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Figure 1 in The sting of Mesobuthus gibbosus (Scorpiones: Buthidae): morphological and ultrastructural characterization
Figure 1: A. Lateral view of M. gibbosus sting. The venom pore located at the tip of the sting, and several setae situated more basally are visible, x30. B. A cuticular seta, seta base, and a cuticular pit on the sting at higher magnification, x2,200
Fig. 6 in Identification and characterization of three juvenile hormone genes from Bactrocera dorsalis (Diptera: Tephritidae)
Fig. 6. Effects of starvation on the expression of BdJHEH2, BdJHEH3, and BdJH- DK in Bactrocera dorsalis. The 2-d-old 3rd instars were fed or starved for 24 and 48 h before collection. The re-fed larvae were initially starved for 24 h, then refed for an additional 24 h prior to collection. F24: feeding 24 h; F48: feeding 48 h; S24: starvation 24 h; S48: starvation 48 h; RF: re-fed. Different letters indicate significant differences based on 1-way ANOVA followed by an LSD test (P <0.05).
Fig. 3 in Identification and characterization of three juvenile hormone genes from Bactrocera dorsalis (Diptera: Tephritidae)
Fig. 3. Relative expression levels of BdJHEH2, BdJHEH3, and BdJHDK in developmental stages of Bactrocera dorsalis. Expression levels at 19 time points in (A) 3rd instar larvae and pupae and (B) adults were detected by qPCR. 3L1: 1-d-old 3rd instar larvae; P1: 1-d-old pupae. Females or males were collected for qPCR analysis at 1, 4, 7, and 10 d afer eclosion. Different letters indicate significant differences among females or males based on 1-way ANOVA followed by an LSD test (P <0.05). Significant differences between the females and males determined with a t-test are indicated by * (P <0.05).
Fig. 4 in Identification and characterization of three juvenile hormone genes from Bactrocera dorsalis (Diptera: Tephritidae)
Fig. 4. Relative expression levels of BdJHEH2, BdJHEH3, and BdJHDK in tissues of Bactrocera dorsalis. Expression levels in the head (HD), thorax (TH), midgut (MG), Malpighian tubules (MT), and fat body (FB) were detected by qPCR. Different letters indicate significant differences among tissues based on 1-way ANOVA followed by an LSD test (P <0.05).
Fig. 2 in Identification and characterization of three juvenile hormone genes from Bactrocera dorsalis (Diptera: Tephritidae)
Fig. 2. Multiple sequence alignment and phylogenetic analysis of JHDK from Bactrocera dorsalis and other insects. (A) Sequence alignment. The sequence of B. dorsalis JHDK is compared to SCP2 from Drosophila melanogaster (Dm), and JHDK from Leptinotarsa decemlineata (Ld), Manduca sexta (Ms), Plutella xylostella (Px), and Spodoptera litura (Sl). The α-helices (H1-8) and EF hands are indicated above the alignment according to results from B. mori (Li et al. 2005).Three predicted GTP-binding motifs (Σ1–3) are labeled with an arrow. (B) Phylogenetic analysis of JHDK homologs. The tree was generated with MEGA 5 using the neighbor-joining method. Nodes with>50% bootstrap values (1,000 replicates) are indicated on branches. GenBank accession numbers of all sequences are listed in the tree.
Fig. 1 in Identification and characterization of three juvenile hormone genes from Bactrocera dorsalis (Diptera: Tephritidae)
Fig. 1. Multiple sequence alignment and phylogenetic analysis of JHEHs from Bactrocera dorsalis and other insects. (A) Sequence alignment. The sequences of B. dorsalis JHEHs are compared with JHEH from Drosophila melanogaster (Dm), Bombyx mori (Bm), Manduca sexta (Ms), and Apis mellifera (Am). The catalytic triad (Asp232, Glu409, and His436), 2 tyrosine residues (Tyr 304 and Tyr380), and HGXP motif are labeled with asterisks. The HGXP motif is underlined. (B) Phylogenetic analysis of JHEH homologs. The tree was generated with MEGA 5 using the neighbor-joining method. Nodes with>50% bootstrap values (1,000 replicates) are indicated on branches. GenBank accession numbers of all sequences are listed in the tree.
