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Fig. 2 in Using occupancy models to investigate the prevalence of ectoparasitic vectors on hosts: An example with fleas on prairie dogs

Fig. 2. The robust design for occupancy models of flea prevalence on black-tailed prairie dogs (Cynomys ludovicianus). Prairie dogs were sampled during primary occasions in different months of the year (May–September 2012). Each primary occasion comprised three secondary occasions (combings) during which fleas might be detected (p = probability of detection, given presence). A prairie dog was ''open'' to colonization by fleas between primary occasions. Once a prairie dog was colonized, it was occupied by fleas during all subsequent primary occasions (thus, the extinction probability, E, was fixed at zero, once a prairie dog was occupied by fleas). Closure was assumed during the secondary occasions, but we used behavioral covariates to account for removal of fleas from hosts during each secondary combing (REMOVAL1 and REMOVAL2, see text). In the example encounter history, a '1' indicates that at least one flea was detected during a combing event, and a '0' indicates that no fleas were detected.

Fig. 5 in Using occupancy models to investigate the prevalence of ectoparasitic vectors on hosts: An example with fleas on prairie dogs

Fig. 5. Model-averaged probabilities of flea occupancy (W) and flea colonization (γ) for black-tailed prairie dogs (Cynomys ludovicianus) in old and young colonies, and natural and translocation colonies during May–September 2011, at the Vermejo Park Ranch, New Mexico (see Fig. 1 and text for colony descriptions). Bars depict 95% confidence intervals. We do not report estimates of colonization for September, because few prairie dogs were sampled in that month.

Fig. 1 in Using occupancy models to investigate the prevalence of ectoparasitic vectors on hosts: An example with fleas on prairie dogs

Fig. 1. Map of the study area within the Vermejo Park Ranch, Colfax County, New Mexico, showing old and young, and natural and translocation colonies of black-tailed prairie dogs (Cynomys ludovicianus). Gray areas indicate extent of prairie dog colonies in 2009.

Fig. 6 in Using occupancy models to investigate the prevalence of ectoparasitic vectors on hosts: An example with fleas on prairie dogs

Fig. 6. Probabilities of flea occupancy (W) and flea colonization (γ) for black-tailed prairie dogs (Cynomys ludovicianus) in plots with differing densities of prairie dogs during May–September 2011, at the Vermejo Park Ranch, New Mexico. Solid lines depict estimates and dotted lines depict 95% confidence intervals.

Fig. 3 in Differential patterns of definitive host use by two fish acanthocephalans occurring in sympatry: Pomphorhynchus laevis and Pomphorhynchus tereticollis

Fig. 3. Relative flow rates of P. tereticollis and P. laevis in their hosts in the Ouche (a) and Vingeanne (b) localities, taking into account the abundance of adult intestinal parasites and the relative abundance of each fish host species in the local community. The thickness of arrows and percentages indicate the estimated relative value of each fish species as a resource for each Pomphorhynchus species.

Fig. 6 in Differential patterns of definitive host use by two fish acanthocephalans occurring in sympatry: Pomphorhynchus laevis and Pomphorhynchus tereticollis

Fig. 6. Reproductive parameters of P. laevis and P. tereticollis: (a) testes volume as a function of adult worm size, according to parasite species and fish host; (b) number of ovarian balls according to parasite species and fish host; (c, d) number of eggs as a function of female size according to parasite species, in barbel (c) and chub (d).

