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78 results for “host-specificity”
FIGURE 4 in An updated concept and revised composition for Hamacreadium Linton, 1910 (Opecoelidae: Plagioporinae) clarifies a previously obscured pattern of host-specificity among species
FIGURE 4. Reproduction with modification of type illustrations for (a) Hamacreadium longivesiculum (Yamaguti, 1952) n. comb. and (b) Hamacreadium lutiani (Shen, 1990) n. comb. Abbreviations: C, caecum; Cs, cirrus-sac; Ep, excretory pore; Ev, excretory vesicle; Gp, genital pore; O, ovary; Oes, oesophagus; Os, oral sucker; P, pharynx; T, testis; U, uterus; Vf, vitelline follicles; Vs, ventral sucker. Scale: 1000 µm, 1000 µm.
FIGURE 3 in An updated concept and revised composition for Hamacreadium Linton, 1910 (Opecoelidae: Plagioporinae) clarifies a previously obscured pattern of host-specificity among species
FIGURE 3. Reproduction with modification of type illustrations for (a) Hamacreadium interruptum Nagaty, 1941 and (b) Hamacreadium lethrini Yamaguti, 1934. Abbreviations: C, caecum; Cs, cirrus-sac; Ep, excretory pore; Ev, excretory vesicle; Gp, genital pore; O, ovary; Oes, oesophagus; Os, oral sucker; P, pharynx; T, testis; U, uterus; Vf, vitelline follicles; Vs, ventral sucker. Scale: 1000 µm, 1000 µm.
FIGURE 2 in An updated concept and revised composition for Hamacreadium Linton, 1910 (Opecoelidae: Plagioporinae) clarifies a previously obscured pattern of host-specificity among species
FIGURE 2. Reproduction with modification of type illustrations for (a) Hamacreadium cribbi Bray & Justine, 2016 and (b) Hamacreadium hainanense Shen, 1990. Abbreviations: C, caecum; Cs, cirrus-sac; Ep, excretory pore; Ev, excretory vesicle; Gp, genital pore; O, ovary; Oes, oesophagus; Os, oral sucker; P, pharynx; T, testis; U, uterus; Vf, vitelline follicles; Vs, ventral sucker. Scale: 1000 µm, 1000 µm.
FIGURES 19, 20 in Tapeworms (Cestoda: Proteocephalidae) of Australian reptiles: hidden diversity of strictly host-specific parasites
FIGURES 19, 20. Australophiotaenia striata (Johnston, 1914) n. comb., syntype from Townsville, Queensland, Australia [Coll. No. AHC 28406.] 19. Mature proglottid, ventral view. 20. Pregravid proglottid, dorsal view.
FIGURE 18 in Tapeworms (Cestoda: Proteocephalidae) of Australian reptiles: hidden diversity of strictly host-specific parasites
FIGURE 18. Australophiotaenia striata (Johnston, 1914) n. comb., syntype from Townsville, Queensland, Australia [Coll. No. AHC 28406.], scolex.
FIGURES 5–10 in Tapeworms (Cestoda: Proteocephalidae) of Australian reptiles: hidden diversity of strictly host-specific parasites
FIGURES 5–10. Australophiotaenia longmani (Johnston, 1916) n. comb., 5, 6. Syntype from Yuleba, Queensland, Australia [Coll. No. QMNH G16/468] 5. Scolex. 6. Mature proglottid, dorsal. 7–10. Voucher MHNG-PLAT-36551 (field No. AUS 013). 7. Pregravid proglottid, dorsal. 8. Cross section at level of uterus with eggs in capsules. 9, 10. Eggs in capsules.
FIGURES 11–17 in Tapeworms (Cestoda: Proteocephalidae) of Australian reptiles: hidden diversity of strictly host-specific parasites
FIGURES 11–17. Australophiotaenia mjobergi (Nybelin, 1917) n. comb., syntype from Police Camp, Northern Territory, Australia [Coll. No. SMNH 3272]. 11. Scolex. 12. Pregravid proglottid, ventral. 13. Terminal genitalia. 14–16. Cross sections at level of ovary, testes and uterus, respectively. 17. Eggs in capsules.
FIGURES 1–4 in Tapeworms (Cestoda: Proteocephalidae) of Australian reptiles: hidden diversity of strictly host-specific parasites
FIGURES 1–4. Australophiotaenia amphiboluri (Nybelin, 1917) n. comb., syntype from Mount Tamborine, Queensland, Australia [Coll. No. SMNH 3279]. 1. Mature proglottid, ventral. 2. Cross sections at level of testes. 3. Cross sections at level of uterus with forming eggs. 4. Cross sections at level of eggs in capsules. Abbreviations (Figs. 1–20): cc—chromophilic cells; ci—cirrus; cs—cirrus-sac; doc—dorsal osmoregulatory canal; ec—egg capsule; em—bi-layered embryophore; ga—genital atrium; gc—gland cells; lm—internal longitudinal musculature; gm—Mehlis' glands; mf—muscle fibres; mi—microtriches; oe—outer envelope; od—oviduct; on—oncosphere; ov—ovary; st—subtegumental muscle fibres; su—subtegumental cells; te—testes; tg—tegument; ud—uterine diverticula; us—uterine stem; va—vas deferens; vc—vaginal canal; vd—vitelloduct; vf—vitelline follicles; voc—ventral osmoregulatory canal; vs—vaginal sphincter.
