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7 results for “Blarina”
Blarina brevicauda populations in three different habitats in east-central Illinois, 1972 to 1997.
The population demography of the northern short-tailed shrew, Blarina brevicauda, was monitored monthly from 1972-1997 in bluegrass, alfalfa, and tallgrass prairie habitats in east-central Illinois. Blarina brevicauda were incidentally collected as part of a 25-year prairie vole (Microtus ochrogaster) and meadow vole (M. pennsylvanicus) trapping study. The study sites were located in the University of Illinois Biological Research Area (Phillips Tract) and Trelease Prairie. Animals were trapped with wooden multiple-capture live-traps. Over the span of 25 years, three 3-day trapping sessions monthly were conducted to cover the three habitats. All live shrews were toe-clipped at first capture for individual identification. Animals known to be present on study site, but not captured, were included in totals for that month. While trap mortality of shrews was high in this study (40%), the data obtained were sufficient for analysis of many aspects of the demography of the species.
Text-fig. 7. Enamel microstructure in the molars of (a) Allosorex stenodus (KOE-4196 / Z 28207 – OF 6512) and (b) Blarina brevicauda (KOE-1435). SEM photos showing transverse sections of the paralophid in a lower molar. The schmelzmuster of Allosorex is one-layered and formed exclusively from radial enamel (RE), whereas the schmelzmuster of Blarina is two-layered, with an inner enamel of laterally inclined radial enamel with distinct interrow sheets (REIS). The radial enamel of the outer layer is heavily pigmented and therefore poorly etched. The pigmentation does not affect the schmelzmuster. in Allosorex Stenodus Fejfar, 1966 (Eulipotyphla, Soricidae): Re-Description Of Type Material And Re-Interpretation Of Its Fossil Record
Text-fig. 7. Enamel microstructure in the molars of (a) Allosorex stenodus (KOE-4196 / Z 28207 – OF 6512) and (b) Blarina brevicauda (KOE-1435). SEM photos showing transverse sections of the paralophid in a lower molar. The schmelzmuster of Allosorex is one-layered and formed exclusively from radial enamel (RE), whereas the schmelzmuster of Blarina is two-layered, with an inner enamel of laterally inclined radial enamel with distinct interrow sheets (REIS). The radial enamel of the outer layer is heavily pigmented and therefore poorly etched. The pigmentation does not affect the schmelzmuster.
Text-fig. 8. The limited differentiation the schmelzmuster in Allosorex (Allosoricinae) indicates its phylogenetic position in Soricomorpha. The numbers to the right refer to the different types of schmelzmuster found in Soricidae: 1 – Soricella-schmelzmuster, 2 – Anourosorex- schmelzmuster, 3 – Crocidura-schmelzmuster, 4 – Notiosorex-schmelzmuster, 5 – Sorex-schmelzmuster, and 6 – Blarina-schmelzmuster (ex Koenigswald and Reumer 2020). in Allosorex Stenodus Fejfar, 1966 (Eulipotyphla, Soricidae): Re-Description Of Type Material And Re-Interpretation Of Its Fossil Record
Text-fig. 8. The limited differentiation the schmelzmuster in Allosorex (Allosoricinae) indicates its phylogenetic position in Soricomorpha. The numbers to the right refer to the different types of schmelzmuster found in Soricidae: 1 – Soricella-schmelzmuster, 2 – Anourosorex- schmelzmuster, 3 – Crocidura-schmelzmuster, 4 – Notiosorex-schmelzmuster, 5 – Sorex-schmelzmuster, and 6 – Blarina-schmelzmuster (ex Koenigswald and Reumer 2020).
