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Fig. 1 in Is parasite load dependent on host aggregation size? The case of the greater mouse-eared bat Myotis myotis (Mammalia: Chiroptera) and its parasitic mite Spinturnix myoti (Acari: Gamasida)

Fig. 1 Distribution of greater mouse-eared bat (Myotis myotis) maternity aggregations (grey circles) and single founding individuals (black dots) in the Beskids (Carpathian Mountains, Poland). Data pooled from Kozakiewicz (2003), Szkudlarek et al. (2008), and our data. For the investigated maternity colonies, forested areas within a 10-km radius is shown

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Fig. 3 in Is parasite load dependent on host aggregation size? The case of the greater mouse-eared bat Myotis myotis (Mammalia: Chiroptera) and its parasitic mite Spinturnix myoti (Acari: Gamasida)

Fig. 3 Proportions of protonymph (PN), deutonymph (DN), and adult (AD) stages of S. myoti mites, collected from Myotis myotis (bars), and the sex ratios of deutonymph and adult mites (circles)

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Fig. 2 in Is parasite load dependent on host aggregation size? The case of the greater mouse-eared bat Myotis myotis (Mammalia: Chiroptera) and its parasitic mite Spinturnix myoti (Acari: Gamasida)

Fig. 2 Micrograph of the adult female Spinturnix myoti, dorsal view. Scanning electron microscopy image, original magnification ×40

Fig. 3 in Understanding growth relationships of African cymothoid fish parasitic isopods using specimens from museum and field collections

Fig. 3. The relationship between cymothoid size and host size for female and male parasites by parasite species.

Fig. 3 in Molecular insights into the identification and phylogenetics of the cosmopolitan marine fish blood parasite, Haemogregarina bigemina (Adeleorina: Haemogregarinidae)

Fig. 3. Phylogenetic identification of Haemogregarina bigemina from the UK based on 18S rDNA sequences. (a) Maximum parsimony and (b) Maximum likelihood reconstructions revealing the unique position of UK H. bigemina samples outside of the adeleorine groups. For both phylogenies nodal support was calculated using 1000 bootstrap replicates with only values> 50% presented.

Fig. 1 in Molecular insights into the identification and phylogenetics of the cosmopolitan marine fish blood parasite, Haemogregarina bigemina (Adeleorina: Haemogregarinidae)

Fig. 1. Photograph of the fish host Lipophrys pholis, one of the type hosts of Haemogregarina bigemina, screened in this study.

Fig. 2 in Understanding growth relationships of African cymothoid fish parasitic isopods using specimens from museum and field collections

Fig. 2. The total number, mean number and standard deviation of parasites collected from the South African Institute for Aquatic Biodiversity and fieldwork, respectively. J = juvenile, M = male, F = female. Attachment type indicated as: B = buccal, T = tongue, P = palate, G = gill.

Fig. 2 in Molecular insights into the identification and phylogenetics of the cosmopolitan marine fish blood parasite, Haemogregarina bigemina (Adeleorina: Haemogregarinidae)

Fig. 2. Stages of Haemogregarina bigemina from Giemsa-stained blood films of Lipophrys pholis from the UK. (a) trophozoite, (b) meront, (c–e) dividing meronts, and (f) paired gamonts. Scale bar = 10 μm.

Fig. 1 in Understanding growth relationships of African cymothoid fish parasitic isopods using specimens from museum and field collections

Fig. 1. Isopods preserved along with their fish hosts from the South African Institute for Aquatic Biodiversity (SAIAB). A. Ceratothoa famosa Hadfield, Bruce & Smit, 2014 in the mouth of Diplodus capensis (Smith, 1844); B. Mothocya affinis Hadfield, Bruce & Smit, 2015 in the gills of Hyporhamphus affinis (Günther, 1866); C. Cymothoa sodwana Hadfield, Bruce & Smit, 2013 in the mouth of Trachinotus botla (Shaw, 1803).

Fig. 4 in Understanding growth relationships of African cymothoid fish parasitic isopods using specimens from museum and field collections

Fig. 4. The relationship between cymothoid size and host size for juvenile parasites by parasite species.

