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FIGURE. Images of representative members of tribe Phyllantheae. (A) Flowers of Nellica maderaspatensis. (B) Pistillate flowers of Cathetus gracilis, note the unique disc covering the ovary. (C) Pistillate and staminate flowers of Cathetus glaucophyllus. (D) Staminate flowers of Nymphanthus glaucescens. (E) Fruits of Kirganelia muelleriana. (F) Fruits of Lysiandra subcrenulata. (G) Phylloclade with flowers of Phyllanthus angustifolius. (H) Flowering branchlet of Phyllanthus incrustatus, note the ornamentation on the axes. (I) Habit of Moeroris tenella. (J) Fruiting branch of Dendrophyllanthus tenuirhachis. (K) Fruits of Cicca profusa. (L) Fruiting branch of Emblica officinalis. (M) Flowering plant of Emblica urinaria. (N) Pistillate flower of Breynia disticha. (O) Staminate flower of Breynia disticha. (P) flower of Glochidion dunnianum. (Q) Staminate of Glochidion lanceolarium. (R) Dehisced capsule of Glochidion sp. showing seeds covered with a red sarcotesta. Photos: A & F by J.J. Bruhl; B & P by M.S. Nuraliev; C by T. Williams; E & K by C. Jongkind; H by B. Falcón; J by R.-Y. Yu; D, G, I, L, M, N, O, Q & R by R.W.Bouman. in A revised phylogenetic classification of tribe Phyllantheae (Phyllanthaceae)
FIGURE. Images of representative members of tribe Phyllantheae. (A) Flowers of Nellica maderaspatensis. (B) Pistillate flowers of Cathetus gracilis, note the unique disc covering the ovary. (C) Pistillate and staminate flowers of Cathetus glaucophyllus. (D) Staminate flowers of Nymphanthus glaucescens. (E) Fruits of Kirganelia muelleriana. (F) Fruits of Lysiandra subcrenulata. (G) Phylloclade with flowers of Phyllanthus angustifolius. (H) Flowering branchlet of Phyllanthus incrustatus, note the ornamentation on the axes. (I) Habit of Moeroris tenella. (J) Fruiting branch of Dendrophyllanthus tenuirhachis. (K) Fruits of Cicca profusa. (L) Fruiting branch of Emblica officinalis. (M) Flowering plant of Emblica urinaria. (N) Pistillate flower of Breynia disticha. (O) Staminate flower of Breynia disticha. (P) flower of Glochidion dunnianum. (Q) Staminate of Glochidion lanceolarium. (R) Dehisced capsule of Glochidion sp. showing seeds covered with a red sarcotesta. Photos: A & F by J.J. Bruhl; B & P by M.S. Nuraliev; C by T. Williams; E & K by C. Jongkind; H by B. Falcón; J by R.-Y. Yu; D, G, I, L, M, N, O, Q & R by R.W.Bouman.
Subspecies and Distribution. F. s. silvestris Schreber, 1777 — Europe E to the Carpathian Mts and the River Dnieper N of the Black Sea. F.s. brockmani Pocock, 1944 — Somalia. F. s. cafra Desmarest, 1822 — Zimbabwe, S Mozambique and South Africa. F. s. caucasica Satunin, 1905 — Caucasus Mts and Turkey. F.s. caudata Gray, 1874 — deserts E Caspian Sea to NW China (Xinjiang) and Mongolia. F. s. foxi Pocock, 1944 — Senegal to Lake Chad. F.s. gordoni Harrison, 1968 — Batinah coast of Oman. F.s. grampia G. S. Miller, 1907 — N Scotland. F. s. griselda Thomas, 1926 — Kalahari region to S Angola. F. s. iraki Cheesman, 1920 — Arabian Desert regions. F. s. lybica Forster, 1780 — desert regions of N Africa to Sudan and N Niger. F.s. melland: Schwann, 1904 — SC Africa. F.s. nesterovi Biurla, 1916 — Mesopotamian region to SW Iran. F. s. ocreata Gmelin, 1791 — Ethiopian highlands. F. s. ornata Gray, 1830 — India. Probably W through Iran. F.s. pyrrhus Pocock, 1944 — N Angola and SW Zaire. F. s. sarda Lataste, 1885 — coastal Maghreb region of Morocco and Algeria. F.s. tristrami Pocock, 1944 — Palestine and Red Sea coast of Arabia. F.s. ugandae Schwann, 1904 — E Africa. in Felidae
