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45 results for “Red coloration”
Red, yellow, green and blue are not particularly colorful
<p>This is supplementary material accompanying the article:</p> <p>Witzel, C., Maule, M., & Franklin, A. (2019) Red, yellow, green and blue are not particularly colorful. <em>Journal of Vision</em>.</p> <p>focsat_data.xlsx contains all the individual data, including adjustments of typical and unique hues (+ super-saturated condition), detection (JND0) and discrimination (JND) data for red, yellow, green, and blue; and average saturation matches (subjective saturation) from Witzel and Franklin (2014). Nine sheets overall. IMPORTANT: All data is matched by participants (rows); but participant ids are not provided for reasons of data protection. Empty rows correspond to missing data. Also note that detection thresholds are provided in the first 10 columns, discrimination thresholds in the following 10 columns (11-20).</p> <p>focsat_tables.xlsx contains the exact data from tables in the article (Table 2) and the Supplementary Tables S1-S10. Eleven sheets in total.</p> <p>weberfechner.m is a Matlab function that allows for calculating Weber fractions and discriminable saturation as reported in the article.</p>
Text-fig. 6. Stratigraphic and phylogenetic placement inferred for fossil Fraxinus fruits. Only Fraxinus fossil fruits identified on the section level are included. The black color represents selected fossil fruits from published literature (excluding some Eocene North American occurrences not assigned to section), the red color represents the fossil fruits from the Lühe flora, Yunnan, Southwest China. The phylogenetic relationships are based on Hinsinger et al. (2013). in Fraxinus L. (Oleaceae) Fruits From The Early Oligocene Of Southwest China And Their Biogeographic Implications
Text-fig. 6. Stratigraphic and phylogenetic placement inferred for fossil Fraxinus fruits. Only Fraxinus fossil fruits identified on the section level are included. The black color represents selected fossil fruits from published literature (excluding some Eocene North American occurrences not assigned to section), the red color represents the fossil fruits from the Lühe flora, Yunnan, Southwest China. The phylogenetic relationships are based on Hinsinger et al. (2013).
Рис. 3. Схема миграций виΑов рыб, участвующих в современной Αинамике ихтиофауны на территории НТТ: 1 — разΛивы; 2 — намывы. Черным цветом обозначены направΛения миграций из реки Туманной; красным — из оз. Хасан и РазΛивов; синим — с мест зимовки в реках южного Приморья; зеΛеным — сезонные миграции из южных морей Fig. 3. Scheme of migration of fish species involved in the modern dynamics of ichthyofauna on the territory of LRT: 1 — spills; 2 — alluvial. Black color indicates the direction of migration from the Tumannaya river; red — from lake Khasan and spills; blue — from wintering places in the rivers of southern Primorye; green — seasonal migration from the south seas in Transboundary Migration And The Local Constraints In The Dynamic Of Fish Fauna In The Lower Reaches Of Tumannaya River
Рис. 3. Схема миграций виΑов рыб, участвующих в современной Αинамике ихтиофауны на территории НТТ: 1 — разΛивы; 2 — намывы. Черным цветом обозначены направΛения миграций из реки Туманной; красным — из оз. Хасан и РазΛивов; синим — с мест зимовки в реках южного Приморья; зеΛеным — сезонные миграции из южных морей Fig. 3. Scheme of migration of fish species involved in the modern dynamics of ichthyofauna on the territory of LRT: 1 — spills; 2 — alluvial. Black color indicates the direction of migration from the Tumannaya river; red — from lake Khasan and spills; blue — from wintering places in the rivers of southern Primorye; green — seasonal migration from the south seas
Рис. 1. ФиΛогенетические Αеревья хантавируса AMRV и его прироΑного носитеΛя восточноазиатской мыши Apodemus peninsulae Thomas, 1906. А. ФиΛогенетическое Αерево восточноазиатской мыши Apodemus peninsulae, построенное метоΑом «максимаΛьного правΑопоΑобия» (ML) и поΛученное на основе анаΛиза участка гена цитохрома b мтΔНК (744 п.н.). В узΛах ветвΛения указаны бутстреп-поΑΑержки, рассчитанные ΑΛя 1000 повторов. Цветными Λиниями обозначены фиΛогенетические Λинии: Αве Китайские (зеΛеный), Корейская «Korea» (синий), Амурская «Amur» (красный). ПоΛужирным шрифтом выΑеΛены собственные образцы. Названия образцов из GenBank/NCBI быΛи сокращены; B. ФиΛогенетическое Αерево из работы Α. Н. Яшиной с ΑопоΛнениями, построенное метоΑом «бΛижайшего сосеΑа» (NJ) на основе посΛеΑоватеΛьностей фрагмента М-сегмента (2737–2980 н.