Fig. 1 in Differential patterns of definitive host use by two fish acanthocephalans occurring in sympatry: Pomphorhynchus laevis and Pomphorhynchus tereticollis

Fig. 1. Percentage of P. laevis (plain bars) and P. tereticollis (striped bars) among Pomphorhynchus parasites collected from amphipod intermediate hosts in the benthos, and from the fish definitive hosts, either in the intestine or collected as extraintestinal cystacanths, in (a) the Ouche and (b) Vingeanne localities. The proportion of P. laevis and P. tereticollis in each fish species was compared to their proportion in intermediate gammarid hosts using Fisher exact test (P-values given after correction for multiple test: ***: P <0.001; **: P <0.01; *: P <0.05; ns: not significant). Letters above bars refer to post-hoc comparison between fish species (Tukey HSD). Numbers below bars are sample sizes (number of parasites). Bentho-pelagic fish species are abbreviated in italics, benthic species in standard font. Species abbreviations: Barbus barbus, Bbs; Gobio gobio, Gg; Squalius cephalus, Sc; Telestes souffia, Ts; Chondrostoma nasus, Cn; Rutilus rutilus, Rr; Scardinius erythrophthalmus, Se; Leuciscus leuciscus, Ll; Phoxinus phoxinus, Pp; Barbatula barbatula, Bba; Ameiurus melas, Am; Perca fluviatilis, Pf; Cottus gobio, Cg; Gasterosteus aculeatus, Ga.

Fig. 2 in Differential patterns of definitive host use by two fish acanthocephalans occurring in sympatry: Pomphorhynchus laevis and Pomphorhynchus tereticollis

Fig. 2. Relationship between fish biomass (g.100m-2) and P. laevis (a) or P. tereticollis (b) mean abundance per fish species, across fish species and localities (Ouche and Vingeanne localities). Both variables were log10-transformed.

Fig. 4 in Differential patterns of definitive host use by two fish acanthocephalans occurring in sympatry: Pomphorhynchus laevis and Pomphorhynchus tereticollis

Fig. 4. Relationship between the phylo-structural index of specificity towards definitive hosts and observed prevalence in gammarid intermediate hosts (%) in P. laevis and P. tereticollis. The phylo-structural index of specificity increases with the taxonomic diversity of fish species used.

Fig. 3 in Parasite community structure as a predictor of host population structure: An example using Callorhinchus capensis

Fig. 3. Randomized individual-based species accumulation curve of parasites infecting Callorhinchus capensis (n = 259) caught off the West and South Coasts of South Africa between 2010 and 2015.

Fig. 1 in Parasite community structure as a predictor of host population structure: An example using Callorhinchus capensis

Fig. 1. Map showing sample locations (St Helena Bay, False Bay and trawl locations shown by diamond symbols) in which Callorhinchus capensis were caught (dashed line indicates 200 m depth contour). The inset shows the distribution range of Callorhinchus capensis (IUCN, 2012).

Fig. 2. a–d in Parasite community structure as a predictor of host population structure: An example using Callorhinchus capensis

Fig. 2. a–d: Four metazoan parasites found infecting Callorhinchus capensis caught in False Bay, South Africa in 2013. Photos taken by T. Morris.

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Fig. 5. Native plant hosting M in Distribution, habitat use and plant associations of Moluchia brevipennis (Saussure, 1864) (Blattodea: Ectobiidae): an endemic cockroach from Chilean Mediterranean Matorral biome

Fig. 5. Native plant hosting M. brevipennis cockroaches at our study site. (A) Cockroaches feeding pollen from Oenothera acaulis (Onagraceae) flowers, (B) Puya venusta (Bromeliaceae) inflorescences with cockroaches reaching nectar sources, (C) Carpobrotus chilensis (Aizoaceae) with cockroach inside inflorescence and climbing branch, (D) Haplopappus chrysantemifolius (Asteraceae) with adult and nymphs feeding on florets from floral head, black bar indicates a 1 cm scale.

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Fig. 2. Bayesian tree reconstructed using 478 in Detection of haemosporidian parasites in wild and domestic birds in northern and central provinces of Iran: Introduction of new lineages and hosts

Fig. 2. Bayesian tree reconstructed using 478-bp mitochondrial cytb gene for avian blood parasites lineages. The amplified sequences in the current study are highlighted in bold. Posterior probability support of>0.8 is displayed for each branch. Schematic tree is summarized in section A and separated clade for each genus is given in sections of B (Plasmodium), C (Haemoproteus), and D (Leucocytozoon).