Subspecies and Distribution. F. s. silvestris Schreber, 1777 — Europe E to the Carpathian Mts and the River Dnieper N of the Black Sea. F.s. brockmani Pocock, 1944 — Somalia. F. s. cafra Desmarest, 1822 — Zimbabwe, S Mozambique and South Africa. F. s. caucasica Satunin, 1905 — Caucasus Mts and Turkey. F.s. caudata Gray, 1874 — deserts E Caspian Sea to NW China (Xinjiang) and Mongolia. F. s. foxi Pocock, 1944 — Senegal to Lake Chad. F.s. gordoni Harrison, 1968 — Batinah coast of Oman. F.s. grampia G. S. Miller, 1907 — N Scotland. F. s. griselda Thomas, 1926 — Kalahari region to S Angola. F. s. iraki Cheesman, 1920 — Arabian Desert regions. F. s. lybica Forster, 1780 — desert regions of N Africa to Sudan and N Niger. F.s. melland: Schwann, 1904 — SC Africa. F.s. nesterovi Biurla, 1916 — Mesopotamian region to SW Iran. F. s. ocreata Gmelin, 1791 — Ethiopian highlands. F. s. ornata Gray, 1830 — India. Probably W through Iran. F.s. pyrrhus Pocock, 1944 — N Angola and SW Zaire. F. s. sarda Lataste, 1885 — coastal Maghreb region of Morocco and Algeria. F.s. tristrami Pocock, 1944 — Palestine and Red Sea coast of Arabia. F.s. ugandae Schwann, 1904 — E Africa.
On following pages: 368. Himalayan Broad-muzzled Bat (Submyotodon caliginosus); 369. Moupin Broad-muzzled Bat (Submyotodon moupinensis); 370. Northern Myotis (Myotis septentrionalis); 371. South-western Myotis (Myotis auriculus); 372. Western Small-footed Myotis (Myotis ciliolabrum); 373. Dark-nosed Small-footed Myotis (Myotis melanorhinus); 374. Eastern Small-footed Myotis (Myotis leibil); 375. California Myotis (Myotis californicus); 376. Little Brown Myotis (Myotis lucifugus); 377. Keen's Myotis (Myotis keenii); 378. Long-eared Myotis (Myotis evotis); 379. Fringed Myotis (Myotis thysanodes); 380. Arizona Myotis (Myotis occultus); 381. Longlegged Myotis (Myotis volans); 382. Flat-headed Myotis (Myotis planiceps); 383. Indiana Myotis (Myotis sodalis); 384. Cinnamon Myotis (Myotis fortidens); 385. Findley's Myotis (Myotis findley)); 386. Northern Hairy-legged Myotis (Myotis pilosatibialis); 387. Southern Hairy-legged Myotis (Myotis keaysi); 388. Red Myotis (Myotis ruber); 389. Riparian Myotis (Myotis riparius); 390. Velvety Myotis (Myotis simus); 391. Golden Myotis (Myotis midastactus); 392. Elegant Myotis (Myotis elegans); 393. Fish-eating Myotis (Myotis vives). in Vespertilionidae
On following pages: 368. Himalayan Broad-muzzled Bat (Submyotodon caliginosus); 369. Moupin Broad-muzzled Bat (Submyotodon moupinensis); 370. Northern Myotis (Myotis septentrionalis); 371. South-western Myotis (Myotis auriculus); 372. Western Small-footed Myotis (Myotis ciliolabrum); 373. Dark-nosed Small-footed Myotis (Myotis melanorhinus); 374. Eastern Small-footed Myotis (Myotis leibil); 375. California Myotis (Myotis californicus); 376. Little Brown Myotis (Myotis lucifugus); 377. Keen's Myotis (Myotis keenii); 378. Long-eared Myotis (Myotis evotis); 379. Fringed Myotis (Myotis thysanodes); 380. Arizona Myotis (Myotis occultus); 381. Longlegged Myotis (Myotis volans); 382. Flat-headed Myotis (Myotis planiceps); 383. Indiana Myotis (Myotis sodalis); 384. Cinnamon Myotis (Myotis fortidens); 385. Findley's Myotis (Myotis findley)); 386. Northern Hairy-legged Myotis (Myotis pilosatibialis); 387. Southern Hairy-legged Myotis (Myotis keaysi); 388. Red Myotis (Myotis ruber); 389. Riparian Myotis (Myotis riparius); 390. Velvety Myotis (Myotis simus); 391. Golden Myotis (Myotis midastactus); 392. Elegant Myotis (Myotis elegans); 393. Fish-eating Myotis (Myotis vives).