п.) генома хантавирусов. В узΛах ветвΛения указаны бутстреппоΑΑержки, рассчитанные ΑΛя 1000 повторов. Жирным выΑеΛены иссΛеΑованные РНК изоΛяты (Яшина 2012; Яшина и Αр. 2019) Fig. 1. Phylogenetic trees of AMRV and its natural reservoir host — the Korean field mouse Apodemus peninsulae Thomas, 1906. A. Phylogenetic tree of the Korean field mouse Apodemus peninsulae constructed by the "maximum likelihood" method (ML). The data are obtained from the analysis of the cytochrome b mtDNA gene fragments (744 bp). Bootstrap supports calculated for 1,000 repeats are indicated in the branching nodes. Colored lines indicate phylogenetic lines: two Chinese (green), Korea (blue), and Amur (red). Own samples are highlighted in bold. The names of the samples from GenBank/NCBI have been shortened; B. Phylogenetic tree from L. N. Yashina's work with additions constructed by the neighbour joining method (NJ). It is based on the sequences of an M-segment fragment (2737–2980 bp) of the hantavirus genome. Bootstrap supports calculated for 1,000 repeats are indicated in the branching nodes. The researched RNA isolates are highlighted in bold (Yashina 2012; Yashina et al. 2019) in Variability of the gene cyt b in the Korean field mouse Apodemus peninsulae Thomas, 1906 - a reservoir host of AMRV in the Khasansky District of Primorsky Krai
Рис. 1. ФиΛогенетические Αеревья хантавируса AMRV и его прироΑного носитеΛя восточноазиатской мыши Apodemus peninsulae Thomas, 1906. А. ФиΛогенетическое Αерево восточноазиатской мыши Apodemus peninsulae, построенное метоΑом «максимаΛьного правΑопоΑобия» (ML) и поΛученное на основе анаΛиза участка гена цитохрома b мтΔНК (744 п.н.). В узΛах ветвΛения указаны бутстреп-поΑΑержки, рассчитанные ΑΛя 1000 повторов. Цветными Λиниями обозначены фиΛогенетические Λинии: Αве Китайские (зеΛеный), Корейская «Korea» (синий), Амурская «Amur» (красный). ПоΛужирным шрифтом выΑеΛены собственные образцы. Названия образцов из GenBank/NCBI быΛи сокращены; B. ФиΛогенетическое Αерево из работы Α. Н. Яшиной с ΑопоΛнениями, построенное метоΑом «бΛижайшего сосеΑа» (NJ) на основе посΛеΑоватеΛьностей фрагмента М-сегмента (2737–2980 н.п.) генома хантавирусов. В узΛах ветвΛения указаны бутстреппоΑΑержки, рассчитанные ΑΛя 1000 повторов. Жирным выΑеΛены иссΛеΑованные РНК изоΛяты (Яшина 2012; Яшина и Αр. 2019) Fig. 1. Phylogenetic trees of AMRV and its natural reservoir host — the Korean field mouse Apodemus peninsulae Thomas, 1906. A. Phylogenetic tree of the Korean field mouse Apodemus peninsulae constructed by the "maximum likelihood" method (ML). The data are obtained from the analysis of the cytochrome b mtDNA gene fragments (744 bp). Bootstrap supports calculated for 1,000 repeats are indicated in the branching nodes. Colored lines indicate phylogenetic lines: two Chinese (green), Korea (blue), and Amur (red). Own samples are highlighted in bold. The names of the samples from GenBank/NCBI have been shortened; B. Phylogenetic tree from L. N. Yashina's work with additions constructed by the neighbour joining method (NJ). It is based on the sequences of an M-segment fragment (2737–2980 bp) of the hantavirus genome. Bootstrap supports calculated for 1,000 repeats are indicated in the branching nodes. The researched RNA isolates are highlighted in bold (Yashina 2012; Yashina et al. 2019)
3D Parameter Maps of Red Clump Stars in the Milky Way -- Absolute Magnitudes and Intrinsic Colors
<p>Catalog, application and supplementary figures for the RC paper.</p> <p>1100mRC_WC2021new.fits.zip -- RC catalog</p> <p>Gaia photometric RC sample with 11 million RCs selected based on {\it Gaia}'s EDR3 parallax, our 3D parameter maps, and extinction--distance profile.</p> <p>ra, dec -- position; dm -- distance modulus; av -- V-band extinction</p> <p>dm1 -- corrected distance modulus; av1 -- corrected extinction. used for low-extinction RCs</p> <p>edm -- uncertainty of distance modulus; flag -- 1: high probability RCs, 0: low probability RCs.</p> <p> </p> <p>RC_calculator_linux.zip -- application RC2021 and installer for linux</p> <p>RC_calculator_mac.zip -- application RC2021 and installer for mac </p> <p>The application to estimate the absolute magnitudes and intrinsic colors for RCs with the APOGEE or LAMOST parameters</p> <p> </p> <p>eps -- the best applicable ranges for using application.</p> <p>pd_APOGEE.eps and pd_LAMOST.eps -- Probability densities of each parameter for absolute magnitude training set (orange) and the whole sample (blue). </p> <p>pd_APOGEE_color.eps and pd_LAMOST_color.eps -- Probability densities of each parameter for intrinsic color training set (orange) and the whole sample (blue). </p>
Data from: The role of red coloration and song in peacock spiders: insights into complex signaling systems
