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Fig. 1 in Not playing by the rules: Unusual patterns in the epidemiology of parasites in a natural population of feral horses (Equus caballus) on Sable Island, Canada

Fig. 1. Map of Sable Island, Canada, which is about 50 km long, 1 km wide at its widest point, and in total, 34 km2 (from Gold et al., 2019).

Fig. 3 in Not playing by the rules: Unusual patterns in the epidemiology of parasites in a natural population of feral horses (Equus caballus) on Sable Island, Canada

Fig. 3. Proportions of third-stage larvae of large and small strongyle species cultured from feces of 81 Sable Island horses in summer 2014, showing an unusual dominance of S. equinus in adult horses. Larvae with a rhabditiform pharynx were rare in young (1–3 years) and adult horses (≥3 years), but common in foals, which could represent larvae of Strongyloides westeri.

Fig. 1 in Investigating the role of urbanisation, wetlands and climatic conditions in nematode parasitism in a large Australian elapid snake

Fig. 1. The abundance of nematode infection for each tiger snake specimen in South-West Western Australia. Arrows indicate major cities used as urban centres. Colour indicates the number of worms (intensity) found in the stomach of each specimen. (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 Investigating the role of urbanisation, wetlands and climatic conditions in nematode parasitism in a large Australian elapid snake

Fig. 3. Probability of tiger snake stomach nematode infection in relation to a) distance to wetlands, b) mean annual precipitation and c) topographic wetness index (TWI). Shaded areas represents 95% confidence intervals.

Fig. 1 in Stable isotope analysis spills the beans about spatial variance in trophic structure in a fish host - parasite system from the Vaal River System, South Africa

Fig. 1. Map of the Vaal River showing the position of sampling sites (I: below Grootdraai Dam; II: Vaal Dam; III: below Vaal River Barrage; IV: Bloemhof Dam; V: below Vaalharts Weir; VI: Douglas Weir) along the Vaal River. The block (B) indicates the position of the Vaal River within South Africa and insert A indicates the position of South Africa shaded on the African continent.

Fig. 2 in Parasite species co-occurrence patterns on Peromyscus: Joint species distribution modelling

Fig. 2. Results of variance partitioning for variation in ectoparasite prevalence explained by fixed and random effects for each ectoparasite species (columns). Explained variance presented for the constrained model for deer mice (n = 229 individuals). DM, deer mice; RBV, southern red-backed vole; WJM, woodland jumping mouse; PA, population abundance. Population abundance of small mammal species measured as captures per 100 trap nights. (For interpretation of the references to colour in this figure legend, the reader is referred to the Web version of this article.)

Fig. 2 in Patterns of parasite eggs, oocysts and larvae shedding by moose in the Biebrza marshland (NE Poland)

Fig. 2. The relationship between the EPG of Trichostrongylidae family members, Moniezia spp., Parafasciolopsis fasciolaemorpha, the LPG of Elaphostrongylus sp. and mean monthly temperature (red continuous line). Blue continuous line shows the median EPG/LPG and broken lines indicate its 95% confidence interval. (For interpretation of the references to colour in this figure legend, the reader is referred to the Web version of this article.)

Fig. 1 in Patterns of parasite eggs, oocysts and larvae shedding by moose in the Biebrza marshland (NE Poland)

Fig. 1. The relationship between the prevalence of Nematodirella alcidis and Moniezia spp. eggs and mean monthly temperature (red continuous line). Blue continuous line shows the prevalence and broken lines indicate its 95% confidence interval. (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 Patterns of parasite eggs, oocysts and larvae shedding by moose in the Biebrza marshland (NE Poland)

Fig. 3. The relationship between the median EPG of Parafasciolopsis fasciolaemorpha, the median LPG of Elaphostrongylus sp. and the presence of snow cover.

