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FIGURES 16–18 in The genital area of Halacaridae (Acari), life stages and development of morphological characters and implication on the classification
FIGURES 16–18. Arhodeoporus gracilipes (Trouessart, 1889), 16. genital region, female; 17. everted ovipositor, ventrolateral aspect; 18. genital opening, male. FIGURES 19–20. Arhodeoporus corallinus Otto, 2000, 19. genital region, female; 20. genital opening, male. FIGURES 21–22. Arhodeoporus bonairensis (Viets, 1936), 21. genital opening, female; 22. genital plate, deutonymph. FIGURES 23–24. Atelopsalis pacifica Bartsch, 1985, 23. posterior part of genitoanal plate, female, ventrolateral aspect (genital acetabula of the other side not illustrated); 24. genital plate, protonymph. FIGURES 25–26. Australacarus pustulatus Bartsch, 1993, 25. genitoanal plate, female; 26. genitoanal plate, male. FIGURE 27. Bathyhalacarus acutus Bartsch, 1982, genital opening, female. (fo, foveate surface structure; gac, genital acetabula; ov, ovipositor; pa, porose surface structure) Scale = 50 µm.
FIGURES 1–2. Acanthohalacarus reticulatus Bartsch, 2001, 1 in The genital area of Halacaridae (Acari), life stages and development of morphological characters and implication on the classification
FIGURES 1–2. Acanthohalacarus reticulatus Bartsch, 2001, 1. genital opening, female; 2. genital opening, male. FIGURES 3–4. Acarochelopodia biunguis Bartsch, 1988, 3. genitoanal plate, male; 4. genital plate, deutonymph. FIGURES 5–6. Acaromantis armatus Bartsch, 1977, 5. genital opening, female; 6. genital plate, protonymph. FIGURE 7. Actacarus spinosus Otto, 2000, posterior part of genitoanal plate, female. FIGURE 8. Agaue galatea Otto, 1999, genital plate, deutonymph. FIGURES 9–10. Agauopsis arborea Bartsch, 2003, 9. genital opening, female; 10. genital opening, male. FIGURE 11. Agauopsis arabia Bartsch & Chatterjee, 2001, everted ovipositor and two genital spines (enlarged), ventral aspect, female. FIGURE 12. Agauopsis mokari Otto, 1994, genital opening, male. FIGURE 13. Agauopsis sp. (A. furcata group), genital opening, female. FIGURES 14–15. Agauopsis moorea Bartsch, 1992, 14. genital opening, female; 15. genital opening, male. (agsp, apical genital spine; bgsp, basal genital spine; gac, genital acetabula; GF, genital foramen; gsc, genital sclerite; gsp, genital spines; pgs, perigenital setae; sgs, subgenital seta) Scale = 50 µm
FIGURES 41–43. Enterohalacarus minutipalpus Viets, 1938, 41 in The genital area of Halacaridae (Acari), life stages and development of morphological characters and implication on the classification
FIGURES 41–43. Enterohalacarus minutipalpus Viets, 1938, 41. anterior part of genital plate, female; 42. genital opening, lateral aspect, female; 43. genital plate, nymph. FIGURE 44. Halacarellus capuzinus (Lohmann, 1893), genital region, female. FIGURES 45–46. Halacarellus balticus (Lohmann, 1889), female, 45. everted ovipositor, lateral aspect; 46. apex of ovipositor, posteroapical part, lateral aspect. FIGURE 47. Halacaropsis hirsuta (Trouessart, 1889), genital opening, female. FIGURES 48–49. Halacarus bisulcus Viets, 1927, 48. genital opening, female; 49. genital opening, male. FIGURES 50–52. Halixodes novaezelandiae Bartsch, 1986, male, 50. genitoanal plate; 51. genital sclerites with stump-like, spinose subgenital setae; 52. genital opening (subgenital setae omitted). (aagsp, anteroapical genital spines; bgsp, basal genital spine; gac, genital acetabulum; gsp, genital spines; ov, ovipositor (at rest); pagsp, posteroapical genital spines; sgs, subgenital seta) Scale = 50 µm.