On following pages: 97. Savi's Pipistrelle (Hypsugo savii); 98. Alashanian Pipistrelle (Hypsugo alaschanicus); 99. Cadorna's Pipistrelle (Hypsugo cadornae); 100. Joffre's Pipistrelle (Hypsugo joffrel); 101. Chinese Pipistrelle (Hypsugo pulveratus); 102. Chocolate Pipistrelle (Hypsugo affinis); 103. Pungent Pipistrelle (Hypsugo mordax); 104. Peters's Pipistrelle (Hypsugo peters); 105. Long-toothed Pipistrelle (Hypsugo dolichodon); 106. Burmese Pipistrelle (Hypsugo lophurus); 107. Red-brown Pipistrelle (Hypsugo kitcheneri); 108. Brown Pipistrelle (Hypsugo imbricatus); 109. Big-eared Pipistrelle (Hypsugo macrotis); 110. Vordermann's Pipistrelle (Hypsugo vordermanni), 111. Anchieta's Pipistrelle (Hypsugo anchieta); 112. Kirindy Pipistrelle (Hypsugo bemainty); 113. Mouse-like Pipistrelle (Hypsugo musciculus); 114. Broad-headed Pipistrelle (Hypsugo crassulus); 115. Eisentraut's Pipistrelle (Hypsugo eisentrautl); 116. Schlieffen's Bat (Nycticeinops schlieffenii); 117. Moloney's Mimic Bat (Mimetillus moloney)). in Vespertilionidae
On following pages: 97. Savi's Pipistrelle (Hypsugo savii); 98. Alashanian Pipistrelle (Hypsugo alaschanicus); 99. Cadorna's Pipistrelle (Hypsugo cadornae); 100. Joffre's Pipistrelle (Hypsugo joffrel); 101. Chinese Pipistrelle (Hypsugo pulveratus); 102. Chocolate Pipistrelle (Hypsugo affinis); 103. Pungent Pipistrelle (Hypsugo mordax); 104. Peters's Pipistrelle (Hypsugo peters); 105. Long-toothed Pipistrelle (Hypsugo dolichodon); 106. Burmese Pipistrelle (Hypsugo lophurus); 107. Red-brown Pipistrelle (Hypsugo kitcheneri); 108. Brown Pipistrelle (Hypsugo imbricatus); 109. Big-eared Pipistrelle (Hypsugo macrotis); 110. Vordermann's Pipistrelle (Hypsugo vordermanni), 111. Anchieta's Pipistrelle (Hypsugo anchieta); 112. Kirindy Pipistrelle (Hypsugo bemainty); 113. Mouse-like Pipistrelle (Hypsugo musciculus); 114. Broad-headed Pipistrelle (Hypsugo crassulus); 115. Eisentraut's Pipistrelle (Hypsugo eisentrautl); 116. Schlieffen's Bat (Nycticeinops schlieffenii); 117. Moloney's Mimic Bat (Mimetillus moloney)).
