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Fig. 3 in Arthropod parasites of Antarctic and Subantarctic birds and pinnipeds: A review of host-parasite associations

Fig. 3. Genera of sucking lice (Echinophthiriidae – 1) and chewing lice (Menoponidae – 2, Philopteridae – 3) recorded infesting Antarctic birds and mammals.

Fig. 2 in Untapped potential: The utility of drylands for testing eco-evolutionary relationships between hosts and parasites

Fig. 2. Worldwide endemic and imported cases of (A) cutaneous leishmaniasis (CL) and (B) viceral leishmaniasis (VL) as of 2018. Warmer colors indicate a higher number of cases reported that year. Source: World Health Organization (2019).

Fig. 2 in Fish out of water: Aquatic parasites in a drying world

Fig. 2. Network of predicted changes in host and parasite communities in intermittent rivers as a consequence of decreased streamflow because of climate change.

Fig. 2 in Arthropod parasites of Antarctic and Subantarctic birds and pinnipeds: A review of host-parasite associations

Fig. 2. Host-parasite associations at the family level between Antarctic birds and mammals and arthropods, excluding stragglers and contaminants. Phylogenetic trees are not drawn to scale (adapted from Dabert and Mironov, 1999; Whiting, 2002; Dowling and O'Connor, 2010; Zhang, 2011; Prum et al., 2015).

Fig. 2. Flow diagrams showing a in "Weight of evidence" as a tool for evaluating disease in wildlife: An example assessing parasitic infection in Northern bobwhite (Colinus virginianus)

Fig. 2. Flow diagrams showing a weight of evidence framework using the (A) 7 questions proposed by Burkhardt-Holm and Scheurer (2007) and the (B) modified questions for addressing disease(s) in wildlife.

Fig. 3 in Adaptations, life-history traits and ecological mechanisms of parasites to survive extremes and environmental unpredictability in the face of climate change

Fig. 3. Flow chart outlining factors that can influence the response of parasites to climate change.

Fig. 4 in Mosquito identification and haemosporidian parasites detection in the enclosure of the African penguins (Spheniscus demersus) at the SANBI zoological garden

Fig. 4. Maximum likelihood tree showing the clustering of Leucocytozoon sp. (Clade I) and Plasmodium sp. (Clade II) with Haemoproteus sp. as outgroup. Sequences from this study are highlighted with red circles. (For interpretation of the references to color in this figure legend, the reader is referred to the Web version of this article.)

Fig. 3. Maximum likelihood tree for Culex species showing the 5 clades representing 5 subgroups. Clade I in Mosquito identification and haemosporidian parasites detection in the enclosure of the African penguins (Spheniscus demersus) at the SANBI zoological garden

Fig. 3. Maximum likelihood tree for Culex species showing the 5 clades representing 5 subgroups. Clade I is the Trifilatus Subgroup (Mattingly and Rageau, 1958) for Cx. torrentium; Clade II and III are the Pipiens Complex; Clade IV the Theileri Subgroup (Sirivanakarn, 1976) for Cx. theileri; and Clade V is the Tarsalis (Edwards, 1932) for Cx. declaratory and Apicinus Subgroups (Edwards, 1932) for Cx. mollis. Lutzia sp. used as outgroups. Sequences from this study are indicated by asterisks (*).

Fig. 1 in Mosquito identification and haemosporidian parasites detection in the enclosure of the African penguins (Spheniscus demersus) at the SANBI zoological garden

Fig. 1. Map of South Africa showing the National Zoological Gardens (NZG). The red star indicates where the African penguin enclosure is located and where mosquito samples were collected (Labuschagne et al., 2008). (For interpretation of the references to color in this figure legend, the reader is referred to the Web version of this article.)

Fig. 2. Predicted probabilities and 95 in Is it best on the nest? Effects of avian life-history on haemosporidian parasitism

Fig. 2. Predicted probabilities and 95% confidence intervals of haemosporidian parasitism (Plasmodium, Haemoproteus, and Leucocytozoon). Expected prevalence illustrated according to haemosporidia genera; Plasmodium represented with "P" (a), Haemoproteus represented with "H" (b–c), Leucocytozoon represented with "L" (d–f). Note that in some instances symbol size exceeded the range of confidence intervals.