Research on animal signaling enhances our understanding of links between sensory processing, decision-making, behavior, and evolution. Studies of sexually-selected signals may be particularly informative as mate choice provides access to decision patterns in the way that courtship leads to an easily observable behavioral output in choosers, i.e. mating. Male peacock spiders have some of the most elaborate and varied courtship displays known among animals. Particularly striking to human observers is the diversity of red, orange and yellow ornaments that males exhibit across the genus. The primary objective of our research was to investigate how these visual ornaments interact with vibratory songs to affect female mating behavior of one species, Maratus volans. Accordingly, we conducted mating trials under a series of experimentally manipulated vibratory and lighting conditions. Contrary to expectation, chromatic characteristics of longer wavelength ornaments are not driving female mate choice decisions, despite their extensive presence on male fans. Instead, our results suggest that contrast is important to females. Additionally, we found that vibratory signals were not necessary and did not increase mating rates. Our study demonstrates the intricacies inherent in complex signaling systems.
A mechanism for red coloration in vertebrates
<p>Red coloration is a salient feature of the natural world. Many vertebrates produce red color by converting dietary yellow carotenoids into red ketocarotenoids via an unknown mechanism. Here, we show that two enzymes, cytochrome P450 2J19 (CYP2J19) and 3-hydroxybutyrate dehydrogenase 1-like (BDH1L), are sufficient to catalyze this conversion. In birds, both enzymes are expressed at sites of ketocarotenoid biosynthesis (feather follicles and red cone photoreceptors), and genetic evidence implicates these enzymes in yellow/red color variation in feathers. In fish, homologs of CYP2J19 and BDH1L are required for ketocarotenoid production, and we show that these enzymes are sufficient to produce ketocarotenoids in cell culture and when ectopically expressed in fish skin. Finally, we demonstrate that the red-cone-enriched tetratricopeptide repeat protein 39B (TTC39B) enhances ketocarotenoid production when co-expressed with CYP2J19 and BDH1L. The discovery of this mechanism of ketocarotenoid biosynthesis has major implications for understanding the evolution of color diversity in vertebrates.</p>
From green to red: Urban heat stress drives leaf color evolution
<p><span>Urban environments, occupying approximately 1% of total land area, often impose novel biotic and abiotic selective pressures on organisms and provide valuable opportunities to understand the eco-evolutionary dynamics between nature and human societies. Prevalence of impervious surface and resulting higher temperatures in urban areas, known as urban heat islands, comprises prominent characteristics in global cities. However, it is not known whether and how urban plants adapt to such heat stress. This study focused on <em>Oxalis</em> <em>corniculata</em>, which has intraspecific polymorphism in leaf color (green, red), and examined whether the leaf color variation is associated with urban heat stress. Field observations revealed consistent associations between leaf color and habitat types (green vs. urban) at local (< 500m), landscape (< 50km), and global scales. Green-leaved plants were dominant in green habitats, and red-leaved individuals had increased in number in urban habitats. Growth and photosynthesis experiments indicated the adaptive benefit and cost of red/green leaves associated with heat stresses. Red-leaved individuals had higher growth rates and photosynthetic efficiency under heat stress, while green-leaved individuals displayed higher growth rates and photosynthetic efficiency under non-stressful conditions. Genome-wide SNP analysis suggests that the red leaf trait may have evolved multiple times from the ancestral green leaf, rather than spreading from a single origin of red leaf evolution. Overall, the results suggested that the dominance of red leaves of <em>O. corniculata</em> seen in cities worldwide would be evidence of plant adaptative evolution due to urban heat islands.</span></p>
Data from: The role of red coloration and song in peacock spiders: insights into complex signaling systems
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From green to red: Urban heat stress drives leaf color evolution
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A mechanism for red coloration in vertebrates
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Female ornaments: is red skin color attractive to males and related to condition in rhesus macaques?
