overview of the geographical distribution of chewing lice parasitizing chicken
Figs 9–13. Gallancyra dentata (Sugimoto, 1934) gen. et comb. nov. ex Gallus gallus (Linnaeus, 1758) (NHMUK010682393). 9. Male head, dorsal and ventral views. 10. Female antenna, ventral view. 11. Male genitalia, dorsal view. 12. Male paramere, dorsal view. 13. Male mesosome, ventral view. Female antenna at same scale as male head. Abbreviations: ads = anterior dorsal seta; as2 = anterior seta 2; pst1–2 = parameral setae 1–2. All genitalic component drawn at same scale.
Fig. 2 in Gallancyra gen. nov. (Phthiraptera: Ischnocera), with an overview of the geographical distribution of chewing lice parasitizing chicken
Fig. 2. Geographical distribution of four species of ischnoceran chewing lice parasitizing wild and domestic chicken (Gallus spp.). Each circle is divided into four sectors, representing the four louse species: upper left = Lipeurus caponis (Linnaeus, 1758); upper right = Lipeurus tropicalis Peters, 1931; lower left = Cuclotogaster heterographus (Nitzsch, 1866); lower right = Lagopoecus sinensis (Sugimoto, 1930). Black sectors indicate that this louse species is known from this country, whereas hollow sectors indicate that we have found no published records of this species in this country. Presence of the four species of chewing lice in a country is based on the reports summarized in Table 1.
Data from: Comparing methods for mapping global parasite diversity
Aim Parasites are a major component of global ecosystems, yet spatial variation in parasite diversity is poorly known, largely because their occurrence data are limited and thus difficult to interpret. Using a recently compiled database of parasite occurrences, we compare different models which we use to infer parasite geographic ranges and parasite species richness across the globe. Innovation To date, most studies exploring spatial patterns of parasite diversity assumed, with little validation, that the geographic range of a parasite species can be represented by the collective geographic range of its host species. Our study compares this assumption with a suite of other methods to infer parasite distribution from parasite occurrence data (e.g. based on data density, ecoregions and climatic conditions). We highlight diversity hotspots identified by the various methods and compare the effects of sampling intensities in different regions, a crucial factor of observed parasite diversity. Main conclusions The type of model used to infer parasite distributions affects estimates of both total species richness and spatial patterns of hotspots of parasite richness. Overall, the models based on reported occurrences share similar areas of high parasite richness that tends to be biased towards areas of high sampling effort. In contrast, the model based on host distributions showed hotspots of parasite diversity which are biased towards areas of high host species richness. Accounting for sampling effort could only help to reconcile the outcome from the different models in some regions. Further, the non-saturated species accumulation curves even for the best studied regions of the world such as Europe and North America as a call for further sampling effort and development of effective analytic tools that can provide robust accounts of global parasite diversity.
Figure 2 in Host-parasite relationships and life cycles of cuckoo wasps in agro-ecosystems in Argentina (Hymenoptera: Chrysididae: Chrysidini)
Figure 2. Emergence patterns of (a) Chrysis boutheryi (Brèthes) (squares; n = 20) and (b) C. saltana Bohart (triangles, n = 19) adults reared from trap-nests in Toay, La Pampa Province.
Figure 1 in Host-parasite relationships and life cycles of cuckoo wasps in agro-ecosystems in Argentina (Hymenoptera: Chrysididae: Chrysidini)
Figure 1. (a) Study sites: Toay (inverted triangle), Hortensia (square), Pila (triangle) and Colonia Elía (hexagon), situated in the Pampean region. The area encircled by thick line indicates the location of the Río de la Plata grasslands. Subdivisions are limited by dotted lines and identified by capital letters. A: Rolling Pampa; B: Inland Pampa; C: Southern Pampa; D: Flooding Pampa; E: Mesopotamic Pampa; F: Campos (modified from Medan et al.2011). (b–c) Trap-nests located in one tree and on fence posts.
Figure 1 in Coccidian parasites of red squirrels (Sciurus vulgaris) and grey squirrels (Sciurus carolinensis) in England
Figure 1. Outline map of England indicating collection sites. 1. Isle of Wight; 2. Fursey Island; 3. Epping Forest; 4. Thetford Chase; 5. Formby; 6. Cumbria.
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These curated guides explain access requirements, typical timelines, costs, and reuse considerations for widely used research datasets.
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.