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opencc-by-4.0Feb 2014View details →

Fig. 3 in Morphology of the first instar larva of obligatory traumatic myiasis agents (Diptera: Calliphoridae, Sarcophagidae)

Fig. 3 First instar of Cochliomyia hominivorax. a Abdominal segments 1–3, ventral view. b Abdominal segment 2, spines. c Anal division, ventral view. d Anal division, posterior spiracles. Abbreviations: a1–a3 abdominal segments 1–3, ao anal opening, ap anal pad, asb anterior spinose band, at anal tuft, lcw lateral creeping welt, psb posterior spinose band, st peristigmatic tufts

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Fig. 2 in Morphology of the first instar larva of obligatory traumatic myiasis agents (Diptera: Calliphoridae, Sarcophagidae)

Fig. 2 First instar of Cochliomyia hominivorax. a Anterior end of body, antero-lateral view. b Anterior end of body, ventral view. c Antennal complex. d Maxillary palpus. e Functional mouth opening. f Ventral organ. g Keilin's organ. h First thoracic segment, spines

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Fig. 1 in Morphology of the first instar larva of obligatory traumatic myiasis agents (Diptera: Calliphoridae, Sarcophagidae)

Fig. 1 First instar of Chrysomya bezziana. a Anterior end of body, lateral view. b Functional mouth opening. c Antennal complex. d Maxillary palpus. e Ventral organ. f Third abdominal segment, ventral view, anterior spines. g Anal division, posterior view. Abbreviations: abr antennal basal ring, ad antennal dome, an antennal complex, ao anal opening, ap anal pad, cir cirri, mh mouthhooks, mp maxillary palpus, ll labial lobe, lo labial organ, ns1 first additional sensillum coeloconicum, ns2 second additional sensillum coeloconicum, or oral ridges, p1–p7 papillae 1–7 sb1–sb3 sensilla basiconica, sc1–sc3 sensilla coeloconica, sp posterior spiracle, vo ventral organ

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Fig. 1 4. Terellia tussilaginis, larva. 1 in Description Of The Third Instar Larvae Of Terellia Tussilaginis And T. Gynaecochroma (Diptera, Tephritidae)

Fig. 1 4. Terellia tussilaginis, larva. 1 — habitus, left; 2 — habitus5, ventral; 3 — larvae in seed; 4 — anterior spiracle; 5 — gnathocephalon, anterior view: ant — antenna; lso — lateral sensory organ; mh — mouthhook; max org — maxillary sensory organ; or rdg — oral ridges; s — sensillum.

Fig. 9 in Description of the female, nymph and larva and mitochondrial genome, and redescription of the male of Ixodes barkeri Barker, 2019 (Acari: Ixodidae), from the short-beaked echidna, Tachyglossus aculeatus, with a consideration of the most suitable subgenus for this tick

Fig. 9 Mitochondrial genomes of Ixodes (Endopalpiger) australiensis, I. (Endo.) barkeri, I. (Endo.) woyliei and I. (Exopalpiger) fecialis. Protein-coding genes are shown in green, tRNAs are in yellow, rRNAs are in red, and the two control regions are in blue. Protein-coding genes are labelled by their four-character abbreviations, tRNAs are labelled by their one-letter amino acid abbreviations, and the two control regions are labelled as CR1 and CR2. Mitochondrial genome size variation is indicated in parentheses. The arrangement of genes in these four species is identical except that the main cluster of tRNA genes has the arrangement ARNSEF in the three species of Endopalpiger [I. (Endo.) australiensis, I. (End.) barkeri and I. (End.) woyliei], whereas in the one species of Exopalpiger [I. (Exo.) fecialis] the arrangement is ARNESF. The arrangement in I. (Exo.) fecialis is the first known arrangement in an Ixodidae tick that is different from ARNSEF.Thus, ARNESF might be a synapomorphy for the subgenus Exopalpiger

Fig. 7 Ixodes barkeri Barker, 2019 in Description of the female, nymph and larva and mitochondrial genome, and redescription of the male of Ixodes barkeri Barker, 2019 (Acari: Ixodidae), from the short-beaked echidna, Tachyglossus aculeatus, with a consideration of the most suitable subgenus for this tick

