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FIGURE 2 in Neotypification of Cistanche tubulosa (Schenk) Wight ex Hook.f.: a name applied to a widely distributed, polyphyletic group of plants
FIGURE 2. Cistanche tubulosa: A. Inflorescence, B. Corolla, showing calyx and serrated bract and small central lower lobe vs. lateral lobe in profile, C. View of compressed calyx showing sub-equal lobes. Specimens were illustrated based on the neotype and with reference to living material from the Middle East. Illustrations by CJ Thorogood.
FIGURE 1. A in Neotypification of Cistanche tubulosa (Schenk) Wight ex Hook.f.: a name applied to a widely distributed, polyphyletic group of plants
FIGURE 1. A typical specimen of Cistanche tubulosa growing in Eilat, c. 230 km from Wadi Feiran Oasis where the neotype was collected. B–E: morphological features of C. tubulosa including the calyx (B), filaments (C), corolla tube (D) and bract (E). F. Cistanche violacea, a putative species that co-occurs with C. tubulosa in the Middle East; G. An intermediate morphotype of the former two species where they co-occur in a wadi in southern Israel. Photos by CJ Thorogood.
FIGURE 3 in Neotypification of Cistanche tubulosa (Schenk) Wight ex Hook.f.: a name applied to a widely distributed, polyphyletic group of plants
FIGURE 3. The name Cistanche tubulosa is applied to entities in different clades, but with overlapping distributions: A. Relationships within the 'Widespread Clade' (Ataei et al., 2020) summarised to show the placement of species identified by the authors as C. tubulosa (plants from China in the 'C. laxiflora clade'; plants from the Middle East in the 'C. tubulosa clade') or as aff. C. tubulosa (plants from Oman and Yemen in the 'C. senegalensis clade'). Figure 3B. Distribution map of the specimens sequenced by Ataei et al. (2020) included in the phylogeny. The map shows specimens in the C. tubulosa clade in red, and specimens placed in the C. senegalensis clade in blue (note the overlap), including the four specimens referred to as C. aff. tubulosa (three in Oman and one in Yemen). The type localities of C. tinctoria and C. tubulosa are shown as a black circle and black square respectively.
FIGURE 1 in Aphids of Iran: their host plants and distribution
FIGURE 1. Map of the provinces of Iran (By Ali Zifan - Own work; derived from File:BlankMap-World6-Equirectangular.svg, CC BY-SA 4.0, https://commons.wikimedia.org/w/index.php?curid=58124425).
FIGURE 2 in Aphids of Iran: their host plants and distribution
FIGURE 2. Aphid species diversity index for each province of Iran based on the number of aphid species reported from each province and its area.
FIGURES 78–83 in Exploring the diversity of Gracillariidae (Lepidoptera) in South Africa: host plants, distribution, and DNA barcoding analysis, with the description of nine new species
FIGURES 78–83. Immature stages of South African Gracillariidae. 78, Phodoryctis tephrosiella, mines on Tephrosia rhodesica (Fabaceae). Gauteng, Tshwane, A. Sharp leg. 79, Conopobathra carbunculata, mines on Peltophorum africanum (Fabaceae), Limpopo, York, A. & I. Sharp leg. 80, Phyllonorycter grewiella, mine on Grewia hexamita (Malvaceae), A. & I. Sharp leg. 81, P. pseudogrewiella sp. nov., mines on Grewia flavescens (Malvaceae), Limpopo, Hoedspruit, A. & I. Sharp leg. 82, P. pseudogrewiella sp. nov., larva, ibidem. 83, P. pseudogrewiella sp. nov., mines and pupae, ibidem.
FIGURES 67–72 in Exploring the diversity of Gracillariidae (Lepidoptera) in South Africa: host plants, distribution, and DNA barcoding analysis, with the description of nine new species
FIGURES 67–72. Immature stages of South African Gracillariidae. 67, Conopomorphina aptata, mine on Schotia brachypetala (Fabaceae), Limpopo, Hoedspruit, A. & I. Sharp leg. 68, Acrocercops syzygiena, mine on Syzygium cordatum (Myrtaceae), Gauteng, Tshwane, A. & I. Sharp leg. 69, A. syzygiena, larva, ibidem. 70, A. combreticola, mine on Combretum zeyheri (Combretaceae) Gauteng, Tshwane, A. Sharp leg. 71, Cryptolectica capnodecta, larva on Syzygium cordatum (Myrtaceae), Limpopo, Hoedspruit, A. & I. Sharp leg. 72, C. terminalina, mine on Terminalia sericea (Combretaceae), A. & I. Sharp leg.
FIGURES 84−88 in Exploring the diversity of Gracillariidae (Lepidoptera) in South Africa: host plants, distribution, and DNA barcoding analysis, with the description of nine new species
FIGURES 84−88. Immature stages of South African Gracillariidae species. 84, Cameraria melhaniella sp. nov., larva on Melhania acuminata (Malvaceae), Limpopo, Hoedspruit, A. & I. Sharp leg. 85, Metriochroa pergulariae, mine on Pergularia daemia (Apocynaceae), Limpopo, Hoedspruit, A. & I. Sharp leg. 86, Phyllocnistis allisonae, mine on Protea rubropilosa (Proteaceae), Limpopo, Lopez Vaamonde leg. 87, Phyllocnistis magalismontani sp. nov., mine on Englerophytum magalismontanum (Sapotaceae), Limpopo, Hoedspruit, leg. A. & I. Sharp. 88, Phyllocnistis faureae, mine on Faurea saligna (Proteaceae), Gauteng, Tshwane, leg. A. Sharp.