Example BAM file for Galaxy Visualization Development Tutorial
<p>An example BAM file that can be used for the tutorial on visualization plugin development in Galaxy.</p>
FIGURES 76–77. Subimaginal genitals developing under larval cuticle. 76 in Systematics of Indobaetis Müller-Liebenau & Morihara 1982, and related implications for some other Baetidae genera (Ephemeroptera)
FIGURES 76–77. Subimaginal genitals developing under larval cuticle. 76, Nigrobaetis niger (Linnaeus 1761)) («Nigrobaetis- type» of gonostyli folding); 77, Acerpenna pygmaea (Hagen 1861) («Labiobaetis-type» of gonostyli folding). (Abbreviations: gs1–gs3, 1st–3rd segments of gonostylus; m.s, median sterno-styligeral muscle; us, unistyliger.
FIGURE 6 in Complete larval development of Thor amboinensis (De Man, 1888) (Decapoda: Thoridae) described from laboratory-reared material and identified by DNA barcoding
FIGURE 6. Thor amboinensis. Decapodid: A, total animal, lateral view; B, antennule; C, antenna; D, mandibles; E, maxillule; F, maxilla; G, first maxilliped; H, second maxilliped; I, third maxilliped; I', detail of proximal segment of third maxilliped endopod; I'', detail of distal segment of third maxilliped endopod; J, first pereiopod; K, second pereiopod; L, third pereiopod; M, fourth pereiopod; N, fifth pereiopod; O, first pleopod; P, second pleopod; Q, third pleopod; R, fourth pleopod; S, fifth pleopod; T, telson and uropods; T', detail of telson. Scale bars: 0.5 mm (B–D, I–T); 0.1 mm (A, E–H, I', I'', T').
FIGURE 3 in Complete larval development of Thor amboinensis (De Man, 1888) (Decapoda: Thoridae) described from laboratory-reared material and identified by DNA barcoding
FIGURE 3. Thor amboinensis. Third zoea: A, total animal, lateral view; B, second maxilliped; C, first pereiopod; D, second pereiopod; E, third pereiopod; F, telson and uropods. Fourth zoea: G, antenna; H, maxillule; I, maxilla; J, second pereiopod; K, third pereiopod; L, telson and uropods. Fifth zoea: M, total animal, lateral view; N, antennule; O, third pereiopod; P, fourth and fifth pereiopods; Q, telson and uropods; Q', detail of telson. Scale bars: 0.5 mm (F, L, O, Q); 0.1 mm (A–E, G–K, M–N, P, Q').
FIGURE 1 in Complete larval development of Thor amboinensis (De Man, 1888) (Decapoda: Thoridae) described from laboratory-reared material and identified by DNA barcoding
FIGURE 1. Phylogenetic tree of representatives of the family Thoridae and one outgroup based on the mitochondrial cytochrome c oxidase I (COI) barcode gene sequences (658 bp). Sequences were aligned with those available in GenBank. The phylogenetic reconstruction was carried out with the Maximum Likelihood (ML) analysis, implemented in MEGA version 6.0 (Tamura et al. 2013). Numbers are support values for 10000 bootstraps.
FIGURE 5 in Complete larval development of Thor amboinensis (De Man, 1888) (Decapoda: Thoridae) described from laboratory-reared material and identified by DNA barcoding
FIGURE 5. Thor amboinensis. Eighth zoea: A, total animal, lateral view; B, antennule; C, antenna; D, mandibles; E, first pereiopod; E', detail of propodus and dactylus of first pereiopod; F, second pereiopod; G, third pereiopod; G', detail of propodus and dactylus of third pereiopod; H, fourth pereiopod; I, fifth pereiopod; J, first pleopod; K, second pleopod; L, third pleopod; M, fourth pleopod; N, fifth pleopod; O, telson and uropods. Scale bars: 0.5 mm (B, C, E–G, I–O); 0.1 mm (A, D, E', G', H).
FIGURE 2 in Complete larval development of Thor amboinensis (De Man, 1888) (Decapoda: Thoridae) described from laboratory-reared material and identified by DNA barcoding
FIGURE 2. Thor amboinensis. First zoea: A, total animal, lateral view; B, antennule; C, antenna; D, mandibles; E, maxillule; F, maxilla; G, first maxilliped; H, second maxilliped; I, third maxilliped; J, first pereiopod; K, telson. Second zoea: L, total animal, lateral view; M, antennule; N, antenna; O, first pereiopod; P, second pereiopod; Q, telson. Scale bars: 0.5 mm (A, K, Q); 0.1 mm (B–J, L–P).