Sexual selection produces extravagant male traits, such as colorful ornaments, via female mate choice. More rarely, in mating systems in which males allocate mating effort between multiple females, female ornaments may evolve via male mate choice. Females of many anthropoid primates exhibit ornaments that indicate intra-individual cyclical fertility, but which have also been proposed to function as inter-individual quality signals. Rhesus macaque females are one such species, exhibiting cyclical facial color variation that indicates ovulatory status, but in which the function of inter-individual variation is unknown. We collected digital images of the faces of 32 rhesus macaque adult females. We assessed mating rates, and consortship by males, according to female face coloration. We also assessed whether female coloration was linked to physical (skinfold fat, BMI) or physiological (fecal glucocorticoid metabolite fGCM, urinary C-peptide concentrations) condition. We found that redder-faced females were mated more frequently, and consorted for longer periods by top-ranked males. Redder females had higher fGCM concentrations, perhaps related to their increased mating activity and consequent energy mobilization, and blood-flow. Prior analyses have shown that female facial redness is a heritable trait, and that redder-faced females have higher annual fecundity, while other evidence suggests that color expression is likely to be a signal rather than a cue. Collectively, the available evidence suggests that female coloration has evolved at least in part via male mate choice. Its evolution as a sexually-selected ornament attractive to males is probably attributable to the high female reproductive synchrony found in this species.
Red coloration and the evolution of aposematism in arboreal sciurids
<p>An animal's coloration is associated with a variety of processes and is therefore subjected to multiple selective pressures. Mammals, especially, are typically inconspicuously colored, or cryptic, to avoid detection by predators. Alternatively, an animal may use conspicuous coloration to advertise the presence of an anti-predator defense. The association between signal and defense is called aposematism. Conspicuous black and white coloration has recently been associated with a range of defenses in mammals, including body size (Howell et al. 2021), however red coloration as a potentially aposematic signal has yet to be investigated in mammals. Squirrels, like most mammals, are unable to perceive red for use as a social signal. Here we use a comparative framework to test whether redness could be a means of background matching, serve a thermoregulatory function or be an honest warning of anti-predator defenses across a global distribution of tree squirrels which vary in size from 16g to 2.2kg in this study. We measured redness of the dorsum, the venter and of red accents of study skin specimens of 57 tree squirrel species (N=257) representing 25 genera. We then associated these phenotypic variables with environmental variables using phylogenetic generalized least squares regression. We find that increasing dorsal redness is associated with more humid environments and closed canopies, consistent with prior work that coloration on this body region under selection for crypsis (Sheets and Chavez 2020). However, we find that ventral redness and maximum redness is associated with large body sizes. Our findings suggest that crypsis and aposematism are not mutually exclusive, and that aposematism may be more widespread in mammals than is currently appreciated.<u></u></p>
On following pages: 626. Niceforo Maria's Oldfield Mouse (Thomasomys nicefori); 627. Popayan Oldfield Mouse (Thomasomys popayanus); 628. Short-faced Oldfield Mouse (Thomasomys baeops); 629. Snow-footed Oldfield Mouse (Thomasomys niveipes); 630. Principal Oldfield Mouse (Thomasomys princeps); 631. Silky Oldfield Mouse (Thomasomys bombycinus); 632. Red Andean Oldfield Mouse (Thomasomys auricularis); 633. Cinnamon-colored Oldfield Mouse (Thomasomys cinnameus); 634. Central Andes Oldfield Mouse (Thomasomys contradictus); 635. Ashy-bellied Oldfield Mouse (Thomasomys cinereiventen; 636. Colombian Oldfield Mouse (Thomasomys dispar); 637. Soft-furred Oldfield Mouse (Thomasomys lanigen); 638. Ash-colored Oldfield Mouse (Thomasomys cinereus); 639. Wandering Oldfield Mouse (Thomasomys erro); 640. Paramo Oldfield Mouse (Thomasomys paramorum); 641. Forest Oldfield Mouse (Thomasomyssilvestris); 642. Smoky Oldfield Mouse (Thomasomys fumeus); 643. Pichincha Oldfield Mouse (Thomasomys vulcani); 644. Ucucha Oldfield Mouse (Thomasomys ucucha); 645. Taczanowski's Oldfield Mouse (Thomasomys taczanowskii); 646. Golden Oldfield Mouse (Thomasomys