Konza Prairie site, station Konza Prairie LTER watershed 001d, study of animal abundance of Blarina hylophaga in units of numberPerTransectLinePer4DayTrapSeason on a yearly timescale
The EcoTrends project was established in 2004 by Dr. Debra Peters (Jornada Basin LTER, USDA-ARS Jornada Experimental Range) and Dr. Ariel Lugo (Luquillo LTER, USDA-FS Luquillo Experimental Forest) to support the collection and analysis of long-term ecological datasets. The project is a large synthesis effort focused on improving the accessibility and use of long-term data. At present, there are ~50 state and federally funded research sites that are participating and contributing to the EcoTrends project, including all 26 Long-Term Ecological Research (LTER) sites and sites funded by the USDA Agriculture Research Service (ARS), USDA Forest Service, US Department of Energy, US Geological Survey (USGS) and numerous universities. Data from the EcoTrends project are available through an exploratory web portal (http://www.ecotrends.info). This web portal enables the continuation of data compilation and accessibility by users through an interactive web application. Ongoing data compilation is updated through both manual and automatic processing as part of the LTER Provenance Aware Synthesis Tracking Architecture (PASTA). The web portal is a collaboration between the Jornada LTER and the LTER Network Office. The following dataset from Konza Prairie (KNZ) contains animal abundance of Blarina hylophaga measurements in numberPerTransectLinePer4DayTrapSeason units and were aggregated to a yearly timescale.
Konza Prairie site, station Konza Prairie LTER watershed 004b, study of animal abundance of Blarina hylophaga in units of numberPerTransectLinePer4DayTrapSeason on a yearly timescale
The EcoTrends project was established in 2004 by Dr. Debra Peters (Jornada Basin LTER, USDA-ARS Jornada Experimental Range) and Dr. Ariel Lugo (Luquillo LTER, USDA-FS Luquillo Experimental Forest) to support the collection and analysis of long-term ecological datasets. The project is a large synthesis effort focused on improving the accessibility and use of long-term data. At present, there are ~50 state and federally funded research sites that are participating and contributing to the EcoTrends project, including all 26 Long-Term Ecological Research (LTER) sites and sites funded by the USDA Agriculture Research Service (ARS), USDA Forest Service, US Department of Energy, US Geological Survey (USGS) and numerous universities. Data from the EcoTrends project are available through an exploratory web portal (http://www.ecotrends.info). This web portal enables the continuation of data compilation and accessibility by users through an interactive web application. Ongoing data compilation is updated through both manual and automatic processing as part of the LTER Provenance Aware Synthesis Tracking Architecture (PASTA). The web portal is a collaboration between the Jornada LTER and the LTER Network Office. The following dataset from Konza Prairie (KNZ) contains animal abundance of Blarina hylophaga measurements in numberPerTransectLinePer4DayTrapSeason units and were aggregated to a yearly timescale.