Data from: Genetic architecture of resistance in Daphnia hosts against two species of host-specific parasites
Understanding the genetic architecture of host resistance is key for understanding the evolution of host–parasite interactions. Evolutionary models often assume simple genetics based on few loci and strong epistasis. It is unknown, however, whether these assumptions apply to natural populations. Using a quantitative trait loci (QTL) approach, we explore the genetic architecture of resistance in the crustacean Daphnia magna to two of its natural parasites: the horizontally transmitted bacterium Pasteuria ramosa and the horizontally and vertically transmitted microsporidium Hamiltosporidium tvaerminnensis. These two systems have become models for studies on the evolution of host–parasite interactions. In the QTL panel used here, Daphnia's resistance to P. ramosa is controlled by a single major QTL (which explains 50% of the observed variation). Resistance to H. tvaerminnensis horizontal infections shows a signature of a quantitative trait based in multiple loci with weak epistatic interactions (together explaining 38% variation). Resistance to H. tvaerminnensis vertical infections, however, shows only one QTL (explaining 13.5% variance) that colocalizes with one of the QTLs for horizontal infections. QTLs for resistance to Pasteuria and Hamiltosporidium do not colocalize. We conclude that the genetics of resistance in D. magna are drastically different for these two parasites. Furthermore, we infer that based on these and earlier results, the mechanisms of coevolution differ strongly for the two host–parasite systems. Only the Pasteuria–Daphnia system is expected to follow the negative frequency-dependent selection (Red Queen) model. How coevolution works in the Hamiltosporidium–Daphnia system remains unclear.
Data from: Transcriptomics of host-specific interactions in natural populations of the parasitic plant purple witchweed (Striga hermonthica)
Host-specific interactions can maintain genetic and phenotypic diversity in parasites that attack multiple host species. Host diversity, in turn, may promote parasite diversity by selection for genetic divergence or plastic responses to host type. The parasitic weed purple witchweed [Striga hermonthica (Delile) Benth.] causes devastating crop losses in sub-Saharan Africa and is capable of infesting a wide range of grass hosts. Despite some evidence for host adaptation and host-by-Striga genotype interactions, little is known about intraspecific Striga genomic diversity. Here we present a study of transcriptomic diversity in populations of S. hermonthica growing on different hosts (maize [Zea mays L.] vs. grain sorghum [Sorghum bicolor (L.) Moench]). We examined gene expression variation and differences in allelic frequency in expressed genes of aboveground tissues from populations in western Nigeria parasitizing each host. Despite low levels of host-based genome-wide differentiation, we identified a set of parasite transcripts specifically associated with each host. Parasite genes in several different functional categories implicated as important in host–parasite interactions differed in expression level and allele on different hosts, including genes involved in nutrient transport, defense and pathogenesis, and plant hormone response. Overall, we provide a set of candidate transcripts that demonstrate host-specific interactions in vegetative tissues of the emerged parasite S. hermonthica. Our study shows how signals of host-specific processes can be detected aboveground, expanding the focus of host–parasite interactions beyond the haustorial connection.
Alpha-diversity, Beta-diversity and host-specificity of wood-boring longhorn beetle (Cerambycidea) in Asian tropical and subtropical forests
<p><span>A long-debated question in ecology is whether the hyper-diversity of tropical plant-feeding insects is a direct consequence of high tropical plant diversity and/or should be attributed to increases in host plant specialization. To address this debate, we used the longhorn beetle as a study system because their larval stages feed on the xylems of trees and lianas. We hypothesized that longhorn beetles show higher host-specificity in tropical forests than in other forests; alternatively, the high longhorn beetle diversity in the tropics may simply be owing to more diverse host plants. We therefore designed an investigation in tropical and subtropical forests to test these hypotheses. We adapted several analyses (i.e., non-metric multidimensional scaling analysis, alpha-diversity, beta-dissimilarity indices comparisons, and variation partitioning based on redundancy analysis) to compare the species diversity of plants and longhorn beetles in different forests. Our results show that both the plant and beetle species in the tropical and subtropical areas were well-stratified (non-metric multidimensional scaling analysis). The beetle alpha-diversity in the tropical forests was significantly higher than that in the subtropical forests, but the plant alpha-diversity in the two types of forests were not significantly different. The beta-dissimilarity comparison showed that the plant species exerted a significant influence on beetle compositional assemblage in the tropical forests, but not in the subtropical forests. Finally, the variation partitioning results showed that both plant species and plant phylogenetic beta-diversity possessed significant explanatory power for beetle assemblage composition in the tropical forests, but not in the subtropical forests. We conclude that wood-boring longhorn beetles show higher host-specificity in tropical forests than in subtropical forests, and the high diversity of wood-boring longhorn beetles in tropical forests might be explained to a large extent by their more finely partitioned diet-breadth.</span></p>
Figure 11 in Developmental stages and growth of Pseudocaligus fugu Yamaguti, 1936 (Copepoda: Siphonostomatoida: Caligidae) host-specific to Puffer
Figure 11. Relationships between length of egg string and number of eggs per string (A) and between numbers of eggs per right and left egg strings (B) in females of Pseudocaligus fugu obtained at a culture area off Kosasa Town, Sasebo City on 29 May 2008.