FIGURE 4 in Complete larval development of Thor amboinensis (De Man, 1888) (Decapoda: Thoridae) described from laboratory-reared material and identified by DNA barcoding
FIGURE 4. Thor amboinensis. Sixth zoea: A, total animal, dorsal view; B, antenna; C, third pereiopod; D, fourth pereiopod; E, fifth pereiopod; F, telson and uropods. Seventh zoea: G, total animal, lateral view; H, detail of antennules exopod; I, maxillule; J, maxilla; K, first maxilliped; L, second maxilliped; M, third maxilliped; N, detail of dactylus of third pereiopod; O, fourth pereiopod; P, fifth pereiopod; Q, pleopods. Scale bars: 0.5 mm (B, D, E, F); 0.1 mm (A, C, G–M, O–Q).
Development of a flow chamber system for the reproducible in vitro analysis of biofilm formation on implant materials
<p>The data provided are the original data from the microscopic investigation of oral bacterial biofilms. Bacteria were stained with a life/dead staining. Viable cells are represented in red, non-viable cells are shown in green. The biofilms were gained by 50 stacks each with a CLSM. The biofilms were formed in a flow chamber system with a flow velocity of 100µL/min over 24-72 hours. The biofilms were grown on tintanium discs.</p>
Pernambuco State, Brazil (MZSP 30975). Fig. 1: ventral, lateral & dorsal views; Fig. 2: apical view, V = varix, L = edge of outer lip. Figs. 3–4: paratype from the type lot (MZSP 122078). Fig. 3: ventral view; Fig. 4: apical view (varix not developed in this immature specimen). Figs. 5–6: shell surface SEM images of holotype (MZSP 30975). Fig. 5: image taken at 500x showing pattern of paired grooves separated by wider region; the thin line is parallel to the axis of shell. Fig. 6: a closer look showing pattern of punctae (1,000x). SEM images by Yolanda Villacampa and courtesy of the Smithsonian. in Taxonomic review of tropical western Atlantic shallow water Drilliidae (Mollusca: Gastropoda: Conoidea) including descriptions of 100 new species
Pernambuco State, Brazil (MZSP 30975). Fig. 1: ventral, lateral & dorsal views; Fig. 2: apical view, V = varix, L = edge of outer lip. Figs. 3–4: paratype from the type lot (MZSP 122078). Fig. 3: ventral view; Fig. 4: apical view (varix not developed in this immature specimen). Figs. 5–6: shell surface SEM images of holotype (MZSP 30975). Fig. 5: image taken at 500x showing pattern of paired grooves separated by wider region; the thin line is parallel to the axis of shell. Fig. 6: a closer look showing pattern of punctae (1,000x). SEM images by Yolanda Villacampa and courtesy of the Smithsonian.
FIGURES 6–9 in Ultrastructural observations on the development of Ceratomyxa aegyptiaca (Myxozoa: Bivalvulida) infecting Solea aegyptiaca (Pleuronectiformes: Soleidae) from Tunisian coastal lagoon
FIGURES 6–9. Capsulogenesis of Ceratomyxa aegyptiaca. Fig. 6. Detail of two capsulogenic cells (Cc) in different stages of maturation. One capsulogenic cell (Cc) has a mature polar capsule (Pc). The second capsulogenic cell shows the sections of external tube (Et), a capsular primordium (Cp) and nucleus (CNu) in their cytoplasm. Sutural line (S) can be observed. Fig. 7. Detail of a capsulogenic cell showing a polar capsule with twisted appearance of polar filament (Pf). The apical side of the mature spore was surrounded by fibrous material (arrows) between the capsulogenic cell and the valve (V). Note the presence vacuoles (V) in the cytoplasm of the capsulogenic cell (Cc). Fig. 8. Longitudinal section of mature polar capsule (Pc) with its polar filament (arrows) showing an electron dense matrix (*) an outer electron-dense layer (OL) and an inner electron-lucent layer (IL). Note the channel for filament discharge (D) and externally, a surrounding valve (V) can be seen. Fig. 9. A section of spore showing the mature polar capsule (Pc), the opening for the extrusion of the polar filament (Op), two sporoplasm nucleus (SpNu) and one flattened valvogenic nuclei (VNu).