aureus); 647. White-tipped Oldfield Mouse (Thomasomys caudivarius); 648. Hudson's Oldfield Mouse (Thomasomys hudsoni); 649. Reddish-backed Oldfield Mouse (Thomasomys pyrrhonotus); 650. Montane Oldfield Mouse (Thomasomys oreas); 651. Cajamarca Oldfield Mouse (Thomasomys praetor; 652. Distinguished Oldfield Mouse (Thomasomys notatus); 653. Apeco Oldfield Mouse (Thomasomys apeco); 654. Peruvian Oldfield Mouse (Thomasomys eleusis); 655. Strong-tailed Oldfield Mouse (Thomasomys ischyrus); 656. Reddish-nosed Oldfield Mouse (Thomasomys rosalinda); 657. Large-eared Oldfield Mouse (Thomasomys macrotis); 658. Ashaninka Oldfield Mouse (Thomasomys onkiro); 659. Inca Oldfield Mouse (Thomasomys incanus), 660. Kalinowski's Oldfield Mouse (Thomasomys kalinowskii); 661. Slender Oldfield Mouse (Thomasomys gracilis); 662. Daphne's Oldfield Mouse (Thomasomys daphne); 663. Anderson's Oldfield Mouse (Thomasomys anderson); 664. Austral Oldfield Mouse (Thomasomys australis); 665. Ladew's Oldfield Mouse (Thomasomys ladewi). in Cricetidae
On following pages: 626. Niceforo Maria's Oldfield Mouse (Thomasomys nicefori); 627. Popayan Oldfield Mouse (Thomasomys popayanus); 628. Short-faced Oldfield Mouse (Thomasomys baeops); 629. Snow-footed Oldfield Mouse (Thomasomys niveipes); 630. Principal Oldfield Mouse (Thomasomys princeps); 631. Silky Oldfield Mouse (Thomasomys bombycinus); 632. Red Andean Oldfield Mouse (Thomasomys auricularis); 633. Cinnamon-colored Oldfield Mouse (Thomasomys cinnameus); 634. Central Andes Oldfield Mouse (Thomasomys contradictus); 635. Ashy-bellied Oldfield Mouse (Thomasomys cinereiventen; 636. Colombian Oldfield Mouse (Thomasomys dispar); 637. Soft-furred Oldfield Mouse (Thomasomys lanigen); 638. Ash-colored Oldfield Mouse (Thomasomys cinereus); 639. Wandering Oldfield Mouse (Thomasomys erro); 640. Paramo Oldfield Mouse (Thomasomys paramorum); 641. Forest Oldfield Mouse (Thomasomyssilvestris); 642. Smoky Oldfield Mouse (Thomasomys fumeus); 643. Pichincha Oldfield Mouse (Thomasomys vulcani); 644. Ucucha Oldfield Mouse (Thomasomys ucucha); 645. Taczanowski's Oldfield Mouse (Thomasomys taczanowskii); 646. Golden Oldfield Mouse (Thomasomys aureus); 647. White-tipped Oldfield Mouse (Thomasomys caudivarius); 648. Hudson's Oldfield Mouse (Thomasomys hudsoni); 649. Reddish-backed Oldfield Mouse (Thomasomys pyrrhonotus); 650. Montane Oldfield Mouse (Thomasomys oreas); 651. Cajamarca Oldfield Mouse (Thomasomys praetor; 652. Distinguished Oldfield Mouse (Thomasomys notatus); 653. Apeco Oldfield Mouse (Thomasomys apeco); 654. Peruvian Oldfield Mouse (Thomasomys eleusis); 655. Strong-tailed Oldfield Mouse (Thomasomys ischyrus); 656. Reddish-nosed Oldfield Mouse (Thomasomys rosalinda); 657. Large-eared Oldfield Mouse (Thomasomys macrotis); 658. Ashaninka Oldfield Mouse (Thomasomys onkiro); 659. Inca Oldfield Mouse (Thomasomys incanus), 660. Kalinowski's Oldfield Mouse (Thomasomys kalinowskii); 661. Slender Oldfield Mouse (Thomasomys gracilis); 662. Daphne's Oldfield Mouse (Thomasomys daphne); 663. Anderson's Oldfield Mouse (Thomasomys anderson); 664. Austral Oldfield Mouse (Thomasomys australis); 665. Ladew's Oldfield Mouse (Thomasomys ladewi).
FIGURE. Drosera hirtella (a–i): a, b, c, habit of the "type morphotype"; d, habit of the "western morphotype"; e, rosette of the "type morphotype"; f, emerging inflorescence, highlighting the red scape with red eglandular trichomes characteristic of the species; g, fertile individuals of D. hirtella (left plant, with inflorescence emerging to the bottom) and D. lutescens (right plant, with inflorescence emerging to the top left) growing under shaded conditions side by side, highlighting the morphological differences between the two species regarding leaf shape and scape and indumentum color; h, i, flower. a–c, f and h at Serra do Cipó, MG; d and h at Chapada dos Veadeiros, GO; e at Diamantina, MG; g at Cristalina, GO. Photo credits: all by PMG. in A synopsis of the genus Drosera (Droseraceae) in Brazil
FIGURE. Drosera hirtella (a–i): a, b, c, habit of the "type morphotype"; d, habit of the "western morphotype"; e, rosette of the "type morphotype"; f, emerging inflorescence, highlighting the red scape with red eglandular trichomes characteristic of the species; g, fertile individuals of D. hirtella (left plant, with inflorescence emerging to the bottom) and D. lutescens (right plant, with inflorescence emerging to the top left) growing under shaded conditions side by side, highlighting the morphological differences between the two species regarding leaf shape and scape and indumentum color; h, i, flower. a–c, f and h at Serra do Cipó, MG; d and h at Chapada dos Veadeiros, GO; e at Diamantina, MG; g at Cristalina, GO. Photo credits: all by PMG.