Fig. 1 in Review on parasites of wild and captive giant pandas (Ailuropoda melanoleuca): Diversity, disease and conservation impact

Fig. 1. Distribution of wild giant pandas in six mountain regions (Qinling, Minshan, Qionglai, Liangshan, Daxiangling and Xiaoxiangling) in three Provinces (Gansu, Shaanxi, and Sichuan) of China. Adapted from Wang et al. (2018).

Fig. 1 in "Weight of evidence" as a tool for evaluating disease in wildlife: An example assessing parasitic infection in Northern bobwhite (Colinus virginianus)

Fig. 1. Timeline depicting the history of wildlife diseases in the United States: blue boxes are for disease reports and outbreaks, green for improvements to disease research, and red for events that hindered disease research. Abbreviations: foot-and-mouth disease (FMD), Smoot-Hawley Tariff Act (SHTA), State-Federal Cooperative Brucellosis Eradication Program (SFCBER), Bear River Research Station (BRRS), Wildlife Disease Investigations Laboratory (WDIL), Southeastern Cooperative Wildlife Disease Study (SCWDS), epizootic hemorrhagic disease (EHD), World Organisation for Animal Health's (OIE), National Wildlife Research Center (NWRC), U. S. Fish and Wildlife Service (USFWS). References: 1. Antolin et al. (2002), 2. Creel (1941), 3. Anderson (1978), 4. Locke and Friend (1987), 5. McCoy and Chapin (1912), 6. Wherry and Lamb (1914), 7. Meagher and Meyer (1994), 8. Clements (2007), 9, Bachrach (1968), 10. Busch and Parker (1972), 11. USFWS (1991), 12. Tunnicliff and Marsh (1935), 13. Brooks and Buchanan (1970), 14. Elton (1931), 15. Brown (2007), 16. CDFW 2019, 17. Friend (2014), 18. SCWDS 2019, 19. Shope et al. (1960), 20. Cohen (2000), 21. Cross et al. (2013), 22. Samuel et al. (2007), 23. Carvalho et al. (2017), 24. Dobson and Hudson (1986), 25. Jones et al. (2008), 26. Berger et al. (1998), 27. Laurance et al. (1996), 28. Collins and Crump (2009), 29. OIE 2008, 30. Voyles et al. (2015), 31. Fagerstone (2014), 32. USFWS (2016), 33. Scheele et al. (2019). (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 Fish out of water: Aquatic parasites in a drying world

Fig. 1. Conceptual diagram showing alternating cycles of flow, flow cessation and drying in intermittent rivers, with associated changes in the percentage of lotic, lentic and terrestrial habitats (adapted from Datry et al., 2016).

Fig. 1 in Arthropod parasites of Antarctic and Subantarctic birds and pinnipeds: A review of host-parasite associations

Fig. 1. Sub-areas of the Antarctic region. Legend: AAP = Antarctic Peninsula (including South Shetland Islands and Palmer Archipelago), AWS = Antarctica Weddell Sea sector, AAT = Antarctica Atlantic Ocean sector (including Bouvet Island), AIW = Antarctica Indian Ocean West sector, AIE = Antarctica Indian Ocean East sector, ARS = Antarctica Ross Sea sector (including Scott and Balleny Islands), APW = Antarctica Pacific Ocean West sector, APE = Antarctica Pacific Ocean East sector (including Peter I Island), SOI = South Orkney Island, SGI = South Georgia Island, SSI = South Sandwich Islands, PEI = Prince Edward Islands, CRI = Crozet Islands, KEI = Kerguelen Islands, HMI = Heard and McDonald Islands. The Antarctic Polar Front was drawn from Moore et al. (1999).

Fig. 7 in First molecular investigation of haemosporidian parasites in Thai bat species

Fig. 7. Phylogenetic (ML) relationship of the Polychromophilus cytb gene fragment (378 bp) found in this study (taxa with green circles) and the other global isolates. Polychromophilus isolated from T. melanopogon is placed in the same clade with African bat isolates of P. melanipherus in clade 1. Polychromophilus isolated from My. siligorensis shares the same clade with European bat isolates of P. murinus in clade 2. BS values greater than 50% are shown in the figure. The other clades, consisting of Hepatocystis, Nycteria, Plasmodium, and outgroup are collapsed. (For interpretation of the references to color in this figure legend, the reader is referred to the Web version of this article.)