On following pages: 252. Northern Hoary Bat (Lasiurus cinereus); 253. Hawaiian Hoary Bat (Lasiurus semotus); 254. Eastern Red Bat (Lasiurus borealis); 255. Pfeiffer's Red Bat (Lasiurus pfeiffer); 256. Seminole Bat (Lasiurus seminolus); 257. Greater Red Bat (Lasiurus atratus); 258. Southern Red Bat (Lasiurus blossevilil); 259. Desert Red Bat (Lasiurus frantzi); 260. Cinnamon Red Bat (Lasiurus varius); 261. Minor Red Bat (Lasiurus minon; 262. Tacarcunan Bat (Lasiurus castaneus); 263. Hairy-tailed Bat (Lasiurus ebenus); 264. Jamaican Red Bat (Lasiurus degelidus); 265. Western Yellow Bat (Lasiurus xanthinus); 266. Northern Yellow Bat (Lasiurus intermedius); 267. Cuban Yellow Bat (Lasiurus insularis); 268. Southern Yellow Bat (Lasiurus egal). in Vespertilionidae
On following pages: 252. Northern Hoary Bat (Lasiurus cinereus); 253. Hawaiian Hoary Bat (Lasiurus semotus); 254. Eastern Red Bat (Lasiurus borealis); 255. Pfeiffer's Red Bat (Lasiurus pfeiffer); 256. Seminole Bat (Lasiurus seminolus); 257. Greater Red Bat (Lasiurus atratus); 258. Southern Red Bat (Lasiurus blossevilil); 259. Desert Red Bat (Lasiurus frantzi); 260. Cinnamon Red Bat (Lasiurus varius); 261. Minor Red Bat (Lasiurus minon; 262. Tacarcunan Bat (Lasiurus castaneus); 263. Hairy-tailed Bat (Lasiurus ebenus); 264. Jamaican Red Bat (Lasiurus degelidus); 265. Western Yellow Bat (Lasiurus xanthinus); 266. Northern Yellow Bat (Lasiurus intermedius); 267. Cuban Yellow Bat (Lasiurus insularis); 268. Southern Yellow Bat (Lasiurus egal).
Data from: Blood-red colour as a prey-choice cue for mosquito specialist predators
<p>Specialist predators are innately and distinctively proficient at targeting specific prey types. This is enabled by behavioural, perceptual, and cognitive mechanisms that can only be understood using carefully-designed experiments. <i>Evarcha culicivora </i>is<i> </i>an East African jumping spider that feeds on vertebrate blood acquired indirectly by actively targeting blood-carrying female mosquitoes as preferred prey. Here we asked whether these spiders use the colour red to locate this prey. In Objective 1, we used spectrophotometry to document the changing redness of mosquitoes in the 17 hours following blood meals. In Objective 2, we used vision-based choice tests to document how <i>E</i>. <i>culicivora</i>'s preference for blood-carrying mosquitoes changes over time (using an interval comparable to that in Objective 1). We found that <i>E. culicivora</i> exhibits the strongest preference for blood-carrying mosquitoes during the first 6 hours after those mosquitoes had fed on blood and that this time interval corresponded to when mosquitoes exhibit the reddest coloration. Based on this, we then took mosquitoes that had never fed on blood and manipulated their colour using a blend of red food dye to make them appear blood-fed (i.e., by matching their spectral properties to mosquitoes that had fed on blood within the previous 6 hours). We also created grey-dyed mosquitoes that matched the spectral properties of those that had never fed on blood. In Objective 3, we found that <i>E. culicivora</i> did not visually discriminate between our red-dyed mosquitoes and blood-carrying mosquitoes; this indicated that, to <i>E. culicivora</i>, our red-dyed mosquitoes appropriately resembled blood-carrying mosquitoes. Finally, in Objective 4, we showed that <i>E. culicivora</i> consistently preferred red-dyed mosquitoes over grey-dyed mosquitoes, supporting the hypothesis that <i>E. culicivora</i> uses some aspect of red coloration to locate their preferred prey. We discuss how these findings relate to cognitive processes, colour-based communication, and sensory exploitation. </p>
On following pages: 130. Sulawesi Flying Fox (Acerodon celebensis); 131. Talaud Flying Fox (Acerodon humilis); 132. Golden-capped Flying Fox (Acerodon jubatus); 133. Palawan Flying Fox (Acerodon leucotis): 134. Sunda Flying Fox (Acerodon macklotil); 135. Moluccan Masked Flying Fox (Pteropus personatus); 136. Lombok Flying Fox (Pteropus lombocensis); 137. Little Red Flying Fox (Pteropus scapulatus); 138. Big-eared Flying Fox (Pteropus macrotis): 139. Gilliard's Flying Fox (Pteropus gilliardorum); 140. Sanborn's Flying Fox (Pteropus mahaganus); 141. Dwarf Flying Fox (Pteropus woodford); 142. Pohnpei Flying Fox (Pteropus molossinus); 143. Chuuk Flying Fox (Pteropus pelagicus). in Pteropodidae
On following pages: 130. Sulawesi Flying Fox (Acerodon celebensis); 131. Talaud Flying Fox (Acerodon humilis); 132. Golden-capped Flying Fox (Acerodon jubatus); 133. Palawan Flying Fox (Acerodon leucotis): 134. Sunda Flying Fox (Acerodon macklotil); 135. Moluccan Masked Flying Fox (Pteropus personatus); 136. Lombok Flying Fox (Pteropus lombocensis); 137. Little Red Flying Fox (Pteropus scapulatus); 138. Big-eared Flying Fox (Pteropus macrotis): 139. Gilliard's Flying Fox (Pteropus gilliardorum); 140. Sanborn's Flying Fox (Pteropus mahaganus); 141. Dwarf Flying Fox (Pteropus woodford); 142. Pohnpei Flying Fox (Pteropus molossinus); 143. Chuuk Flying Fox (Pteropus pelagicus).
