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Fig. 2 in Morphological and morphometric differentiation of dorsal-spined first stage larvae of lungworms (Nematoda: Protostrongylidae) infecting muskoxen (Ovibos moschatus) in the central Canadian Arctic

Fig. 2. Laboratory guide for the differentiation of L1 of U. pallikuukensis and V. eleguneniensis. The guide is based on key morphological features supported by morphometric data of heat killed L1. These features were characteristic of each of the species as visible under 400 × magnification.

Fig. 1 in Morphological and morphometric differentiation of dorsal-spined first stage larvae of lungworms (Nematoda: Protostrongylidae) infecting muskoxen (Ovibos moschatus) in the central Canadian Arctic

Fig. 1. Morphology of first stage larva (L1) of Umingmakstrongylus pallikuukensis. Photomicrograph of U. pallikuukensis L1 taken at 400 × magnification in differential interference contrast indicating the location of relevant important anatomical structures.

Fig. 3 in Morphological and morphometric differentiation of dorsal-spined first stage larvae of lungworms (Nematoda: Protostrongylidae) infecting muskoxen (Ovibos moschatus) in the central Canadian Arctic

Fig. 3. Morphology of Cystocaulus ocreatus first stage larva (L1). Photomicrograph of C. ocreatus L1 collected from Uzbekistan (US National Parasite Collection No. 95144) taken at 400 × magnification in differential interference contrast showing ventral post-anal cuticular striations and overall elongated structure of the tail spike similar to U. pallikuukensis.

opencc-by-4.0Aug 2015View details →

Fig. 2 in Species of Angiostrongylus (Nematoda: Metastrongyloidea) in wildlife: A review

Fig. 2. Tawny frogmouth, Podargus strigoides, with severe posterior paresis and unable to right itself due to infection with Angiostrongylus cantonensis.

opencc-by-4.0Aug 2015View details →

Fig. 3 in Species of Angiostrongylus (Nematoda: Metastrongyloidea) in wildlife: A review

Fig. 3. Immature Angiostrongylus cantonensis in the cerebellum of a brushtail possum, Trichosurus vulpecula, with extensive granulomatous and eosinophilic meningoencephalitis and malacia (adapted from Ma et al., 2013, fig. 7).

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Fig. 1 in Species of Angiostrongylus (Nematoda: Metastrongyloidea) in wildlife: A review

Fig. 1. The life cycle of Angiostrongylus cantonensis. Rat definitive hosts acquire thirdstage larvae by ingesting infected intermediate hosts, aquatic or terrestrial snails and slugs. Larvae penetrate the stomach, enter the hepatic portal and mesenteric lymphatic systems and are carried to the heart and lungs. They enter alveoli, invade the pulmonary veins, are returned to the left heart and distributed around the body by the arterial circulation. Larvae reach the CNS, predominantly the cerebrum and cerebellum, grow and moult twice in the parenchyma and young adults invade the subarachnoid space of the brain. After about two weeks they invade the cerebral vein and move to the heart and pulmonary arteries where they mature. Eggs are carried in the blood to the lungs where they embryonate. First-stage larvae escape up the bronchial escalator, are swallowed, pass out in the faeces, are ingested by intermediate hosts, snails and slugs, and develop to third-stage infective larvae. A broad spectrum of animals – planarians, prawns, crabs, frogs and lizards may serve as paratenic hosts in which infective larvae reside but undergo no further development. Humans are an accidental host and infection may occur through ingestion of intermediate or paratenic hosts, the latter often eaten raw or their juices used in preparation of local dishes. Infective larvae may also leave molluscs and contaminate vegetables such as lettuce (adapted from Wang et al., 2008, fig. 1).

opencc-by-4.0Dec 2013View details →

Figure 1 in A Nearctic parasite in a Palearctic host: Parelaphostrongylus andersoni (Nematoda; Protostrongylidae) infecting semi-domesticated reindeer in Alaska

Figure 1. Map of the Seward Peninsula, Alaska, showing semi-domesticated reindeer herd ranges including the Kakarak herd (bold), and reindeer herd loss to Western Arctic caribou herd; and the western limits of caribou migration from 1989 to 2000 (modified from Finstad et al. (2006) and Rattenbury et al. (2009)).

