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Fig. 1 in Environmental conditions predict helminth prevalence in red foxes in Western Australia

Fig. 1. Prevalence of helminths in red foxes (n=147) from sampling locations throughout southwest Western Australia. Numbers in parentheses indicate sample size at each location.

opencc-by-4.0Aug 2014View details →

Fig. 1. 2D in Proteomic profile of Ortleppascaris sp.: A helminth parasite of Rhinella marina in the Amazonian region

Fig. 1. 2D gel containing the somatic extract of Ortleppascaris sp. larvae. See Table 1 for details.

opencc-by-4.0Aug 2016View details →

Fig. 1 in Reduced helminth parasitism in the introduced bank vole (Myodes glareolus): More parasites lost than gained

Fig. 1. Frequency distribution of intestinal helminth species richness in wood mice and bank voles examined in 2011 and 2012.

Fig. 5 in Helminth community structure in two species of arctic-breeding waterfowl

Fig. 5. Predicted helminth infection intensity for combined cestodes identified in Pacific black brant (BLBR) and greater white-fronted geese (GWFG) collected in two locations (i.e., Arctic and Subarctic) in Alaska (2014). Circles represent predicted means and error bars denote 85% confidence intervals. Predictions are based off the most supported model from AICc selection and all interactions include lower-order effects (see title).

Fig. 4 in Helminth community structure in two species of arctic-breeding waterfowl

Fig. 4. Predicted prevalence (top) and helminth infection intensity (bottom) for the cestode Drepanidotaenia lanceolata enumerated in Pacific black brant (BLBR) and greater whitefronted geese (GWFG) collected in Arctic and Subarctic Alaska (2014). Circles represent predicted means and error bars denote 85% confidence intervals. Predictions are based off the most supported model from AICc selection and all interactions include lower-order effects (see titles).

Fig. 6 in Helminth community structure in two species of arctic-breeding waterfowl

Fig. 6. Predicted prevalence (top) and helminth infection intensity (bottom) for the cestode Tschertkovilepis setigera enumerated in Pacific black brant (BLBR) and greater whitefronted geese (GWFG) collected in Arctic and Subarctic Alaska (2014). Circles represent predicted means and error bars denote 85% confidence intervals. Predictions are based off the most supported model from AICc selection and all interactions include lower-order effects (see titles).

Fig. 2 in Helminth community structure in two species of arctic-breeding waterfowl

Fig. 2. Predicted helminth species richness (SR) for Pacific black brant (BLBR) and greater white-fronted geese (GWFG) collected from Subarctic and Arctic Alaska (2014). Circles represent predicted means and error bars denote 85% confidence intervals. Predictions are based on the most supported model from AICc selection and all interactions include lower-order effects (see title).

Fig. 3 in Helminth community structure in two species of arctic-breeding waterfowl

Fig. 3. Predicted prevalence (top) and helminth infection intensity (bottom) for the nematode Trichostrongylus tenuis enumerated in Pacific black brant (BLBR) and greater whitefronted geese (GWFG) collected in Arctic and Subarctic Alaska (2014). Circles represent predicted means and error bars denote 85% confidence intervals. Predictions are based off the most supported model from AICc selection and all interactions include lower-order effects (see titles).

Fig. 1 in Helminth community structure in two species of arctic-breeding waterfowl

Fig. 1. Study sites in Alaska where Pacific black brant and greater white-fronted geese were collected for helminth examination in 2014; A) Yukon-Kuskokwim Delta (61ǫ N 164ǫ W) in Subarctic western Alaska and B) the Arctic Coastal Plain (70ǫ N 154ǫ) in Arctic Alaska.

Fig. 5 in Helminth parasites of howler and spider monkeys in Mexico: Insights into molecular diagnostic methods and their importance for zoonotic diseases and host conservation

Fig. 5. Phylogenetic trees based on 28S (left) and cox1 (right) sequences of Strongyloides eggs. Sequences obtained from the eggs are bold type and indicated with an *. Numbers at the nodes represent posterior probabilities from Bayesian inference.

Fig. 2 in Helminth parasites of howler and spider monkeys in Mexico: Insights into molecular diagnostic methods and their importance for zoonotic diseases and host conservation

Fig. 2. Egg morphotypes found in the faeces of Mexican primates. A) Trypanoxyuris sp., B) Controrchis biliophilus, arrow pointing to the two eyespot remnants; C) trematode, diagnosed as C. biliophilus by molecular data; D) unidentified ancylostomatid; E) Strongyloides sp.; F) unidentified ascarid. Scale bar is equal to 15 Mm.

Fig. 4 in Helminth parasites of howler and spider monkeys in Mexico: Insights into molecular diagnostic methods and their importance for zoonotic diseases and host conservation

Fig. 4. Phylogenetic tree based on 28S sequences of Controrchis biliophilus. Sequences obtained from the eggs are bold type and indicated with an *. Numbers at the nodes represent posterior probabilities from Bayesian inference. Host species are indicated within parenthesis.