Fig. 2 in Sensitive, quantitative detection of Besnoitia darlingi and related parasites in intermediate hosts and to assess felids as definitive hosts for known and as-yet undescribed related parasite species

Fig. 2. Coccidia-specific PCR to confirm the presence of DNA of various parasite species used to test the analytic specificity of the BdanjoRT1 real-time PCR. (1) Besnoitia darlingi, (2) B. neotomofelis, (3) B. oryctofelisi, (4) B. besnoiti (Evora isolate), (5) B. bennetti (Texas), (6) B. tarandi (Bt-CA-Quebec1), (7) Toxoplasma gondii, (8) Hammondia hammondi, (9) Neospora caninum, (10) H. heydorni, (11) Cystoisospora felis, (12) C. rivolta, (13) C. burrowsi, (14) C. canis, (15) Sarcocystis cruzi and (16) Crytosporidium parvum. Presence of (17) Giardia duodenalis and (18) Tritrichomonas foetus DNA was shown by amplification using species or genus-specific primers, respectively. C, negative control; M, marker.

Fig. 3 in Sensitive, quantitative detection of Besnoitia darlingi and related parasites in intermediate hosts and to assess felids as definitive hosts for known and as-yet undescribed related parasite species

Fig. 3. Analytic sensitivity (A) and standard curve (B) obtained for the threshold cycle (Cq) values obtained in the BdanjoRT1 real-time PCR using varying amounts of genomic Besnoitia darlingi DNA (approximately equivalent to the DNA content of 10.000 [blue], 1000 [green], 100 [red], 10 [brown] and 1 [black] B. darlingi tachyzoites) diluted in 100 ng/μl mouse DNA. Cq values used for regression are displayed as circles. Results on samples resembling DNA of 0.1 tachyzoite were not included in regression, since only two of four samples had reacted with a Cq value of 37.8 or 38.3 (displayed grey in A and as crosses in B). (For interpretation of the references to colour in this figure legend, the reader is referred to the Web version of this article.)

Fig. 1 in Sensitive, quantitative detection of Besnoitia darlingi and related parasites in intermediate hosts and to assess felids as definitive hosts for known and as-yet undescribed related parasite species

Fig. 1. Location of the primers and the probe of the Besnoitia darlingi/B.neotomofelis /B. oryctofelisi-specific real-time PCR assay BdanjoRT1 within the ITS-1 region of the rRNA gene. The sequences of the ITS-1 region of B. akodoni (AY545987, bold), B. jellisoni (AF076860, bold), B. neotomofelis (HQ909085, bold), B. darlingi (AF489696, bold) and B. oryctofelisi (AY182000, bold), were aligned relative to sequences of other Besnoitia spp. including B. besnoiti from Portugal, Spain and Germany, B. bennetti reported from the USA and Belgium, B. tarandi from Canada and Finland and those of Neospora caninum, Hammondia heydorni, Toxoplasma gondii and H. hammondi by using Clustal V (DNAStar, Madisin, Wisconsin, USA). Deletions and substitutions in the sequences relative to and within the clade of B. acodoni, B. jellisoni, B. neotomofelis, B. darlingi and B. oryctofelisi are indicated by black background. Sequences of the primer BdanjoRev and the Probe Bb11-12 are displayed in their complementary form. The probe Bb11-12 was established for a real-time PCR to detect B. besnoiti, but it is universal and can be used for the detection of all Besnoitia spp. mentioned here.

Fig. 1 in Diversity of helminth parasites of freshwater fish in the headwaters of the Coatzacoalcos River, in Oaxaca, Mexico

Fig. 1. The upper Coatzacoalcos river in Mexico showing the fish Collection sites; codes: 1. El Platanillo river, tributary to Del Sol river (municipality Santo Domingo Petapa), coordinates 16.951111, −95.244167, altitude 416 m; 2. Río Grande (El Barrio), 16.792167, −95.016083, 220 m; 3. Río Negro (Santa María Chimalapa), 16.898528, −94.693694, 166 m; 4. Río Modelo (Santa María Chimalapa), 17.134778, −94.745000, 115 m; 5. Río Pánfilo (Matías Romero, Oaxaca), 17.083639, −94.873944, 60 m; 6. Río Jaltepec (Jesús Carranza, Veracruz), 17.388444, −95.056111, 40 m; 7. Río Escondido (Paraje San Francisco El Vado, Agencia Municipal Río Escondido, Santa María Chimalapa), 17.091083, −94.751694, 103 m. Note all sites in Oaxaca state, except # 6.