FIGURES 61−66 in Exploring the diversity of Gracillariidae (Lepidoptera) in South Africa: host plants, distribution, and DNA barcoding analysis, with the description of nine new species
FIGURES 61−66. Immature stages of South African Gracillariidae. 61, Caloptilia sp., larva on Cryptocarya transvaalensis (Lauraceae), Limpopo, Lekgalameetse N. R., A. & I. Sharp leg. 62, C. cataractias, larva on Rhynchosia minima (Fabaceae), Limpopo, Hoedspruit, A. & I. Sharp leg. 63, Macarostola noellineae, larva on Syzygium cordatum (Myrtaceae), Limpopo, Hoedspruit, A. & I. Sharp leg. 64, Ectropina spirostachydis sp. nov., larva on Spirostachys africana (Euphorbiaceae), Limpopo, Hoedspruit, A. & I. Sharp leg. 65, Cuphodes melanostola, mine on Euclea divinorum (Ebenaceae), Limpopo, Hoedspruit, A. & I. Sharp leg. 66, Conopomorphina ochnivora, Gauteng, Magaliesburg, on Ochna pretoriensis, (Ochnaceae), H. Staude leg.
FIGURES 58–60 in Exploring the diversity of Gracillariidae (Lepidoptera) in South Africa: host plants, distribution, and DNA barcoding analysis, with the description of nine new species
FIGURES 58–60. Female genitalia of South African Gracillariidae. 58-59, Phyllocnistis magalismontani sp. nov., paratype: 58, lateral view; 59, signa. 60, P. allisonae sp. nov., holotype: ventral view. (All scale bar 190 μm).
FIGURES 56–57 in Exploring the diversity of Gracillariidae (Lepidoptera) in South Africa: host plants, distribution, and DNA barcoding analysis, with the description of nine new species
FIGURES 56–57. Female genitalia of South African Gracillariidae. 56, Telamoptilia cordati sp. nov., paratype: ventral view. 57, Phyllonorycter pseudogrewiella sp. nov., holotype: lateral view. (All scale bar 190 μm).
FIGURES 44–46 in Exploring the diversity of Gracillariidae (Lepidoptera) in South Africa: host plants, distribution, and DNA barcoding analysis, with the description of nine new species
FIGURES 44–46. Male genitalia of South African Gracillariidae. Phodoryctis tephrosiella sp. nov.: 44, holotype, ventral view; 45, phallus with bulbus ejaculatorius; 46, segment VIII in ventral view (ae: aedeagus; pb: phallobase; be: bulbus ejaculatorius). (All scale bars 190 μm).
FIGURES 25−30 in Exploring the diversity of Gracillariidae (Lepidoptera) in South Africa: host plants, distribution, and DNA barcoding analysis, with the description of nine new species
FIGURES 25−30. Forewing pattern of South African Gracillariidae. 25, Leucocercops curatellifoliae sp. nov., Limpopo. 26, Phodoryctis tephrosiella sp. nov., Gauteng. 27, Telamoptilia cordati sp. nov., South Africa, Limpopo. 28, Telamoptilia sp., Madagascar. 29, Cameraria melhaniella sp. nov., Limpopo. 30, Phyllonorycter pseudogrewiella sp. nov., Limpopo.
FIGURES 13–18 in Exploring the diversity of Gracillariidae (Lepidoptera) in South Africa: host plants, distribution, and DNA barcoding analysis, with the description of nine new species
FIGURES 13–18. Adults of South African Gracillariidae. 13, Cryptolectica capnodecta, Limpopo, Hoedspruit, A. & I. Sharp leg. 14, C. terminalina, Limpopo, Hoedspruit, A. & I. Sharp leg. 15, Dialectica pyramidota, Limpopo, Hoedspruit, A. & I. Sharp leg. 16, Amblyptila cynanchi, Western Cape, Knysna, S. Mecenero leg. 17, Leucocercops dasmophora, Gauteng, Tshwane, A. & I. Sharp leg. 18, L. curatellifoliae sp. nov. Limpopo, Hoedspruit, A. & I. Sharp leg.
FIGURES 32–34 in Exploring the diversity of Gracillariidae (Lepidoptera) in South Africa: host plants, distribution, and DNA barcoding analysis, with the description of nine new species
FIGURES 32–34. Forewing pattern of the South African Phyllocnistinae. 32, Phyllocnistis magalismontani sp. nov., drawn by holotype; 33, P. allisonae sp. nov., holotype; 34, P. faureae, holotype.