Fig. 1. Circular Bayesian tree inferred from mtDNA cox-2 in Temporal stability of parasite distribution and genetic variability values of Contracaecum osculatum sp. D and C. osculatum sp. E (Nematoda: Anisakidae) from fish of the Ross Sea (Antarctica)

Fig. 1. Circular Bayesian tree inferred from mtDNA cox-2 sequences obtained from specimens of C. osculatum sp. D and C. osculatum sp. E analysed in the present study, based on Bayesian Inference (BI) method using MrBayes v3.2.2 (Ronquist et al., 2012). Evolutionary distance was estimated using the TrN + G (G = 0.60) substitution model as implemented in jModeltest (Posada, 2008), with the AIC approach (Posada and Buckley, 2004). Posterior probability values are the result of 1.000000 of runs and are reported at the nodes. The coloured icons correspond to the two species considered in this study (red = C. osculatum sp. D and blue = C. osculatum sp. E).

Fig. 2 in Temporal stability of parasite distribution and genetic variability values of Contracaecum osculatum sp. D and C. osculatum sp. E (Nematoda: Anisakidae) from fish of the Ross Sea (Antarctica)

Fig. 2. Schematic distribution of the fish species examined in the present study for larval of C. osculatum sp. D and C. osculatum sp. E, along the continental shelf of the Ross Sea coastal ecosystem. Arrows indicating preferred preys and the diet preference for each fish species are reported according to the literature (La Mesa et al., 2004). The represented pelagic organisms comprise species of euphausiids and fish juveniles, benthic and epibenthic organisms are polychaetes, amphipods, decapods and gastropods. A pie chart with the relative proportions of C. osculatum sp. D and C. osculatum sp. E is given for each fish species. Squares and circles represent the hypothetical distribution of C. osculatum sp. D and C. osculatum sp. E larvae in their intermediate hosts.

Fig. 4 in Detection of cryptic species of Rugopharynx (Nematoda: Strongylida) from the stomachs of Australian macropodid marsupials

Fig. 4. Phylogenetic relationships of species of Rugopharynx and Rugonema labiatum based on a neighbor-joining analysis of the sequence data of the ITS+ nuclear ribosomal DNA. Values above and below branches represent the NJ and MP bootstrap values (respectively) that were greater than 70%.

Fig. 3 in Detection of cryptic species of Rugopharynx (Nematoda: Strongylida) from the stomachs of Australian macropodid marsupials

Fig. 3. Phylogenetic relationships of species of Rugopharynx and Rugonema labiatum based on a Bayesian analysis of the sequence data of the ITS+ nuclear ribosomal DNA. Values above branches indicate posterior probabilities that were greater than 0.8. Abbreviations of Australian state names are provided in Table 1.

Fig. 2 in Detection of cryptic species of Rugopharynx (Nematoda: Strongylida) from the stomachs of Australian macropodid marsupials

Fig. 2. Morphological buccal capsule types in the genus Rugopharynx. I, simple cylindrical buccal capsule, R. macropodis; IIA, bilobed buccal capsule with subequal divisions, R. epsilon; IIB, bilobed buccal capsule with anterior lobe shorter, R. rufogrisea; III, trilobed buccal capsule, R. longibursaris.