On following pages: 190. Hairy Fruit-eating Bat (Artibeus hirsutus); 191. Fringed Fruit-eating Bat (Artibeus fimbriatus); 192. Ecuadorian Fruit-eating Bat (Artibeus aequatorialis); 193. Jamaican Fruit-eating Bat (Artibeus jamaicensis); 194. Dark Fruit-eating Bat (Artibeus obscurus); 195. Schwartz's Fruit-eating Bat (Artibeus schwartz); 196. Great Fruit-eating Bat (Artibeus lituratus); 197. Large Fruit-eating Bat (Artibeus amplus); 198. Flat-faced Fruit-eating Bat (Artibeus planirostris); 199. Rosenberg's Fruit-eating Bat (Artibeus rosenbergi); 200. Thomas's Fruit-eating Bat (Artibeus watson); 201. Toltec Fruit-eating Bat (Artibeus toltecus); 202. Pygmy Fruit-eating Bat (Artibeus phaeotis); 203. Gervais's Fruit-eating Bat (Artibeus cinereus); 204. Andersen's Fruit-eating Bat (Artibeus anderseni); 205. Little Fruit-eating Bat (Artibeusravus); 206. Aztec Fruit-eating Bat (Artibeus aztecus); 207. Bogota Fruit-eating Bat (Artibeus bogotensis); 208. Silvery Fruit-eating Bat (Artibeus glaucus); 209. Dwarf Fruit-eating Bat (Artibeus gnomus); 210. Jamaican Fig-eating Bat (Ariteus flavescens); 211. Tree Bat (Ardops nicholls); 212. Red Fruit Bat (Stenoderma rufum); 213. Wrinkle-faced Bat (Centurio senex): 214. Ipanema Broad-nosed Bat (Pygoderma bilabiatum); 215. Visored Bat (Sphaeronycteris toxophyllum); 216. Little White-shouldered Bat (Ametrida centurio); 217. Cuban Fig-eating Bat (Phyllops falcatus). in Phyllostomidae
On following pages: 190. Hairy Fruit-eating Bat (Artibeus hirsutus); 191. Fringed Fruit-eating Bat (Artibeus fimbriatus); 192. Ecuadorian Fruit-eating Bat (Artibeus aequatorialis); 193. Jamaican Fruit-eating Bat (Artibeus jamaicensis); 194. Dark Fruit-eating Bat (Artibeus obscurus); 195. Schwartz's Fruit-eating Bat (Artibeus schwartz); 196. Great Fruit-eating Bat (Artibeus lituratus); 197. Large Fruit-eating Bat (Artibeus amplus); 198. Flat-faced Fruit-eating Bat (Artibeus planirostris); 199. Rosenberg's Fruit-eating Bat (Artibeus rosenbergi); 200. Thomas's Fruit-eating Bat (Artibeus watson); 201. Toltec Fruit-eating Bat (Artibeus toltecus); 202. Pygmy Fruit-eating Bat (Artibeus phaeotis); 203. Gervais's Fruit-eating Bat (Artibeus cinereus); 204. Andersen's Fruit-eating Bat (Artibeus anderseni); 205. Little Fruit-eating Bat (Artibeusravus); 206. Aztec Fruit-eating Bat (Artibeus aztecus); 207. Bogota Fruit-eating Bat (Artibeus bogotensis); 208. Silvery Fruit-eating Bat (Artibeus glaucus); 209. Dwarf Fruit-eating Bat (Artibeus gnomus); 210. Jamaican Fig-eating Bat (Ariteus flavescens); 211. Tree Bat (Ardops nicholls); 212. Red Fruit Bat (Stenoderma rufum); 213. Wrinkle-faced Bat (Centurio senex): 214. Ipanema Broad-nosed Bat (Pygoderma bilabiatum); 215. Visored Bat (Sphaeronycteris toxophyllum); 216. Little White-shouldered Bat (Ametrida centurio); 217. Cuban Fig-eating Bat (Phyllops falcatus).