Deccan region, Madras, India. Genus Vandeleuria is masculine, so widely used specific name oleracea has been changed for gender agreement. Vandeleuria oleraceusis possibly a composite of species. Polytypic, but subspecific taxonomy requires reassessment. Distribution. Widespread in S Asia (India, Nepal, Bhutan, Bangladesh, and Sri Lan-ka), S China (W & S Yunnan), and mainland SE Asia N of the Isthmus of Kra. Descriptive notes. Head-body 68 mm, tail 105 mm, ear 13 mm, hindfoot 17 mm; weight 10 g. The Indomalayan Long-tailed Climbing Mouse is small, with flat nail on outer finger and outertoe; tail is slender, brown, twice as long as head-body length, and lacks distal tuft. Dorsal pelageis silky and salmon in color; venter is white, with fulvous hues. Habitat. Tall cane and tangled vines in primary and secondary forest such as bamboo forest, moist deciduous forest, temperate forests, montane wet zone, and disturbed secondary forests, and perhaps agricultural areas at elevations of 150-1500 m. Food and Feeding. Indomalayan [Long-tailed Climbing Mice eat fruits, buds, and flowers. Breeding. Litters of the Indomalayan Long-tailed Climbing Mouse have 3-6 young. Activity patterns. Indomalayan Long-tailed Climbing Mice are arboreal and nocturnal, although one individual was caught duringthe day. Movements, Home range and Social organization. Indomalayan Long-tailed Climbing Mice build nests in tall bushes or cane to rear their young. Status and Conservation. Classified as Least Concern on The IUCN Red Last (as V. olacea). The Indomalayan Long-tailed Climbing Mouse occurs in several habitats and a wide distribution that includes national parks. Further taxonomical studies are required to assess conservation status ofthis potentially diverse species complex. Bibliography. Corbet & Hill (1992), Dang Huy Huynh et al. (1994), Ellerman (1941), Marshall (1977b), Musser & Carleton (2005), Osgood (1932), Phillips (1980), Wang Yingxiang (2003). in Muridae
Deccan region, Madras, India. Genus Vandeleuria is masculine, so widely used specific name oleracea has been changed for gender agreement. Vandeleuria oleraceusis possibly a composite of species. Polytypic, but subspecific taxonomy requires reassessment. Distribution. Widespread in S Asia (India, Nepal, Bhutan, Bangladesh, and Sri Lan-ka), S China (W & S Yunnan), and mainland SE Asia N of the Isthmus of Kra. Descriptive notes. Head-body 68 mm, tail 105 mm, ear 13 mm, hindfoot 17 mm; weight 10 g. The Indomalayan Long-tailed Climbing Mouse is small, with flat nail on outer finger and outertoe; tail is slender, brown, twice as long as head-body length, and lacks distal tuft. Dorsal pelageis silky and salmon in color; venter is white, with fulvous hues. Habitat. Tall cane and tangled vines in primary and secondary forest such as bamboo forest, moist deciduous forest, temperate forests, montane wet zone, and disturbed secondary forests, and perhaps agricultural areas at elevations of 150-1500 m. Food and Feeding. Indomalayan [Long-tailed Climbing Mice eat fruits, buds, and flowers. Breeding. Litters of the Indomalayan Long-tailed Climbing Mouse have 3-6 young. Activity patterns. Indomalayan Long-tailed Climbing Mice are arboreal and nocturnal, although one individual was caught duringthe day. Movements, Home range and Social organization. Indomalayan Long-tailed Climbing Mice build nests in tall bushes or cane to rear their young. Status and Conservation. Classified as Least Concern on The IUCN Red Last (as V. olacea). The Indomalayan Long-tailed Climbing Mouse occurs in several habitats and a wide distribution that includes national parks. Further taxonomical studies are required to assess conservation status ofthis potentially diverse species complex. Bibliography. Corbet & Hill (1992), Dang Huy Huynh et al. (1994), Ellerman (1941), Marshall (1977b), Musser & Carleton (2005), Osgood (1932), Phillips (1980), Wang Yingxiang (2003).