Fig. 1 in Detection of cryptic species of Rugopharynx (Nematoda: Strongylida) from the stomachs of Australian macropodid marsupials

Fig. 1. Localities within Australia at which specimens of Rugopharynx used in this study were collected. Coordinates for each locality are provided in Table 1. 1, Lake Clifton; 2, Waroona; 3, Collie, Wellington Dam; 4, Perup River; 5, Kalgoorlie; 6, Wallerberdina Station; 7, Port Augusta; 8, Ashbourne; 9, Kangaroo Island; 10, Naracoorte; 11, Hattah Lakes National Park; 12, Yan Yean; 13, The Gurdies; 14, Launceston; 15, Emu Flat, Bondo State Forest; 16, Trangie; 17, Grafton; 18, Lamington National Park; 19, Miles; 20, Dawes; 21, Mt Sebastopol; 22, Rockhampton; 23, Winton; 24, Proserpine; 25, Bowen; 26, Magnetic Island; 27, Lake Barrine.

Fig. 3 in Temporal stability of parasite distribution and genetic variability values of Contracaecum osculatum sp. D and C. osculatum sp. E (Nematoda: Anisakidae) from fish of the Ross Sea (Antarctica)

Fig. 3. Schematic representation of the hypothetic life-cycle of C. osculatum sp. D (a) and C. osculatum sp. E (b) in the Ross Sea.

Fig. 5 in Detection of cryptic species of Rugopharynx (Nematoda: Strongylida) from the stomachs of Australian macropodid marsupials

Fig. 5. Molecular phylogeny of species of Rugopharynx and Rugonema labiatum based on a consensus of the BI, NJ and MP trees (Figs. 3 and 4), and the relationships of their hosts. This figure includes species of Macropus (M. agilis, M. antilopinus, M. bernardus) which are not hosts to species of Rugopharynx, as well as M. parma, from which no material could be obtained for genetic studies. Only those species of Petrogale included in this study are shown on the host tree. The morphology of the buccal capsule for each nematode taxon is also shown.

opencc-by-4.0Dec 2016View details →

Fig. 2 in A new species of Potoroxyuris (Nematoda: Oxyuridae) from the woylie Bettongia penicillata (Marsupialia: Potoroidae) from southwestern Australia

Fig. 2. Photomicrographs of Potoroxyuris keninupensis n. sp. and P. potoroo. (A) eggs of P. keninupensis n. sp. from faeces, one with the operculum open. (B) En face view of a female specimen of P. keninupensis n. sp. showing pharyngeal lobes. The amphid (am) and both submedian papillae (p) are in focus on the left side of the oral opening. The granule to the right of the dorsal lobe is an artefact. (C) Dorso-lateral optical section of a ventro-lateral pharyngeal lobe of allotype female P. keninupensis n. sp. (D) Lateral view of a ventro-lateral pharyngeal lobe of a female P. potoroo from AHC 47720. Scale bars: (A‾B) 50 Mm, (C‾D) 20 Mm.

Fig. 7 in First description of Onchocerca jakutensis (Nematoda: Filarioidea) in red deer (Cervus elaphus) in Switzerland

Fig. 7. Microfilaria showing transverse annulation and irregular shape of swollen anterior end (magnitude of annulation and swelling might be exaggerated by artefact of fixation).

Fig. 5 in First description of Onchocerca jakutensis (Nematoda: Filarioidea) in red deer (Cervus elaphus) in Switzerland

Fig. 5. External cuticular annulation (A) of O. jakutensis female with interruption over lateral field.

Fig. 4. a and b in First description of Onchocerca jakutensis (Nematoda: Filarioidea) in red deer (Cervus elaphus) in Switzerland

Fig. 4. a and b: Posterior end of O. jakutensis male with 5 pairs of pericloacal papillae without unpaired precloacal papilla.

Fig. 3. O in First description of Onchocerca jakutensis (Nematoda: Filarioidea) in red deer (Cervus elaphus) in Switzerland

Fig. 3. O. jakutensis male and female (bars: 100 Mm): a. Tail of male with five pairs of pericloacal and two pair of closely spaced terminal papillae. b. Tail of another male with more distantly spaced papillae on tail end. Left spicule protruding. c. Spicules in ventral view. d. Head end of male. e. Head end of female with vulva. f. Posterior end of female with conical tail in ventral view, annulations indicated on sides. g. Microfilaria with terminal distribution of nuclei.