Data from: Responses to artificial selection for locomotor activity: A focus on death feigning in red flour beetle
<p>Whole-organism performance, including locomotor activity, is an important fitness trait in many animals. Locomotor activity is often classified into sprint speed and locomotor endurance and differences in sprint speed and locomotor endurance affect on other traits such as life-history traits. Previous studies found that locomotor endurance, sprint speed, and brain dopamine (DA) levels are correlated with artificial selection for death-feigning (an anti-predator behavior that we refer to as "death-feigning syndrome") in some insect species. Thus, if the syndrome has a genetic basis, death feigning, sprint speed, and brain DA levels may be affected by artificial selection for locomotor endurance. We artificially selected for locomotor endurance over 10 generations in the red flour beetle <em>Tribolium castaneum</em>, and established higher (H) and lower activity (L) strains, then compared their death-feigning behavior, sprint speed, and brain DA levels. H-strain beetles exhibited significantly shorter duration of death-feigning, and significantly higher sprint speed, suggesting variation in death-feigning syndrome. Surprisingly, although brain DA expression affects various animal behaviors, we found no significant differences in the brain DA expressions of H- and L-strain beetles. Thus, our results imply genetic correlations between locomotor endurance, sprint speed, and death feigning, but not with brain DA expression, suggesting that differences in the biogenic amine results of our previous studies may reflect differences in behavioral expression mechanisms.</p>
FIGURE 1 in The tadpole of the red-bellied toad, Melanophryniscus dorsalis (Mertens, 1933) (Anura: Bufonidae)
FIGURE 1. Tadpole of Melanophryniscus dorsalis, MCN 14349, Stage 31. (A) coloration in life, (B) lateral view, (C) dorsal view with detail of the position of pineal end–organ (arrow), (D) oral disc (ventral view), (E) labial teeth observed with scanning electron microscope.
On following pages: 147. Swayne's Hartebeest (Alcelaphus swaynel); 148. Kongoni (Alcelaphus coki); 149. Lichtenstein's Hartebeest (Alcelaphus lichtensteinii); 150. Red Hartebeest (Alcelaphus caama); 151. Herola (Beatragus hunter). in Bovidae
On following pages: 147. Swayne's Hartebeest (Alcelaphus swaynel); 148. Kongoni (Alcelaphus coki); 149. Lichtenstein's Hartebeest (Alcelaphus lichtensteinii); 150. Red Hartebeest (Alcelaphus caama); 151. Herola (Beatragus hunter).
On following pages: 259. Western White-bellied Duiker (Cephalophus leucogaster); 260. Uele White-bellied Duiker (Cephalophus arrhenii); 261. Red-flanked Duiker (Cephalophus rufilatus); 262. Natal Red Duiker (Cephalophus natalensis); 263. Harvey's Duiker (Cephalophus harveyi); 264. Black-fronted Duiker (Cephalophus nigrifrons); 265. ltombwe Duiker (Cephalophus hypoxanthus); 266. Rwenzori Duiker (Cephalophus rubidus); 267. Mount Elgon Duiker (Cephalophus fosteri); 268. Mount Kenya Duiker (Cephalophus hooki). in Bovidae
On following pages: 259. Western White-bellied Duiker (Cephalophus leucogaster); 260. Uele White-bellied Duiker (Cephalophus arrhenii); 261. Red-flanked Duiker (Cephalophus rufilatus); 262. Natal Red Duiker (Cephalophus natalensis); 263. Harvey's Duiker (Cephalophus harveyi); 264. Black-fronted Duiker (Cephalophus nigrifrons); 265. ltombwe Duiker (Cephalophus hypoxanthus); 266. Rwenzori Duiker (Cephalophus rubidus); 267. Mount Elgon Duiker (Cephalophus fosteri); 268. Mount Kenya Duiker (Cephalophus hooki).
On following pages: 224. Burmese Red Serow (Capricornis rubidus); 225. Japanese Serow (Capricornis crispus); 226. Formosan Serow (Capricornis swinhoei); 227. Muskox (Ovibos moschatus). in Bovidae
On following pages: 224. Burmese Red Serow (Capricornis rubidus); 225. Japanese Serow (Capricornis crispus); 226. Formosan Serow (Capricornis swinhoei); 227. Muskox (Ovibos moschatus).