On following pages: 428. Greater Congo Shrew (Congosorex polli); 429. Lesser Congo Shrew (Congosorex verheyeni 432. Sclater's Mouse Shrew (Myosorex sclateri); 433. Thin Mouse Shrew (Myosorex tenuis); 434. South African Mouse Mouse Shrew (Myosorex zinki); 437. Geata Mouse Shrew (Myosorex geata); 438. Kihaule's Mouse Shrew (Myosorex 441. Kabogo Mouse Shrew (Myosorex kabogoensis); 442. Babault's Mouse Shrew (Myosorex babaultii); 443. Montane Shrew (Myosorex jeje); 446. Eisentraut's Mouse Shrew (Myosorex eisentrauti); 447. Rumpi Mouse Shrew (Myosorex); 430. Meester's Mouse Shrew (Myosorex meester); 431. Dark-footed Mouse Shrew (Myosorex cafer); Shrew (Myosorex varius); 435. Long-tailed Mouse Shrew (Myosorex longicaudatus); 436. Kilimanjaro kihaulei); 439. Nyika Mouse Shrew (Myosorex gnoskei); 440. Schaller's Mouse Shrew (Myosorex schalleri); Mouse Shrew (Myosorex blarina); 444. Bururi Mouse Shrew (Myosorex bururiensis); 445. Kahuzi Mouse rumpii); 448. Oku Mouse Shrew (Myosorex okuensis). in Soricidae
On following pages: 428. Greater Congo Shrew (Congosorex polli); 429. Lesser Congo Shrew (Congosorex verheyeni 432. Sclater's Mouse Shrew (Myosorex sclateri); 433. Thin Mouse Shrew (Myosorex tenuis); 434. South African Mouse Mouse Shrew (Myosorex zinki); 437. Geata Mouse Shrew (Myosorex geata); 438. Kihaule's Mouse Shrew (Myosorex 441. Kabogo Mouse Shrew (Myosorex kabogoensis); 442. Babault's Mouse Shrew (Myosorex babaultii); 443. Montane Shrew (Myosorex jeje); 446. Eisentraut's Mouse Shrew (Myosorex eisentrauti); 447. Rumpi Mouse Shrew (Myosorex); 430. Meester's Mouse Shrew (Myosorex meester); 431. Dark-footed Mouse Shrew (Myosorex cafer); Shrew (Myosorex varius); 435. Long-tailed Mouse Shrew (Myosorex longicaudatus); 436. Kilimanjaro kihaulei); 439. Nyika Mouse Shrew (Myosorex gnoskei); 440. Schaller's Mouse Shrew (Myosorex schalleri); Mouse Shrew (Myosorex blarina); 444. Bururi Mouse Shrew (Myosorex bururiensis); 445. Kahuzi Mouse rumpii); 448. Oku Mouse Shrew (Myosorex okuensis).
On following pages: 138. Southern Short-tailed Shrew (Blarina carolinensis); 139. Everglades Short-tailed Shrew (Blarina peninsulae); 140. Sherman's Short-tailed Shrew (Blarina sherman); 141. Sichuan Short-tailed Shrew (Blarinella quadraticauda); 142. Burmese Short-tailed Shrew (Blarinella ward); 143. Indochinese Short-tailed Shrew (Blarinella griselda); 144. Chinese Mole Shrew (Anourosorex squamipes); 145. Taiwanese Mole Shrew (Anourosorex yamashinai); 146. Assam Mole Shrew (Anourosorex assamensis); 147. Giant Mole Shrew (Anourosorex schmidi); 148. Desert Gray Shrew (Notiosorex crawford); 149. Cockrum''s Gray Shrew (Notiosorex cockrumi); 150. Large-eared Gray Shrew (Notiosorex evotis); 151. Villa's Gray Shrew (Notiosorex villa); 152. Mexican Shrew (Megasorex gigas); 153. Taiwanese Brown-toothed Shrew (Episoriculus fumidus); 154. Arboreal Brown-toothed Shrew (Episoriculus macrurus); 155. Bailey's Brown-toothed Shrew (Episoriculus baileyi); 156. Long-tailed Brown-toothed Shrew (Episoriculus leucops); 157. Hodgson's Brown-toothed Shrew (Episoriculus caudatus); 158. Sichuan Brown-toothed Shrew (Episoriculus sacratus); 159. Hidden Brown-toothed Shrew (Episoriculus umbrinus); 160. Nepalese Brown-toothed Shrew (Episoriculus soluensis); 161. Himalayan Shrew (Soriculus nigrescens); 162. De Winton's Brown-toothed Shrew (Chodsigoa hypsibia); 163. Pygmy Brown-toothed Shrew (Chodsigoa parva); 164. Smith's Brown-toothed Shrew (Chodsigoa smithii); 165. Salenski's Brown-toothed Shrew (Chodsigoa salenski)); 166. Dusky Brown-toothed Shrew (Chodsigoa furva); 167. Lesser Taiwanese Brown-toothed Shrew (Chodsigoa sodalis); 168. Van Sung''s Brown-toothed Shrew (Chodsigoa caovansunga); 169. Hoffmann's Brown-toothed Shrew (Chodsigoa hoffmanni); 170. Lowe's Brown-toothed Shrew (Chodsigoa parca); 171. Bornean Water Shrew (Chimarrogale phaeura); 172. Sumatran Water Shrew (Chimarrogale sumatrana); 173. Malayan Water Shrew (Chimarrogale hantu); 174. Chinese Water Shrew (Chimarrogale styani); 175. Himalayan Water Shrew (Chimarrogale himalayica); 176. Leander's Water Shrew (Chimarrogale leander); 177. Japanese Water Shrew (Chimarrogale platycephala); 178. Elegant Water Shrew (Nectogale elegans); 179. Mediterranean Water Shrew (Neomys anomalus); 180. Transcaucasian Water Shrew (Neomys teres); 181. Eurasian Water Shrew (Neomys fodiens). in Soricidae
On following pages: 138. Southern Short-tailed Shrew (Blarina carolinensis); 139. Everglades Short-tailed Shrew (Blarina peninsulae); 140. Sherman's Short-tailed Shrew (Blarina sherman); 141. Sichuan Short-tailed Shrew (Blarinella quadraticauda); 142. Burmese Short-tailed Shrew (Blarinella ward); 143. Indochinese Short-tailed Shrew (Blarinella griselda); 144. Chinese Mole Shrew (Anourosorex squamipes); 145. Taiwanese Mole Shrew (Anourosorex yamashinai); 146. Assam Mole Shrew (Anourosorex assamensis); 147. Giant Mole Shrew (Anourosorex schmidi); 148. Desert Gray Shrew (Notiosorex crawford); 149. Cockrum''s Gray Shrew (Notiosorex cockrumi); 150. Large-eared Gray Shrew (Notiosorex evotis); 151. Villa's Gray Shrew (Notiosorex villa); 152. Mexican Shrew (Megasorex gigas); 153. Taiwanese Brown-toothed Shrew (Episoriculus fumidus); 154. Arboreal Brown-toothed Shrew (Episoriculus macrurus); 155. Bailey's Brown-toothed Shrew (Episoriculus baileyi); 156. Long-tailed Brown-toothed Shrew (Episoriculus leucops); 157. Hodgson's Brown-toothed Shrew (Episoriculus caudatus); 158. Sichuan Brown-toothed Shrew (Episoriculus sacratus); 159. Hidden Brown-toothed Shrew (Episoriculus umbrinus); 160. Nepalese Brown-toothed Shrew (Episoriculus soluensis); 161. Himalayan Shrew (Soriculus nigrescens); 162. De Winton's Brown-toothed Shrew (Chodsigoa hypsibia); 163. Pygmy Brown-toothed Shrew (Chodsigoa parva); 164. Smith's Brown-toothed Shrew (Chodsigoa smithii); 165. Salenski's Brown-toothed Shrew (Chodsigoa salenski)); 166. Dusky Brown-toothed Shrew (Chodsigoa furva); 167. Lesser Taiwanese Brown-toothed Shrew (Chodsigoa sodalis); 168. Van Sung''s Brown-toothed Shrew (Chodsigoa caovansunga); 169. Hoffmann's Brown-toothed Shrew (Chodsigoa hoffmanni); 170. Lowe's Brown-toothed Shrew (Chodsigoa parca); 171. Bornean Water Shrew (Chimarrogale phaeura); 172. Sumatran Water Shrew (Chimarrogale sumatrana); 173. Malayan Water Shrew (Chimarrogale hantu); 174. Chinese Water Shrew (Chimarrogale styani); 175. Himalayan Water Shrew (Chimarrogale himalayica); 176. Leander's Water Shrew (Chimarrogale leander); 177. Japanese Water Shrew (Chimarrogale platycephala); 178. Elegant Water Shrew (Nectogale elegans); 179. Mediterranean Water Shrew (Neomys anomalus); 180. Transcaucasian Water Shrew (Neomys teres); 181. Eurasian Water Shrew (Neomys fodiens).
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