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Fig. 1 in Spread of moniesiosis pathogens in livestock in the Ganja-Gazakh Region of the Republic of Azerbaijan: Bio-ecological features

Fig. 1. Spread of moniesiosis pathogens in sheep in different landscape/ecological territories Рис. 1. Распространение возбуÃитеΛей мониезиоза среÃи овец в ΛанÃшафтно-экоΛоги- ческих зонах

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Fig. 2 in Spread of moniesiosis pathogens in livestock in the Ganja-Gazakh Region of the Republic of Azerbaijan: Bio-ecological features

Fig. 2. Spread of moniesiosis pathogens in goats in different landscape/ecological territories Рис. 2. Распространение возбуÃитеΛей мониезиоза коз по ΛанÃшафтно-экоΛогическим зонам

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Fig. 3 in Exploiting parallels between livestock and wildlife: Predicting the impact of climate change on gastrointestinal nematodes in ruminants

Fig. 3. In marginal grazing systems in Europe sheep often occupy separate summer and winter grazing areas, analogous to the summer and winter ranges of migratory ruminants. In the uplands of Wales, UK, (shown here) sheep are often grazed on extensive areas of land at low stocking densities over the summer period, and sent to lowland dairy farms for winter grazing at higher stocking densities. (Photo: Rose, H.).

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Fig. 2 in Exploiting parallels between livestock and wildlife: Predicting the impact of climate change on gastrointestinal nematodes in ruminants

Fig. 2. The relative seasonal incidence of ovine parasitic gastroenteritis (PGE) in the Southwest of England, UK, based on monthly diagnoses of (a) Nematodosis (NOS = species not otherwise specified), (b) Haemonchosis and (c) Nematodirosis (van Dijk et al., 2008).

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Fig. 1 in Exploiting parallels between livestock and wildlife: Predicting the impact of climate change on gastrointestinal nematodes in ruminants

Fig. 1. Comparison of the instantaneous daily development rate of Ostertagia ostertagi (grey) and O. gruehneri (black) at a range of constant temperatures. Instantaneous daily development rates were estimated from the time to 50% development of L3, derived from data published in the literature (O. ostertagi: Rose, 1961; Pandey, 1972; Young et al., 1980) and original data (O. gruehneri: Hoar, 2012) as described by Azam et al. (2012).

Fig. 2 in Occurrence of tongue worm, Linguatula cf. serrata (Pentastomida: Linguatulidae) in wild canids and livestock in south-eastern Australia

Fig. 2. In the petri dishes, adult males and females removed from the left and right nasal cavities of a wild dog; Right image shows the anterior end of a nymph removed from lymph node of a cow.

Fig. 1 in Occurrence of tongue worm, Linguatula cf. serrata (Pentastomida: Linguatulidae) in wild canids and livestock in south-eastern Australia

Fig. 1. Left: A female tongue worm being removed from the nasal cavity of a wild dog; Right: Two encapsulated tongue worm nymphs in a bovine mesenteric lymph node.

Fig. 4 in Occurrence of tongue worm, Linguatula cf. serrata (Pentastomida: Linguatulidae) in wild canids and livestock in south-eastern Australia

Fig. 4. Tongue worm eggs found in the faeces of a wild dog in the present study. Arrows indicate a pair of hooks.

Fig. 3. Localities where animals infected with L in Occurrence of tongue worm, Linguatula cf. serrata (Pentastomida: Linguatulidae) in wild canids and livestock in south-eastern Australia

Fig. 3. Localities where animals infected with L. cf. serrata were found. Open and solid symbols represent uninfected and infected animals, respectively. The stars, the red circles and the blue circles are cattle, wild dogs and foxes, respectively. Those animals that have been collected opportunistically have been excluded from this map. A) Creighton's Creek, B) Tallangatta, C) Mitta Mitta, D) Black Mountain, E) Corryong, F) Tumbarumba, G) Limestone, H) Wee Jasper, I) Brindabella, J) Mullion, K) Bullen Range, L) Booroomba, M) Perisher Valley, N) Orbost, O) Wabonga, P) Wangaratta, Q) Wodonga, R) Holbrook, S) Wagga Wagga, T) Towong, U) Braidwood, V) Carcoar. (For interpretation of the references to colour in this figure legend, the reader is referred to the web version of this article.)