Fig. 2 in Short Communication First record of Scolopendra cingulata Latreille 1829 (Chilopoda: Scolopendromorpha: Scolopendridae) in NW Italy and biotope characterization
Fig. 2 - Juvenile Scolopendra cingulata, observed on 9 April 2021. / Giovane di Scolopendra cingulata osservato il 9 aprile 2021. (Photo / Foto: Luca Anselmo).
Fig. 1 in Short Communication First record of Scolopendra cingulata Latreille 1829 (Chilopoda: Scolopendromorpha: Scolopendridae) in NW Italy and biotope characterization
Fig. 1 - Adult Scolopendra cingulata, observed on 9 April 2021 (a) and the updated distribution in Italy (b): previous data come from CKmap (Zapparoli & Minelli, 2005) and GBIF (2021). / Adulto di Scolopendra cingulata, osservato il 9 aprile 2021 (a) e distribuzione aggiornata in Italia (b): i dati precedenti provengono da CKmap (Zapparoli & Minelli, 2005) e GBIF (2021).
Fig. 1 in Characterization of Bacillus thuringiensis (Bacillaceae) strains pathogenic to Myzus persicae (Hemiptera: Aphididae)
Fig. 1. Protein profiles of the strains virulent to Myzus persicae. Lane 1: GP640, Lane 2: GP399, Lane 3: GP238, Lane 4: GP322, Lane 5: GP139, Lane 6: GP762, Lane 7: GP339, Lane 8: GP300, Lane 9: HD1, Lane 10: GP402, Lane 11: GP382, Lane 12: GP528, Lane 13: GP782, Lane 14: GP209, Lane 15: GP777, Lane 16: GP778, Lane 17: GP60, Lane 18: GP780.
Fig. 3 in Evaluation of blood and muscle tissues for molecular detection and characterization of hematozoa infections in northern pintails (Anas acuta) wintering in California
Fig. 3. Phylogenetic assignment of hematozoa mitochondrial DNA cytochrome b sequences originating from northern pintails collected from the Central Valley of California (asterisks). Reference sequences for Leucocytozoon (white circles), Haemoproteus (grey circles), and Plasmodium (black circles) parasites were obtained from the National Center for Biotechnology Information. Bootstrap support values for differentiation of broad taxonomic groups are indicated. Values reported for Haemoproteus and Plasmodium show support for phylogenetic differentiation from the mixed subclade formed by reference sequences a and b.
Fig. 4. Minimum spanning network for hematozoa mitochondrial DNA cytochrome b in Evaluation of blood and muscle tissues for molecular detection and characterization of hematozoa infections in northern pintails (Anas acuta) wintering in California
Fig. 4. Minimum spanning network for hematozoa mitochondrial DNA cytochrome b haplotypes detected in Central Valley northern pintails. Circles are drawn proportional to the frequency at which haplotypes were observed. Shading represents the sample collection from which haplotypes originated: white (2006– 2007 wing muscle), grey (2011–2012 wing muscle), and black (2011–2012 blood). A single mutation separates nodes unless explicitly indicated by number. Lines separating nodes are drawn to scale unless indicated by a break. Parasite taxa have been abbreviated in haplotype names (Leu = Leucocytozoon, Hae = Haemoproteus and Pla = Plasmodium).
Fig. 1 in Evaluation of blood and muscle tissues for molecular detection and characterization of hematozoa infections in northern pintails (Anas acuta) wintering in California
Fig. 1. Locations in the Central Valley of California from which northern pintail tissue samples were collected. Samples (n in 2006–2007, n in 2011–2012) were collected in the Sacramento Valley sub-region at: (A) Sacramento National Wildlife Refuge (NWR; 44,92), (B) Delevan NWR (35,7), (C) Colusa NWR (0,2), (D) Sutter NWR (0,1), (E) Little Dry Creek State Wildlife Area (SWA; 0,5), (F) Howard Slough SWA (0,1), (G) Yolo SWA (0,8) and (H) a private duck hunting club (0,1). Samples were collected in the San Joaquin Valley sub-region at: (I) San Luis NWR (7,0), (J) Kesterson NWR (4,0), (K) Los Banos SWA (14,0), (L) Volta SWA (10,0), (M) a private duck hunting club (2,0), and (N) Mendota SWA (30,40).
Fig. 2 in Evaluation of blood and muscle tissues for molecular detection and characterization of hematozoa infections in northern pintails (Anas acuta) wintering in California
Fig. 2. Estimated prevalence of Leucocytozoon (white bars), Haemoproteus (grey bars), and Plasmodium (black bars) parasites in northern pintails sampled in the Central Valley of California in 2006–2007 and 2011–2012 using occupancy modeling. Error bars represent 95% confidence intervals around point estimates.