On following pages: 211. Alpine Chamois (Rupicapra rupicapra); 212. Carpathian Chamois (Rupicapra carpatica); 213 Gray Goral (Nemorhaedus bedford); 216. Chinese Goral (Nemorhaedus griseus); 217. Burmese Goral (Nemorhaedus. Asia Minor Chamois (Rupicapra asiatica); 214. Himalayan Brown Goral (Nemorhaedus goral); 215. Himalayan evansi); 218. Long-tailed Goral (Nemorhaedus caudatus); 219. Red Goral (Nemorhaedus baileyi). in Bovidae
On following pages: 211. Alpine Chamois (Rupicapra rupicapra); 212. Carpathian Chamois (Rupicapra carpatica); 213 Gray Goral (Nemorhaedus bedford); 216. Chinese Goral (Nemorhaedus griseus); 217. Burmese Goral (Nemorhaedus. Asia Minor Chamois (Rupicapra asiatica); 214. Himalayan Brown Goral (Nemorhaedus goral); 215. Himalayan evansi); 218. Long-tailed Goral (Nemorhaedus caudatus); 219. Red Goral (Nemorhaedus baileyi).
Home range sizes of red deer in contrasting landscapes and implications for management
<p><span>Knowledge about deer spatial use is essential for damage mitigation and management coordination. Here we assess annual and seasonal home range sizes for </span><span>red deer in Sweden, based on data from GPS-marked deer in two regions with different management systems and contrasting landscapes</span><span>. We compare our findings with reviewed data on European red deer (<em>Cervus elaphus</em>) home range sizes in Europe. We found that female annual</span><span> home ranges (95% kernels) were 2.7 times larger in a mixed agricultural-forest landscape compared to a forest-dominated landscape. Core areas (50% kernels) were approximately 1/5 of the full annual home ranges (90% kernels) regardless of region. Home range size in the forest landscape showed little inter-seasonal variation whereas in the agricultural-forest landscape, home ranges were significantly larger during calving, hunt, and winter-spring compared to summer and rut. In the forest landscape, females had home range sizes that enables single red deer management areas to manage their own females. Whereas, within the agricultural-forest landscape, female spatial use cover several license units. Here, the coordinated license system is needed to reach trade-offs between goals of conservation, game management, and damage mitigation. Males had in general larger home ranges than females. The majority of the males made a seasonal migration to and from the rutting areas. The license system in the agricultural-forest landscape is large enough to manage migrating males, but in the forest landscape a coordination of several deer management areas is needed in order to encompass male spatial use. </span></p>
On following pages 48 Mexıcen Red Brocket (Mazama remomal 49 Small Red Brocket (Mazamo boroml, 50 Lesser Brocket (Mazarno nenas), 51 Lıttle Rod Brocket (Manmo mfinal 52 Mende Bracket (Mazama bncennl in Cervidae
On following pages 48 Mexıcen Red Brocket (Mazama remomal 49 Small Red Brocket (Mazamo boroml, 50 Lesser Brocket (Mazarno nenas), 51 Lıttle Rod Brocket (Manmo mfinal 52 Mende Bracket (Mazama bncennl
On following pages 4 Grant Muntpac (Munrıacus vuquengensıs) 5 Fea s Munt|ac lMunuacus feaol 6 Gongshan munqakl 9 Leaf Munuac (Mumıecus putaoensıs) 10 Puhoat Muntpac (Muntracus puhoatanslsl Annarnıte Munqac Mumpc (Munrıecus gongshanenslsl 7 Black Munqac (Mumracus cnmfronsl 8 Red Muntpc (Munnacus (Munrıocııs rruongsonensıs) 12 Roosevelt: Munqac (Munuacus rooseveltorurrrl in Cervidae
On following pages 4 Grant Muntpac (Munrıacus vuquengensıs) 5 Fea s Munt|ac lMunuacus feaol 6 Gongshan munqakl 9 Leaf Munuac (Mumıecus putaoensıs) 10 Puhoat Muntpac (Muntracus puhoatanslsl Annarnıte Munqac Mumpc (Munrıecus gongshanenslsl 7 Black Munqac (Mumracus cnmfronsl 8 Red Muntpc (Munnacus (Munrıocııs rruongsonensıs) 12 Roosevelt: Munqac (Munuacus rooseveltorurrrl
Data from: Variation in red fox Vulpes vulpes diet in five continents