Fig. 7 Ixodes barkeri Barker, 2019, scanning electron micrographs of larva. A Scutum. B Gnathosoma, dorsal view. C Gnathosoma, ventral view. D Gnathosoma, anteroventral view. E Coxae. Scale bars: A, E 0.1 mm; B–D, 0.05 mm

Fig. 1 in Description of the female, nymph and larva and mitochondrial genome, and redescription of the male of Ixodes barkeri Barker, 2019 (Acari: Ixodidae), from the short-beaked echidna, Tachyglossus aculeatus, with a consideration of the most suitable subgenus for this tick

Fig. 1 The four known localities in Australia, Queensland (Qld), of Ixodes barkeri Barker, 2019, are indicated by white-with-red dots

Fig. 10 in Description of the female, nymph and larva and mitochondrial genome, and redescription of the male of Ixodes barkeri Barker, 2019 (Acari: Ixodidae), from the short-beaked echidna, Tachyglossus aculeatus, with a consideration of the most suitable subgenus for this tick

Fig. 10 Maximum likelihood (ML) phylogenetic tree from entire mt genomes (14,935 bps). The sequence alignment was put though Gblocks to remove regions with alignment gaps.Tip labels indicate NCBI accession numbers and (Barker & Barker Collection reference nos.). Numbers above branches show maximum likelihood bootstrap support, whereas numbers below branches show the Bayesian posterior probability support. Ixodes pavlovskyi Pomerantzev, 1946, one of the species "Other Ixodes" (sensu Barker & Murrell, 2004), for which an entire mitochondrial (mt) genome was available in GenBank, was set as the outgroup. The scale bar indicates 0.06 nucleotide substitutions per nucleotide site for the 14,935 nucleotide sites in our alignment of theses entire mt genomes. So, for example, there were about 896 nucleotide substitutions along the branch that leads to I. (Ceratixodes) uriae plus I. (Sternalixodes) holocyclus plus I. (Exopalpiger) fecialis, which is marked with an asterisk [i.e. 0.06 nucleotide substitutions per nucleotide site × 14,935 nucleotide sites (bps) = 1896 nucleotide substitutions]. Ticks in bold were sequenced in the present study

Fig. 4 Ixodes barkeri Barker, 2019 in Description of the female, nymph and larva and mitochondrial genome, and redescription of the male of Ixodes barkeri Barker, 2019 (Acari: Ixodidae), from the short-beaked echidna, Tachyglossus aculeatus, with a consideration of the most suitable subgenus for this tick

Fig. 4 Ixodes barkeri Barker, 2019, scanning electron micrographs of female. A Idiosoma, dorsal view. B Scutum, dorsal view. C Scutum, dorsolateral view. D Idiosoma showing scutum and alloscutum with punctations and setae, dorsal centrolateral portion. E Idiosoma, ventral view. Scale bars: A, E 0.5 mm; B, C 0.2 mm; D 0.1 mm

Fig. 6 Ixodes barkeri Barker, 2019 in Description of the female, nymph and larva and mitochondrial genome, and redescription of the male of Ixodes barkeri Barker, 2019 (Acari: Ixodidae), from the short-beaked echidna, Tachyglossus aculeatus, with a consideration of the most suitable subgenus for this tick

Fig. 6 Ixodes barkeri Barker, 2019, scanning electron micrographs of nymph. A Scutum. B Spiracular plate (arrows show orientation of spiracular plate: a, anterior; d, dorsal). C Gnathosoma, dorsal view. D Gnathosoma, ventral view. E Gnathosoma, anteroventral view. F Coxae. Scale bars: A, C–F, 0.1 mm; B, 0.05 mm

Fig. 5 Ixodes barkeri Barker, 2019 in Description of the female, nymph and larva and mitochondrial genome, and redescription of the male of Ixodes barkeri Barker, 2019 (Acari: Ixodidae), from the short-beaked echidna, Tachyglossus aculeatus, with a consideration of the most suitable subgenus for this tick

Fig. 5 Ixodes barkeri Barker, 2019, scanning electron micrographs of female. A Spiracular plate (arrows show orientation of spiracular plate: a, anterior; d, dorsal). B Gnathosoma, dorsal view. C Gnathosoma, ventral view (I, palpal article 1; II, palpal article 2; ss the strongly salient part of palpal article 1). D Gnathosoma, anteroventral view. E Coxae. F Trochanter I, dorsal view. Scale bars: A, F, 0.1 mm; B–E, 0.2 mm