Fig. 2 in Plasmodium (Novyella) nucleophilum from an Egyptian Goose in São Paulo Zoo, Brazil: microscopic confirmation and molecular characterization
Fig. 2. Bayesian phylogeny of cytochrome b gene lineages of species of avian haemosporidian parasites. A lineage recorded in the Egyptian Goose Alopochen aegyptiacus is provided underlined. Names of the lineages are given after the species names of parasites. GenBank accession numbers of the lineages are provided before the parasite species names. Nodal support values (in percentage) indicate posterior clade probabilities. Plasmodium species from Novyella subgenus are boxed.
Fig. 1 in Plasmodium (Novyella) nucleophilum from an Egyptian Goose in São Paulo Zoo, Brazil: microscopic confirmation and molecular characterization
Fig. 1. Photomicrographs of Plasmodium parasites visualized from thin blood smears obtained from an Egyptian Goose (Alopochen aegyptiacus) in São Paulo Zoo, Brazil. Characteristic of Plasmodium (Novyella) nucleophilum (lineage EG01, GenBank JX467689) the trophozoite (a), meronts (b–c), macrogametocytes (d, e), and microgametocyte (f) are appressed to erythrocyte nuclei (nucleophilic features). Plasmodium (Haemamoeba) sp. (g–i) lacks nucleophilic blood stages and possesses large roundish trophozoites, each with a prominent centrally located vacuole; pigment granules are gathered around the vacuoles. Note that early Plasmodium (H.) sp. trophozoites markedly displace erythrocyte nuclei (g). Arrows, pigment granules. Scale bar = 10 µm.
Fig. 1 in Genetic characterization of Toxoplasma gondii from Brazilian wildlife revealed abundant new genotypes
Fig. 1. Phylogenetic network analysis of Toxoplasma gondii from wildlife in Brazil. Genotype ID and the representative strain are listed for each taxonomic branch. Reference strains are in black, the strains from this study are in red, and the Amazonic reference strains that did not cluster together are in boxes. Inside the circle are listed all the genotypes obtained from the Amazon region which are in the same branch as other isolates from this biome.
Fig. 2 in Genetic characterization of Toxoplasma gondii from Brazilian wildlife revealed abundant new genotypes
Fig. 2. Geographical distribution of the genotypes of Toxoplasma gondii from wildlife in Brazil. Samples are grouped by states. Sample size is represented by the size of the bar and the number written in brackets. The smallest bar represents one genotype. Color code: green, purple, red and blue are for BrI, BrII, previously described atypical genotypes and new atypical genotypes, respectively.
Fig. 1 in Morphological and molecular characterization of Eimeria purpureicephali n. sp. (Apicomplexa:Eimeriidae) in a red-capped parrot (Purpureicephalus spurius, Kuhl, 1820) in Western Australia
Fig. 1. Nomarski interference-contrast photomicrographs of E. purpureicephali n. sp. oocysts showing spheroidal to subspheroidal sporocysts (scale bar = 20 Mm) (1—5) and line drawing of the sporulated oocyst of E. purpureicephali n. sp. Scale bar = 20 Mm (6).
Fig. 5 in Molecular characterization of trypanosomatid infections in wild howler monkeys (Alouatta caraya) in northeastern Argentina
Fig. 5. Agarose gel (2%) showing RibDNA-amplified fragments stained with ethidium bromide. Samples are indicated by their ID numbers. Reference strains were used as positive controls: TCI: T. cruzi I, TCII: T. cruzi II, and TR: T. rangeli. M: 1 kb DNA molecular ladder. Fragment size is indicated in base pairs.
Fig. 2 in Molecular characterization of trypanosomatid infections in wild howler monkeys (Alouatta caraya) in northeastern Argentina
Fig. 2. Study areas showing the locations of the sampled howler groups and the results of molecular analysis: groups with only RibDNA PCR-positive howler monkeys (gray circle); groups with RibDNA and kDNA-PCR-positive howler monkeys (white circle); groups with RibDNA, kDNA, and SatDNA-PCR-positive howler monkeys positive (black circle). A: Isla Brasilera (IB) and Isla del Cerrito (IC); B: San Cayetano (SC) and Estacíon Bioĺogica de Corrientes (EBCo).
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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.