Fig. 2 in Temporal dynamics of species associations in the parasite community of European eels, Anguilla anguilla, from a coastal lagoon

Fig. 2. Intensity of infection (mean ± SE number of parasites per host, including infected hosts only) of the six most common helminth parasites of eels, Anguilla anguilla, in Comacchio Lagoons, during three sampling periods. Graphs on the right-hand side do not include the 2015–2017 period, as these species were not found during that period. See Table 1 for full species names.

Fig. 4 in Temporal dynamics of species associations in the parasite community of European eels, Anguilla anguilla, from a coastal lagoon

Fig. 4. Pairwise relationships between numbers of parasites per host for the three most common digenean parasites of eels, Anguilla anguilla, in Comacchio Lagoons, across all three sampling periods combined. The line represents the relationship (with 95% confidence intervals) predicted by the generalized linear model; see text. Tick marks indicate partial residuals with either positive (top) or negative values (bottom). See Table 1 for full species names.

Fig. 3 in Temporal dynamics of species associations in the parasite community of European eels, Anguilla anguilla, from a coastal lagoon

Fig. 3. Scatterplots of pairwise relationships between numbers of parasites per host for the three most common digenean parasites of eels, Anguilla anguilla, in Comacchio Lagoons, across all three sampling periods combined. See Table 1 for full species names.

Fig. 1 in First detailed records of water mite larvae (Hydrachnidia: Hydrovolzidae, Hydryphantidae) parasitizing empidid flies (Diptera: Empididae: Clinocerinae)

Fig. 1. Types of habitat where infested and uninfested clinocerine species were found; a – the Mała Łąka Valley, June 2017; b – the same locality, October 2017; c – the Kościeliska Valley, May 2018; d – the same locality, November 2018; e – Białka River, Łysa Polana, August 2017; f – Cracow Gorge, June 2017 (photographs by I. Słowińska).

Fig. 5 in Parasitic development in intestines and oocyst shedding patterns for infection by Eimeria uekii and Eimeria raichoi in Japanese rock ptarmigans, Lagopus muta japonica, protected by cages in the Southern Japanese Alps

Fig. 5. Eimeria oocysts (E. uekii) isolated from soil inside the cage (cage No. 6) (A) on Mt. Kita (35̊40′N, 138̊14′E), one of their habitats in the southern Japanese Alps and on Mt. Norikura as other habitats on northern Japanese Alps (B). In Fig. A, the sporocysts and sporozoites are clearly formed.

Fig. 4 in Parasitic development in intestines and oocyst shedding patterns for infection by Eimeria uekii and Eimeria raichoi in Japanese rock ptarmigans, Lagopus muta japonica, protected by cages in the Southern Japanese Alps

Fig. 4. Number of oocysts per gram (OPG) as seasonal detection rate for E. raichoi of hens (solid bars) and chicks (open bars) in cage Nos. 4–6 in 2019. Double arrows show the periods during which feces of hens were examined. ND indicates that we could not collect feces and did not determine the OPG.

Fig. 2 in Parasitic development in intestines and oocyst shedding patterns for infection by Eimeria uekii and Eimeria raichoi in Japanese rock ptarmigans, Lagopus muta japonica, protected by cages in the Southern Japanese Alps

Fig. 2. Histopathological photomicrograph of a section of the intestines of dead chicks during cage protection in 2018. Figs. A and B show developmental trophozoites (arrows) and schizonts (arrows) of Eimeria spp. at the epithelial cells of the colon (Chick c) and ileum (Chick a), respectively. Some zoites (arrows) invade into submucosa (ileum of Chick b) (Fig. C). Figs. D, E, and F show the sexual zoites or cavities after releasing oocysts (arrows) of the ileum (Chick a), ceca (Chick d), and ileum (Chick c). Arrowheads in Fig. F indicate hemorrhages in the intestinal mucosa.