FIGURES 3–5 in Ultrastructural observations on the development of Ceratomyxa aegyptiaca (Myxozoa: Bivalvulida) infecting Solea aegyptiaca (Pleuronectiformes: Soleidae) from Tunisian coastal lagoon
FIGURES 3–5. Earlier sporogonic stages of Ceratomyxa aegyptiaca Fig. 3. Sporoblast showing two capsulogenic cells (Cc) with its nuclei (CNu) and with a prominent external tube (Et) and a capsular primordium (Cp), a sporoplasmogenic cell (SpC) with two nuclei (SpNu) and laterally two valvogenic cells (Vc) with its nuclei (VNu). Fig. 4. Young capsulogenic cell (Cc) with capsular primordium (Cp) composed of an electron-dense outer layer (1), an electron-lucent median layer (2) and an inner electron-dense core (3). Fig. 5. Immature spore attached to the epithelial cell (Ec) showing capsulogenic cell (Cc) with a prominent external tube (Et) in the cytoplasm and a clup-shaped capsular primordium (Cp) which is surrounded by valvogenic cell (Vc).
FIGURES 10–11 in Ultrastructural observations on the development of Ceratomyxa aegyptiaca (Myxozoa: Bivalvulida) infecting Solea aegyptiaca (Pleuronectiformes: Soleidae) from Tunisian coastal lagoon
FIGURES 10–11. Transmission electron micrographs of myxospores of Ceratomyxa aegyptiaca. Fig. 10. Premature myxosspore showing the sporoplasm cell (Spc) with its two nuclei (SpNu). Note the presence of one of the capsulogenic nuclei (CNu). Fig. 11. Young myxospore in frontal view showing two long valves (V) and the flattened valvogenic nuclei (VNu) of one valve.
FIGURES 1–2 in Ultrastructural observations on the development of Ceratomyxa aegyptiaca (Myxozoa: Bivalvulida) infecting Solea aegyptiaca (Pleuronectiformes: Soleidae) from Tunisian coastal lagoon
FIGURES 1–2. Transmission electron micrographs of Ceratomyxa aegyptiaca from the gallbladder of Solea aegyptiaca. Fig. 1. Initial primary cell (P) with large and condensed nucleus (N) containing several vacuoles (V). Fig. 2. Primary cell (P) attached to the gallbladder epithelium showing pinocytotic invaginations (arrows) extending into the host tissue (H).
Figure 5 in The larval development of the spider crab Rochinia gracilipes (Crustacea: Majoidea: Epialtidae: Pisinae) reared in the laboratory
Figure 5. Megalopa of Rochinia gracilipes A. Milne Edwards, 1875. (A) dorsal view, pereiopods drawn trunctated; (B) lateral view; (C) antennules, aesthetascs drawn truncated; (D) antenna; (E) mandible; (F) maxillule; (G) maxilla. Scale bars = 0.1 mm.
Figure 4 in The larval development of the spider crab Rochinia gracilipes (Crustacea: Majoidea: Epialtidae: Pisinae) reared in the laboratory
Figure 4. Second zoea of Rochinia gracilipes A. Milne Edwards, 1875. (A) maxilliped I; (B) maxilliped II; (C) maxilliped III buds; (D) pereiopod buds; (E) dorsal view of abdomen and telson; (F) magnified view of right furca. Long setae maxillipeds I and II (A and B) were drawn truncated. Scale bars = 0.1 mm.
Figure 1 in The larval development of the spider crab Rochinia gracilipes (Crustacea: Majoidea: Epialtidae: Pisinae) reared in the laboratory
Figure 1. First zoea of Rochinia gracilipes A. Milne Edwards, 1875. (A) lateral view; (B) magnified latero-dorsal view showing the short rostrum; (C) antennule; (D) antenna; (E) mandible; (F) maxillule; (G) maxilla. Scale bars = 0.1 mm.
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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.