<p><span>Understanding variation in the diet of widely distributed species can help us to predict how they respond to future environmental and anthropogenic changes.</span></p> <p><span>We studied the diet of the red fox <em>Vulpes vulpes</em>, one of the world's most widely distributed carnivores. We compiled dietary data from 217 studies at 276 locations in five continents to assess how fox diet composition varied according to geographic location, climate, anthropogenic impact and sampling method.</span></p> <p><span>The diet of foxes showed substantial variation throughout the species' range, but with a general trend for small mammals and invertebrates to be the most frequently occurring dietary items.</span></p> <p><span>The incidence of small and large mammals and birds in fox diets was greater away from the equator. The incidence of invertebrates and fruits increased with mean elevation, while the occurrence of medium-sized mammals and birds decreased. </span></p> <p><span>Fox diet differed according to climatic and anthropogenic variables. Diet richness decreased with increasing temperature and precipitation. The incidence of small and large mammals decreased with increasing temperature. The incidence of birds and invertebrates decreased with increasing mean annual precipitation. Higher Human Footprint Index was associated with lower incidence of large mammals and higher incidence of birds and fruit in fox diet. </span></p> <p><span>Sampling method influenced fox diet estimation: estimated percentage of small and medium-sized mammals and fruit was lower in studies based on stomach contents, while large mammals were more likely to be recorded in studies of stomach contents than in studies of scats.</span></p> <p><span>Our study confirms the flexible and opportunistic dietary behaviour of foxes at the global scale. This behavioural trait allows them to thrive in a range of climatic conditions, and in areas with different degrees of human-induced habitat change. This knowledge can help place the results of local-scale fox diet studies into a broader context and to predict how foxes will respond to future environmental changes.</span></p> <div></div>
Hybridization and low genetic diversity in the endangered Alabama Red-Bellied Turtle (Pseudemys alabamensis)
<p><em><span>Pseudemys</span></em><span> <em>alabamensis</em> is one of the most endangered freshwater turtle species in the United States due to its restricted geographic distribution in coastal Alabama and Mississippi. Populations of <em>P. alabamensis </em>are geographically isolated from one another by land and salt water, which could act as barriers to gene flow. It is currently unknown how differentiated these isolated populations are from one another and whether they have experienced reductions in population size. Previous work found morphological differences between Alabama and Mississippi populations, suggesting that they may be evolutionarily distinct. Other <em>Pseudemys</em> turtles such as <em>P. concinna</em> and <em>P. floridana</em> occur naturally within the same geographic area as <em>P. alabamensis</em> and are known to hybridize with each other. These more abundant species could threaten the unique genetic identity of <em>P. alabamensis</em> through introgression. In order to evaluate the endangered status of <em>P. alabamensis</em> and the level of hybridization with other species, we used mitochondrial and nuclear microsatellite markers to assess genetic variation within and among populations of <em>P. alabamensis</em> throughout its range and estimate admixture with co-occurring <em>Pseudemys</em> species. In <em>P. alabamensis</em>, we found no variation in mitochondrial DNA and observed an excess of homozygosity in the microsatellite data. Our results indicate evidence of genetic differentiation between Alabama and Mississippi populations of <em>P. alabamensis</em>, and low estimated breeding sizes and inbreeding for two populations (Fowl River, Alabama and Biloxi, Mississippi). Our results also support admixture of <em>P. alabamensis</em> with<em> P. concinna/P. floridana</em>. Based on our results, <em>P. alabamensis</em> is highly endangered throughout its range and threatened by both low population sizes and hybridization. In order to improve the species' chances of survival, focus should be placed on habitat preservation, maintenance of genetic diversity within both Mississippi and Alabama populations, and regular population monitoring activities such as nest surveillance and estimates of recruitment. </span></p>
FIGURE 9 in Three new species of the shrimp genus Salmoneus Holthuis, 1955 from the Red Sea, with a new record of S. poupini Anker, 2010 (Malacostraca: Decapoda Alpheidae)
FIGURE 9. Salmoneus kausti sp. nov., holotype, non-ovigerous specimen (cl 3.4 mm) from Thuwal, Saudi Arabia (FLMNH UF 36066), shrimp in life, lateral. Photograph by the author.
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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.