Fig. 1 in Helminth parasites of howler and spider monkeys in Mexico: Insights into molecular diagnostic methods and their importance for zoonotic diseases and host conservation

Fig. 1. Surveyed sites for parasites in Mexican primates. Dots indicate sampling sites, black: Alouatta palliata; white: A. pigra; and grey: Atetes geoffroyi. Polygons indicate the primate distribution range in Mexico, diagonal lines: A. palliata; dashes: A. pigra; and grey: A. geoffroyi.

Fig. 3 in Helminth parasites of howler and spider monkeys in Mexico: Insights into molecular diagnostic methods and their importance for zoonotic diseases and host conservation

Fig. 3. Phylogenetic trees based on 28S (left) and cox1 (right) sequences of Trypanoxyuris sp. Sequences obtained from the eggs are bold type and indicated with an *. Numbers at the nodes represent posterior probabilities from Bayesian inference.

opencc-by-4.0Dec 2018View details →

Fig. 3 in A comparison of helminth infections as assessed through coprological analysis and adult worm burdens in a wild host

Fig. 3. The relationship between adult lungworm (A. falciformis) burden and number of larvae per g of faeces in Irish badgers (n = 289).

opencc-by-4.0Dec 2018View details →

Fig. 2 in A comparison of helminth infections as assessed through coprological analysis and adult worm burdens in a wild host

Fig. 2. The relationship between adult hookworm (U. criniformis) burden and faecal egg counts (e.p.g) in Irish badgers (n = 289).

Fig. 1 in Helminths in common eiders (Somateria mollissima): Sex, age, and migration have differential effects on parasite loads

Fig. 1. Distributions of breeding and wintering populations of S. m. borealis, S. m. dresseri, and S. m. sedentaria in North America and Greenland. Populations of S. m. borealis are wintering in two areas in the pictured region; one in Southwest Greenland and one in East Canada (Newfoundland and Labrador). Individuals wintering in Southwest Greenland migrate to breed in West Greenland or Arctic Canada, whereas individuals wintering in East Canada have breeding areas in Arctic Canada (illustrated by the different direction of the red diagonal lines). For S. m. dresseri the breeding and wintering ranges overlap in one area that covers Newfoundland and Labrador as well as the northeastern part of the US, as shown by the grey horizontal lines. The subspecies S. m. sedentaria has its year-round residence in the Hudson Bay area as shown by the blue vertical lines. Dark stars mark the sampling locations of eiders in this study, whereas the white star marks the sampling location of eiders by Tourangeau et al. (2018). (For interpretation of the references to colour in this figure legend, the reader is referred to the Web version of this article.)

Fig. 3 in Helminths in common eiders (Somateria mollissima): Sex, age, and migration have differential effects on parasite loads

Fig. 3. Prevalences of the cestodes Lateriporus sp. (A) and Microsomacanthus spp. (B), and the acanthocephalan Profilicollis sp. (C) in common eiders. Abbreviations: bor, CD = S. m. borealis, Cape Dorset; bor, GRLD = S. m. borealis, Greenland; bor, NFLD = S. m. borealis, Newfoundland; dre, NFLD = S. m. dresseri, Newfoundland; sed, BI = S. m. sedentaria [data published in Tourangeau et al. (2018)]. n-m = non-migratory, po-m = post-migratory, prm = pre-migratory. Letters describe significant differences between groups: if two groups share a letter, there is no significant difference in their prevalences.

Fig. 2 in Helminths in common eiders (Somateria mollissima): Sex, age, and migration have differential effects on parasite loads

Fig. 2. Examples of gastrointestinal parasites retrieved from common eiders in this study. (A) The trematode Notocotylus sp., (B) the cestode Lateriporus sp., (C) the acanthocephalan, Profilicollis sp. (D) microphallid trematodes, and (E) Microsomacanthus spp. cestodes.

Fig. 5 in Helminths in common eiders (Somateria mollissima): Sex, age, and migration have differential effects on parasite loads

Fig. 5. Cumulative percentages of birds infected with (A) Microsomacanthus spp., (B) Microphallus spp. and (C) Gymnophallus spp. from five different locations. Legend denotes the five infection levels: 0, 1s, 10s, 100s, and 1000s of parasite individuals within a single host. Abbreviations: bor, GRLD = S. m. borealis, Greenland; bor, NFLD = S. m. borealis, Newfoundland; dre, NFLD = S. m. dresseri, Newfoundland; bor, CD = S. m. borealis, Cape Dorset; sed, BI = S. m. sedentaria, Belcher Islands [data published in (Tourangeau et al., 2018)]. nm = non-migratory, po-m = post-migratory, pr-m = pre-migratory.