Fig. 5 in Host associations of gall-inducing Prodiplosis longifila (Diptera: Cecidomyiidae) from Bolivia: implications for its use as a biological control agent for Jatropha gossypiifolia (Euphorbiaceae)

Fig. 5. Susceptibility of Jatropha gossypiifolia to Prodiplosis longifila. Morphologically similar galls induced on control Jatropha clavuligera (A) and Jatropha gossypiifolia (B) plants under no-choice conditions in quarantine in Pretoria, South Africa. Galling on a transplanted Jatropha gossypiifolia plant under the natural field conditions in Bolivia (C).

Fig. 4 in Host associations of gall-inducing Prodiplosis longifila (Diptera: Cecidomyiidae) from Bolivia: implications for its use as a biological control agent for Jatropha gossypiifolia (Euphorbiaceae)

Fig. 4. Relationship between the number of branches per plant and number of shoot tips with Prodiplosis longifila galls (A) and percentage of shoots (± SE) with Prodiplosis longifila galls (B) across 3 sites in Bolivia in Feb 2016. Different letters above SE bars indicate significant differences (Tukey's test, P <0.001).

Fig. 3 in Host associations of gall-inducing Prodiplosis longifila (Diptera: Cecidomyiidae) from Bolivia: implications for its use as a biological control agent for Jatropha gossypiifolia (Euphorbiaceae)

Fig. 3. Incidence of Prodiplosis longifila galls on Jatropha clavuligera (percentage of plants with galls) in relation to (A) sites and (B) sampling yr and season (± SE). Different letters above SE bars indicate significant differences (Tukey's test, P <0.001).

Fig. 1 in Host associations of gall-inducing Prodiplosis longifila (Diptera: Cecidomyiidae) from Bolivia: implications for its use as a biological control agent for Jatropha gossypiifolia (Euphorbiaceae)

Fig. 1. Rosette galls induced by Prodiplosis longifila in the growing shoot tip of Jatropha clavuligera. (A) Rosette gall in the terminal and axillary buds of Jatropha clavuligera with swollen petiole and rachis and malformed leaves. (B) Young gall in Jatropha clavuligera showing early stage larvae feeding on the nutritive tissue of the gall with no evidence of cell necrosis and fungal growth.

Fig. 2 in Host associations of gall-inducing Prodiplosis longifila (Diptera: Cecidomyiidae) from Bolivia: implications for its use as a biological control agent for Jatropha gossypiifolia (Euphorbiaceae)

Fig. 2. Morphological and histological changes in the rosette galls induced by Prodiplosis longifila on Jatropha clavuligera. (A) Vertical sectional view of a portion of the galled shoot showing the inflamed axillary position (*) (bar = 1 mm). (B) Cross sectional view of a portion of the galled shoot showing the hyperplasied stems and leaves (bar = 0.33 mm); * indicates the location where the larvae congregate; le = leaf. (C) Cross sectional view of a galled leaf showing the intense hyperplasied upper mesophyll (bar = 0.18 mm); arrows point to the hypertrophied epidermis. (D) Cross sectional view of a leaf axil (bar = 0.33 mm); * indicates the location where the larvae congregate; note the vertically dividing epidermal cells towards the bottom, whereas the laterally occurring epidermal cells are hypertrophied. (E) Cross sectional view of the stem showing variously dividing cortical cells (bar = 30 μm); the arrow indicates cell proliferation. (F) Compactly arranged parenchymatous nutritive cells, each with a prominent nucleus (n) and dense cytoplasm (cyt) (bar = 10 μm). (G) Low magnification image of galled stem showing layers of nutritive cells (nc), where the larvae feed (lfs) (bar = 30 μm). In fact, the cells lying opposite, which appear to be empty, show signs of regeneration (rc). In the far interior of the stem cortex, numerous phenol-included cells (pic) occur. Far interior into the stem cortex numerous phenol including cells (pic) occur. (H) An enlarged view of a phenol-including cell (bar = 18 μm).