On following pages: 347. Plains Mouse (Pseudomys australis); 348. Bolam''s Mouse (Pseudomys bolami); 349. Kakadu Pebble Mouse (Pseudomys calabyi); 350. Western Pebble Mouse (Pseudomys chapmani); 351. Desert Mouse (Pseudomys desertor); 352. Shark Bay Mouse (Pseudomys field); 353. Australian Smoky Mouse (Pseudomys fumeus); 354. Eastern Chestnut Mouse (Pseudomys gracilicaudatus); 355. Sandy Inland Mouse (Pseudomys hermannsburgensis); 356. Long-tailed Mouse (Pseudomys higginsi); 357. Central Pebble Mouse (Pseudomys johnson); 358. Western Chestnut Mouse (Pseudomys nanus); 359. New Holland Mouse (Pseudomys novaehollandiae); 360. Western Mouse (Pseudomys occidentalis); 361. Hastings River Mouse (Pseudomys oralis); 362. Eastern Pebble Mouse (Pseudomys patrius); 363. Heath Mouse (Pseudomys shortridgel); 364. Common Australian Rock Rat (Zyzomys argurus); 365. Arnhem Land Rock Rat (Zyzomys maini); 366. Carpentarian Rock Rat (Zyzomys palatalis); 367. Central Australian Rock Rat (Zyzomys pedunculatus); 368. Kimberley Rock Rat (Zyzomys woodward); 369. Malayan Tree Rat (Pithecheir parvus); 370. Red Tree Rat (Pithecheir melanurus); 371. Bornean Tree Rat (Pithecheirops otion); 372. Cutch Rat (Cremnomys cutchicus); 373. Elvira Rat (Cremnomys elvira); 374. Crump's Rat (Diomys crumpi); 375. White-tailed Wood Rat (Madromys blanfordi); 376. Sand-colored Soft-furred Rat (Millardia gleadowi); 377. Kondana Soft-furred Rat (Millardia kondana); 378. Common Soft-furred Rat (Millardia in Muridae
On following pages: 347. Plains Mouse (Pseudomys australis); 348. Bolam''s Mouse (Pseudomys bolami); 349. Kakadu Pebble Mouse (Pseudomys calabyi); 350. Western Pebble Mouse (Pseudomys chapmani); 351. Desert Mouse (Pseudomys desertor); 352. Shark Bay Mouse (Pseudomys field); 353. Australian Smoky Mouse (Pseudomys fumeus); 354. Eastern Chestnut Mouse (Pseudomys gracilicaudatus); 355. Sandy Inland Mouse (Pseudomys hermannsburgensis); 356. Long-tailed Mouse (Pseudomys higginsi); 357. Central Pebble Mouse (Pseudomys johnson); 358. Western Chestnut Mouse (Pseudomys nanus); 359. New Holland Mouse (Pseudomys novaehollandiae); 360. Western Mouse (Pseudomys occidentalis); 361. Hastings River Mouse (Pseudomys oralis); 362. Eastern Pebble Mouse (Pseudomys patrius); 363. Heath Mouse (Pseudomys shortridgel); 364. Common Australian Rock Rat (Zyzomys argurus); 365. Arnhem Land Rock Rat (Zyzomys maini); 366. Carpentarian Rock Rat (Zyzomys palatalis); 367. Central Australian Rock Rat (Zyzomys pedunculatus); 368. Kimberley Rock Rat (Zyzomys woodward); 369. Malayan Tree Rat (Pithecheir parvus); 370. Red Tree Rat (Pithecheir melanurus); 371. Bornean Tree Rat (Pithecheirops otion); 372. Cutch Rat (Cremnomys cutchicus); 373. Elvira Rat (Cremnomys elvira); 374. Crump's Rat (Diomys crumpi); 375. White-tailed Wood Rat (Madromys blanfordi); 376. Sand-colored Soft-furred Rat (Millardia gleadowi); 377. Kondana Soft-furred Rat (Millardia kondana); 378. Common Soft-furred Rat (Millardia
Distribution. Now restricted to the Channel Country of SW Queensland and the Lake Eyre Basin in NE South Australia. Descriptive notes. Head-body 95-120 mm, tail 105-160 mm, ear 23-29 mm, hindfoot 32-37 mm; weight 30-50 g. The Fawn Hopping Mouse has body form typical of hopping mice, with very long hindfeet, long tail with distal brush of longer hairs, very long ears, and large protruberant eyes. Dorsal fur is of variable color, from pale pinkish fawn to gray; ventral fur white. Unlike most other hopping mice, it has no throat pouch, but males have a glandular area of naked skin on the chest. Habitat. Occurs in low shrublands and tussock grasslands on stony ("gibber") plains and claypans. Shows marked habitat segregation from the Dusky Hopping Mouse (N. fuscus), which is closely associated with sandy substrates. Food and Feeding. The Fawn Hopping Mouse is mostly granivorous, but also eats other plant material (stems, leaves) and occasionally invertebrates. It uses succulent, salt-adapted plants around edges of claypans as a source of water. Breeding. Reproduction is probably largely opportunistic and aseasonal, with high reproductive output from near-continuous breeding after periods of high rainfall; reported littersize is 1-5, most commonly three; gestation period 38-43 days for nonlactating females. Females may mature later than other hopping mice, with reproductive maturity reached at about six months. Activity patterns. Terrestrial and nocturnal. Fawn Hopping Mice shelter during day in burrow systems that are typically simpler and shallower than those of other hopping mice. Movements, Home range and Social