Fig. 8 Ixodes barkeri Barker, 2019 in Description of the female, nymph and larva and mitochondrial genome, and redescription of the male of Ixodes barkeri Barker, 2019 (Acari: Ixodidae), from the short-beaked echidna, Tachyglossus aculeatus, with a consideration of the most suitable subgenus for this tick

Fig. 8 Ixodes barkeri Barker, 2019, light microscopy image of female (Barker & Barker Collection reference #B5321), male (# B4994), nymph (#B5321) and larva (# B5321). Horizontal broken scale bars: 1 mm; vertical scale bars also in mm

Fig. 2 Ixodes barkeri Barker, 2019 in Description of the female, nymph and larva and mitochondrial genome, and redescription of the male of Ixodes barkeri Barker, 2019 (Acari: Ixodidae), from the short-beaked echidna, Tachyglossus aculeatus, with a consideration of the most suitable subgenus for this tick

Fig. 2 Ixodes barkeri Barker, 2019, scanning electron micrographs of idiosoma of male. A Dorsal view; B dorsolateral view; C ventral view. Scale bars: 0.5 mm

Fig. 3 Ixodes barkeri Barker, 2019 in Description of the female, nymph and larva and mitochondrial genome, and redescription of the male of Ixodes barkeri Barker, 2019 (Acari: Ixodidae), from the short-beaked echidna, Tachyglossus aculeatus, with a consideration of the most suitable subgenus for this tick

Fig. 3 Ixodes barkeri Barker, 2019, scanning electron micrographs of male. A Spiracular plate (arrows show orientation of spiracular plate: a, anterior; d, dorsal). B Gnathosoma, dorsal view. C Gnathosoma, ventral view. D Gnathosoma, anteroventral view. E Coxae. F Trochanter I, dorsal view. Scale bars: A–D, F 0.1 mm; E, 0.2 mm

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Growth, nutrient uptake, blood metabolites and bone properties in broilers consuming feed with mineral-enriched whole black soldier fly larvae

<p>Recycling critical minerals like phosphorus is essential for future circular agriculture. This study explored adding mineral-enriched black soldier fly larvae (BSFL), grown on substrates with sewage sludge recyclates (SSR), to broiler feed to reintroduce minerals from restricted waste streams into the nutrient cycle. Results showed that including 15% mineral-enriched BSFL in broiler diets for 42 days had no adverse effects on growth, nutrient intake, or bone condition, and maintained acceptable levels of heavy metals.</p>

Fig. 7 in Preliminary larvicidal effect of seed oil from Cleome arabica L. on fifth instar larvae of Schistocerca gregaria (Orthoptera: Acrididae)

Fig. 7. Variation of the weight compared to the initial weight of L5 larvae of control S. gregaria and treated with C. arabica seed oil.

Fig. 2 in Preliminary larvicidal effect of seed oil from Cleome arabica L. on fifth instar larvae of Schistocerca gregaria (Orthoptera: Acrididae)

Fig. 2. Variation over time of mortality rate observed in control L5 larvae treated with C. arabica seed oil.

Fig. 1 C-D in Preliminary larvicidal effect of seed oil from Cleome arabica L. on fifth instar larvae of Schistocerca gregaria (Orthoptera: Acrididae)

Fig. 1 C-D. Cleome arabica at fruiting stage (Oued Metlili, Region of Ghardaïa-Northern Algerian Sahara, April 2020). C) Fruiting brunches. D) Fruiting close-up.

Fig. 1 A-B in Preliminary larvicidal effect of seed oil from Cleome arabica L. on fifth instar larvae of Schistocerca gregaria (Orthoptera: Acrididae)

Fig. 1 A-B. Cleome arabica at fruiting stage (Oued Metlili, Region of Ghardaïa-Northern Algerian Sahara, April 2020). A) Whole plant. B) Seed.

Fig. 5 A-B in Preliminary larvicidal effect of seed oil from Cleome arabica L. on fifth instar larvae of Schistocerca gregaria (Orthoptera: Acrididae)

Fig. 5 A-B. Morphological abnormalities observed in L5 muer larvae of S. gregaria treated with C. arabica seed oil.