FIGURE 31 in Exploring the diversity of Gracillariidae (Lepidoptera) in South Africa: host plants, distribution, and DNA barcoding analysis, with the description of nine new species
FIGURE 31. Forewing pattern of the Afrotropical Phyllocnistinae. A, Phyllocnistis pharetrucha; B, P. saligna; C, P. magalismontani sp. nov.; D, P. faureae sp. nov.; E, P. loxosticha; F, P. citrella; G, P. allisonae sp. nov..
FIGURES 1−6 in Exploring the diversity of Gracillariidae (Lepidoptera) in South Africa: host plants, distribution, and DNA barcoding analysis, with the description of nine new species
FIGURES 1−6. Adults of South African Gracillariidae. 1, Caloptilia sp., Limpopo, Lekgalameetse N. R., A. & I. Sharp leg. 2, Caloptilia rhusina, Western Cape, Robberg Nature Reserve, S. Mecenero leg. 3, Macarostola noellineae, Limpopo, Hoedspruit, A. & I. Sharp leg. 4, Caloptilia cataractias, Limpopo, Hoedspruit, leg. A. & I. Sharp. 5, Ectropina spirostachydis sp. nov., Limpopo, Hoedspruit, A. & I. Sharp leg. 6, Graphiocephala barbitias, Gauteng, Tshwane, A. Sharp leg.
FIGURES 7–12 in Exploring the diversity of Gracillariidae (Lepidoptera) in South Africa: host plants, distribution, and DNA barcoding analysis, with the description of nine new species
FIGURES 7–12. Adults of South African Gracillariidae. 7, Semnocera procellaris, Limpopo, Hoedspruit, A. & I. Sharp leg. 8, Conopomorphina ochnivora, Gauteng, Magaliesburg, A. & I. Sharp leg. 9, C. aptata, Limpopo, Hoedspruit, A. & I. Sharp leg. 10, Cuphodes melanostola, Limpopo, Hoedspruit, A. & I. Sharp leg. 11, Acrocercops syzygiena, Gauteng, Tshwane, A. Sharp leg. 12, A. combreticola, Gauteng, Tshwane, A. Sharp leg.
Quantifying niche similarity among new world seed plants--Species Distribution Models (SDMs) & associated metadata
<p>Niche shift and conservatism are often framed as mutually exclusive. However, both processes could contribute to biodiversity patterns. We tested this expectation by quantifying the degree of climatic niche similarity among New World seed plants.</p> <p>To incorporate the biological reality that species experience varied abiotic conditions across their range, we assembled distribution models and used these to characterize temperature, precipitation, and elevation niches for species as continuously-valued distributions. We then quantified niche similarity (distributional overlap) and identified statistically significant differences compared to a randomized null.</p> <p>The degree of niche similarity differed among climate variables, plant lineages, and at different phylogenetic scales. For example, ~17% of all seed plants were significantly different in elevational niche from their closest relative(s), whereas for precipitation, this value was only ~4%. Average niche similarity decreased with increasing phylogenetic distance, consistent with niche conservatism; however, variance in niche similarity among close relatives was large, such that there always existed niche differences equaling those among distantly related species.</p> <p>Our results suggest researchers should incorporate both niche shift and conservatism as important, scale-dependent factors shaping biodiversity patterns as these processes are not mutually exclusive, nor do they contribute equally to patterns among different plant lineages or niche variables.</p>
Climatic niche shifts in 815 introduced plant species affect their predicted distributions: Data and scripts
<p class="CxSpFirst"><u>Aim:</u> Introduced species often occupy different climates in their introduced than their native range, but to what degree do such 'climatic niche shifts' interfere with our ability to predict invasions? Answering this question is crucial if we are to understand the threat invasive species pose to human and natural systems, especially given the ever increasing use of species distribution models as tools for invasive species risk assessment and management. Here we investigated how strongly climatic niche shifts interfered with the transferability of native- and introduced-range species distribution models.</p> <p class="CxSpMiddle"><u>Location:</u> Our dataset consisted of ~14 million occurrences distributed worldwide.</p> <p class="CxSpMiddle"><u>Time Period:</u> Occurrence data were collected from online repositories dating from ca. 1600 with the vast majority being from the 20<sup>th</sup> century. Climatic data represent means between 1970–2000.</p> <p class="CxSpMiddle"><u>Major Taxa Studied:</u> Our database represented 815 terrestrial plant species.</p> <p class="CxSpMiddle"><u>Methods:</u> We used ordination to identify climatic niche shifts as species moved between continents. Next, we trained separate MAXENT models using native- or introduced-range occurrences, and projected those models into each species' introduced range. We compared the ordination and MAXENT models to determine whether niche shifts were associated with errors in MAXENT predictions.</p> <p class="CxSpMiddle"><u>Results:</u> Models trained on native-range occurrences poorly predicted introduced-range occurrences, and transferability was lowest in species with large climatic niche shifts. Directional shifts in species' predicted geographic distributions mirrored their niche dynamics. This is concerning because native-range data are often used to predict introduced-range distributions.</p> <p><u>Main Conclusions: </u>Our results highlight the importance of considering niche shifts when modeling the potential geographic distributions of introduced species, and cast doubt on the assumption that the climatic niche of a species can be transferred between native and invasive ranges.</p>
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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.