organization. Fawn Hopping Mice generally live singly or in small groups; typically uncommon within range, but population density may increase by an order of magnitude following periods of high rainfall. Status and Conservation. Classified as Near Threatened on The IUCN Red List. The Fawn Hopping Mouse has shown marked decline in range (estimated at greater than 50%), and presumably population size, since European settlement of Australia. This is mostlikely due to predation by the introduced house cat and Red Fox (Vulpes vulpes), and to habitat degradation associated with pastoralism. Bibliography. Brazenor (1934), Burbidge et al. (2008), Finlayson (1939), Gould (1853), Jackson & Groves (2015), Murray et al. (1999), Ogilby (1892), Thomas (1921h), Van Dyck & Strahan (2008), Waite (1898), Watts & Aslin (1981), Woinarski et al. (2014), Wood Jones (1925). in Muridae
Distribution. Now restricted to the Channel Country of SW Queensland and the Lake Eyre Basin in NE South Australia. Descriptive notes. Head-body 95-120 mm, tail 105-160 mm, ear 23-29 mm, hindfoot 32-37 mm; weight 30-50 g. The Fawn Hopping Mouse has body form typical of hopping mice, with very long hindfeet, long tail with distal brush of longer hairs, very long ears, and large protruberant eyes. Dorsal fur is of variable color, from pale pinkish fawn to gray; ventral fur white. Unlike most other hopping mice, it has no throat pouch, but males have a glandular area of naked skin on the chest. Habitat. Occurs in low shrublands and tussock grasslands on stony ("gibber") plains and claypans. Shows marked habitat segregation from the Dusky Hopping Mouse (N. fuscus), which is closely associated with sandy substrates. Food and Feeding. The Fawn Hopping Mouse is mostly granivorous, but also eats other plant material (stems, leaves) and occasionally invertebrates. It uses succulent, salt-adapted plants around edges of claypans as a source of water. Breeding. Reproduction is probably largely opportunistic and aseasonal, with high reproductive output from near-continuous breeding after periods of high rainfall; reported littersize is 1-5, most commonly three; gestation period 38-43 days for nonlactating females. Females may mature later than other hopping mice, with reproductive maturity reached at about six months. Activity patterns. Terrestrial and nocturnal. Fawn Hopping Mice shelter during day in burrow systems that are typically simpler and shallower than those of other hopping mice. Movements, Home range and Social organization. Fawn Hopping Mice generally live singly or in small groups; typically uncommon within range, but population density may increase by an order of magnitude following periods of high rainfall. Status and Conservation. Classified as Near Threatened on The IUCN Red List. The Fawn Hopping Mouse has shown marked decline in range (estimated at greater than 50%), and presumably population size, since European settlement of Australia. This is mostlikely due to predation by the introduced house cat and Red Fox (Vulpes vulpes), and to habitat degradation associated with pastoralism. Bibliography. Brazenor (1934), Burbidge et al. (2008), Finlayson (1939), Gould (1853), Jackson & Groves (2015), Murray et al. (1999), Ogilby (1892), Thomas (1921h), Van Dyck & Strahan (2008), Waite (1898), Watts & Aslin (1981), Woinarski et al. (2014), Wood Jones (1925).
Figure S1: Two-tailed intergroup Pearson analysis between patient age, COVID-19 detection, de-livery time, gravida, and termination week (Y-axis) with maternal and neonatal anti-COVID-19 immunoglobulins (IgG and IgM) (X-axis), depicted as a three-point color map (lowest = blue, 0 = white, highest = red) showed no significant correlation (at p>0.05).
<p>Figure S1: Two-tailed intergroup Pearson analysis between patient age, COVID-19 detection, de-livery time, gravida, and termination week (Y-axis) with maternal and neonatal anti-COVID-19 immunoglobulins (IgG and IgM) (X-axis), depicted as a three-point color map (lowest = blue, 0 = white, highest = red) showed no significant correlation (at p>0.05).</p>
FIGURE 12. Salmacis spp. A. S. bicolor. Red color variant. B. S in New Ecological Observations and Occurrence for Asteroidea and Echinoidea in Hong Kong
FIGURE 12. Salmacis spp. A. S. bicolor. Red color variant. B. S. sphaeroides. In situ observation of S. sphaeroides. C. S. sphaeroides. Showing individuals scavenging on a dead fish. Photographer: A. Josy Lai; B. and C. Sam King Fung Yiu.
ScienceDex guides
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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.