Fig. 5 in Value of forest remnants for montane amphibians on the livestock grazed Mount Mbam, Cameroon

Fig. 5. Non-metric dimensional scaling plot of amphibian community structure divided by land use type on Mount Mbam based on visual encounter surveys with equal effort for each land use. The PERMANOVA p-value is shown in the top right corner.

Fig. 4 in Value of forest remnants for montane amphibians on the livestock grazed Mount Mbam, Cameroon

Fig. 4. Montane endemic amphibian species observed in recent surveys of Mount Mbam, West-Region, Cameroon. a) Astylosternus rheophilus, b) Astylosternus montanus, c) Afrixalus aff. fulvovittatus, d) Hyperolius balfouri, e) Hyperolius igbettensis, f) Hyperolius nitidulus, g) Hyperolius concolor, h) Hyperolius cinnamomeoventris, i) Hyperolius tuberculatus, j) Leptopelis nordequatorialis, k) Leptopelis boulengeri, l) Phrynobatrachus steindachneri, m) Xenopus cf. eysoole, n) Hoplobatrachus occipitalis, and o) Sclerophrys maculata.

Fig. 2 in Value of forest remnants for montane amphibians on the livestock grazed Mount Mbam, Cameroon

Fig. 2. Montane habitats of amphibian species observed in recent surveys of Mount Mbam, West-Region, Cameroon: a) gallery forest during the rainy season; b): gallery forest during the dry season after a bushfire; c) savanna area transformed by overgrazing; and d): effects of bushfire started for pasture on the same site during the dry season.

Fig. 1 in Value of forest remnants for montane amphibians on the livestock grazed Mount Mbam, Cameroon

Fig. 1. Maps showing (top) the topography of the Bamenda Highlands, white circle showing Mount Mbam in the West Region of Cameroon; and (bottom) the layout of sample sites on Mount Mbam.

Fig. 6 in Prevalence and gross pathology of liver fluke in macropods cohabiting livestock farms in north eastern NSW, Australia, and diagnosis using cELISA

Fig. 6. Livestock farms in the Northern Tablelands region of NSW, Australia, with Macropods harbouring liver fluke infections (December 2018–June 2021).

Fig. 5 in Prevalence and gross pathology of liver fluke in macropods cohabiting livestock farms in north eastern NSW, Australia, and diagnosis using cELISA

Fig. 5. Scatter plot of Fasciola hepatica coproantigen concentration (optical density, 450 nm) and total fluke count in Macropods.

Fig. 4. A in Prevalence and gross pathology of liver fluke in macropods cohabiting livestock farms in north eastern NSW, Australia, and diagnosis using cELISA

Fig. 4. A. Common wallaroo liver (visceral surface) with prominent fibrotic capsules. B. Liver cross-section of fibrous capsules. C. Eastern grey kangaroo liver (visceral surface) with irregular form, hepatomegaly, fibrotic lesions and bile duct hyperplasia. D. Necrotic tracks generated by immature fluke. E. Immature fluke (mm).

Fig. 3. Rainfall and temperature data throughout 2019–2020 in Prevalence and gross pathology of liver fluke in macropods cohabiting livestock farms in north eastern NSW, Australia, and diagnosis using cELISA

Fig. 3. Rainfall and temperature data throughout 2019–2020 recorded at the Armidale airport NSW, Australia (Australian Government of Bureau of Meteorology, 2019, 2020).

Fig. 1 in Prevalence and gross pathology of liver fluke in macropods cohabiting livestock farms in north eastern NSW, Australia, and diagnosis using cELISA

Fig. 1. Geographical location of livestock farms (A–G) surveyed in the Northern Tablelands region of NSW, Australia, to assess liver fluke prevalence in Macropods (ArcGIS 10.4.1 software, 2018).

Fig. 2 in Prevalence and gross pathology of liver fluke in macropods cohabiting livestock farms in north eastern NSW, Australia, and diagnosis using cELISA

Fig. 2. Liver fluke prevalence in Macropods (infected/total sampled) cohabiting farms in the Northern Tablelands region of NSW, Australia. Number of farms by risk site: low – 2 farms, medium – 3 farms, high – 2 farms.