Figure 9 in Developmental stages and growth of Pseudocaligus fugu Yamaguti, 1936 (Copepoda: Siphonostomatoida: Caligidae) host-specific to Puffer
Figure 9. Pseudocaligus fugu, young adult female. (A) Habitus, dorsal view; (B) frontal filament; (C) genital compound somite, ventral view, note sinuate lateral margins of somite and attachment of paired spermatophores; (D) spermatophore; (E) right leg 4. Scales in mm.
Figure 8 in Developmental stages and growth of Pseudocaligus fugu Yamaguti, 1936 (Copepoda: Siphonostomatoida: Caligidae) host-specific to Puffer
Figure 8. Pseudocaligus fugu, fourth chalimus stage. Female (A–N), male (O–R). (A, O) Habitus, dorsal view; (B) frontal filament; (C) left caudal ramus, ventral view; (D) antennule, antenna and postantennary process in situ; (E) distal segment of antennule; (F) maxillule; (G) maxilla; (H) maxilliped; (I) right leg 1, anterior surface; (J) endopod of right leg 1, anterior surface; (K) leg 2, anterior surface; (L) leg 3, anterior surface; (M, Q) left leg 4; (N, R) leg 5; (P) antenna and postantennary process. Scales in mm.
Figure 7 in Developmental stages and growth of Pseudocaligus fugu Yamaguti, 1936 (Copepoda: Siphonostomatoida: Caligidae) host-specific to Puffer
Figure 7. Pseudocaligus fugu, third chalimus stage. Male (A–P), female (Q–S). (A, Q) Habitus, dorsal view; (B) frontal filament; (C) right caudal ramus, ventral view; (D) antennule; (E) antenna; (F) postantennary process; (G) mandible; (H) maxillule; (I) maxilla; (J) maxilliped; (K) leg 1, anterior surface; (L) endopod of right leg 1, anterior surface; (M) leg 2, anterior surface; (N) leg 3, anterior surface; (O) leg 4; (P, S) leg 5; (R) antenna and postantennary process. Scales in mm.
Figure 6 in Developmental stages and growth of Pseudocaligus fugu Yamaguti, 1936 (Copepoda: Siphonostomatoida: Caligidae) host-specific to Puffer
Figure 6. Pseudocaligus fugu, second chalimus stage. (A) Habitus, dorsal view; (B) frontal filament; (C) left caudal ramus, dorsal surface; (D) antennule, antenna and postantennary process in situ; (E) mandible; (F) maxillule; (G) maxilla; (H) maxilliped; (I) leg 1, anterior surface; (J) leg 2, anterior surface; (K).leg 3, anterior surface; (L) leg 4. Scales in mm.
Figure 5 in Developmental stages and growth of Pseudocaligus fugu Yamaguti, 1936 (Copepoda: Siphonostomatoida: Caligidae) host-specific to Puffer
Figure 5. Pseudocaligus fugu, first chalimus stage. (A) Habitus, dorsal view; (B) right caudal ramus, dorsal surface; (C) antennule; (D) antenna; (E) mandible; (F) maxillule; (G) maxilla; (H) maxilliped; (I) leg 1, anterior surface; (J) endopod of right leg 1, anterior surface; (K) leg 2, anterior surface; (L) endopod of right leg 2, anterior surface; (M) leg 3, anterior surface. Scales in mm.
Figure 4 in Developmental stages and growth of Pseudocaligus fugu Yamaguti, 1936 (Copepoda: Siphonostomatoida: Caligidae) host-specific to Puffer
Figure 4. Pseudocaligus fugu, copepodid stage. (A) Habitus, dorsal view; (B) rostrum; (C) left caudal ramus, dorsal surface; (D) antennule; (E) antenna; (F) mandible; (G) maxillule; (H) maxilla; (I) maxilliped; (J) postoral process; (K) leg 1, anterior surface; (L) leg 2, anterior surface; (M) leg 3, dorsal view. Scales in mm.
Figure 3 in Developmental stages and growth of Pseudocaligus fugu Yamaguti, 1936 (Copepoda: Siphonostomatoida: Caligidae) host-specific to Puffer
Figure 3. Pseudocaligus fugu, naupliar stages. First (A–D) and second (E–G) stages. (A, E) Habitus, ventral view; (B, F) antennule; (C, G) antenna; (D) mandible. Scales in mm.
Figure 2 in Developmental stages and growth of Pseudocaligus fugu Yamaguti, 1936 (Copepoda: Siphonostomatoida: Caligidae) host-specific to Puffer
Figure 2. Body lengths of developmental stages of Pseudocaligus fugu collected from Nagasaki Prefecture.
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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.