Fig. 5 in First molecular investigation of haemosporidian parasites in Thai bat species

Fig. 5. Phylogenetic (ML) relationship of the Hepatocystis cytb gene fragment (378 bp) in the present study (taxa with red circles) and the other global isolates from the GenBank™ database. Clade (number) and subclade (letter) are indicated. Haplotype 10 of Hepatocystis in the current study originated from the frugivorous bat Cy. brachyotis shares the same clade with Malaysian isolate (in clade 2). The percentage of trees in which the associated taxa clustered together is shown next to the branches. BS values greater than 50% are shown in the figure. The other clades, consisting of Nycteria, Polychromophilus, Plasmodium, and outgroup are collapsed. (For interpretation of the references to color in this figure legend, the reader is referred to the Web version of this article.)

Fig. 2 in First molecular investigation of haemosporidian parasites in Thai bat species

Fig. 2. Images of bats with Hepatocystis positive infection based on either microscopic examination or cytb gene PCR amplification and sequencing results: H. larvatus (a), H. bicolor (b), H. armiger (c), H. lekakuli (d), R. malayanus (e), R. thomasi (f), R. pearsonii (g), C. thonglongyai (h), and Cy. brachyotis (i). Giemsa-stained blood smears depicting young gametocytes of Hepatocystis in ring and amoeboid forms (j–l), growing and fully-grown macrogametocytes (female) (m–o), growing and fully-grown microgametocytes (male) (p–r) observed in H. larvatus (sample ID THBat19-037, cytb accession no. MT136132). All parasite images were taken at the same magnification. Scale bar = 5 μm. (For viewing the images in color, the reader is referred to the online version of this article). (For interpretation of the references to color in this figure legend, the reader is referred to the Web version of this article.)

Fig. 3 in First molecular investigation of haemosporidian parasites in Thai bat species

Fig. 3. Images of bats with Nycteria positive infection based on either microscopic examination or cytb gene PCR amplification and sequencing results: M. spasma (a), C. thonglongyai (b), T. melanopogon (c), and E. spelaea (d). Giemsa-stained blood smears depicting very early gametocyte (e) with chromatin dot pointed by arrowhead, young and growing macrogametocytes (female) (f–g), growing and fully-grown macrogametocytes (h–k), and growing and fullygrown microgametocytes (l–m) of Nycteria observed in M. spasma (sample ID THBat19- 177, cytb accession no. MT136163). All parasite images were taken at the same magnification. Scale bar = 5 μm. (For viewing the images in color, the reader is referred to the online version of this article). (For interpretation of the references to color in this figure legend, the reader is referred to the Web version of this article.)

Fig. 1 in First molecular investigation of haemosporidian parasites in Thai bat species

Fig. 1. Map of Thailand depicting the bat sampling sites, forested areas, and altitudes. The map was drawn using ArcGIS version 10.2. Altitudes (in meter scale) are indicated in parentheses after each sampling site and were calculated by DIVA-GIS version 7.5.0.0 using spatial data from SRTM DEM Digital Elevation Database.

Fig. 4 in First molecular investigation of haemosporidian parasites in Thai bat species

Fig. 4. Images of bats with Polychromophilus positive infection based on either microscopic examination or cytb gene PCR amplification and sequencing results: My. siligorensis (a) and T. melanopogon (b). Giemsa-stained blood smears depicting growing and fully-grown macrogametocytes of Polychromophilus murinus observed in My. siligorensis (c–e) (sample ID THBat19-211, cytb accession no. MT136168). Growing and fully-grown macrogametocytes of Polychromophilus melanipherus observed in T. melanopogon (f–h) (sample ID THBat19- 170, cytb accession no. MT136167). Fullygrown microgametocytes of P. murinus (i–k). All parasite images were taken at the same magnification. Scale bar = 5 μm. (For viewing the images in color, the reader is referred to the online version of this article). (For interpretation of the references to color in this figure legend, the reader is referred to the Web version of this article.)