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. 1 in Phylogeny of Hepatocystis parasites of Australian flying foxes reveals distinct parasite clade

Fig. 1. Concatenated analysis. Bayesian analysis of concatenated alignment of two mitochondrial (cytb, cox1), one apicoplast (clpC), and two nuclear genes (ef2, PAT) rooted with Leucocytozoon species from birds. Posterior probabilities are given. (A) Clade of Hepatocystis presents the sister clade to mammalian Plasmodium (Plasmodium) and Plasmodium (Vinckeia) species. The parasite sequences of the study from Australian Pteropus species, form one distinct clade (together with three sequences from Asian Pteropus species) (highlighted in yellow). The sister clade contains all sequences from primate Hepatocystis (highlighted in blue), the African bat Hepatocystis parasites (highlighted in red) and sequences of Hepatocystis from Asian flying foxes of the genus Cynopterus and Hipposideros. (B) section from (A), uncollapsed Hepatocystis clades. Sequences of the study are highlighted in bold. Australasian Hepatocystis sequences from Pteropus hosts fall in two subclades and no host species specificity is apparent as the sequences from all three-host species group in two main clades.(For interpretation of the references to colour in this figure legend, the reader is referred to the web version of this article.)

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Fig. 2 in Phylogeny of Hepatocystis parasites of Australian flying foxes reveals distinct parasite clade

Fig. 2. Gametocyte blood stages of haemosporidian parasites of Australian bats. Giemsa-stained thin blood smears were investigated using oil immersion with a light microscope at a magnification of 1,000×. A) ex P. conspicillatus (P_conspicillatus_L2, A1 = macro-, A2 = microgametocyte), B) ex P. conspicillatus (P_conspicillatus_L15, B1 = macro-, B2 = microgametocyte), C) ex P. conspicillatus (P_conspicillatus_L20, C1+2 = early gametocyte stages, C3 = macro-, C 4 = microgametocyte), D) ex P. scapulatus (P_scapulatus_A4, D 1+2 = micro-, D 3+4 macrogametocytes), E) ex P. scapulatus (P_scapulatus_A3, E1+2 = macrogametocytes, E3+4 = unusual microgametocytes) F) ex P. alecto (P_alecto_L8, F1+2 = microgametocytes G) ex P. alecto (P_alecto_L5, G1 = macrogametocyte, G2 = microgametocyte), ''ex'' denotes that parasites were isolated from the respective host species. Bar = 5 μm.

Fig. 1 in Metazoan parasites of California sea lions (Zalophus californianus): A new data and review

Fig. 1. Microphotographs of the metazoan parasites of California sea lions Zalophus califronianus. A, B – Apophallus zalophi, intestine (Digenea), C – Zalophotrema hepaticum, liver (Digenea), D, G – Contracaecum ogmorhini s. l., stomach (Nematoda), E, H – Pseudoterranova decipiens s. l., stomach (Nematoda), F – Anisakis simplex s. l., stomach (Nematoda), I, S – Diphyllobothrium sp., intestine (Cestoda), J, K – Orthohalarchne attenuata, nasal cavity (Acarina), L – Orthohalarchne diminuata (Acarina), M, R – Anophryocephalus sp., intestine (Cestoda), N, O – Corynosoma obtuscens, intestine (Acanthocephala), P, Q – Profilicollis altmani, intestine (Acanthocephala). A, C, F–I, K, M, N, P – under dissecting scope. B, Q – at light microscope. D, E, O, R, S, – at scanning electron microscope. J – in situ.

Fig. 2 in Molecular confirmation of Hymenolepis hibernia in field mice (Apodemus sylvaticus) from St Kilda has potential to resolve a host-parasite relationship

Fig. 2. The Sanger sequence of 385 bp of mt-Cox-1 fragments of cyclophyllidean DNA generated from eggs recovered from mouse 9 and faeces X (6. H-HA25), aligned with 5 corresponding H. hibernia sequences published in the NCBI Genbank database. There are 12 intraspecific SNPs at positions 5, 60, 78, 84, 90, 156, 216, 219, 252, 262, 291 and 318. GeneBank submission ID: 2151861.

Fig. 1 in Molecular confirmation of Hymenolepis hibernia in field mice (Apodemus sylvaticus) from St Kilda has potential to resolve a host-parasite relationship

Fig. 1. Examples of cyclophyllidean tapeworms and eggs recovered from Apodemus sylvaticus viscera and faeces. A: Unarmed scolex of an intact tapeworm - mouse 5. B: Anoplocephalid tapeworm egg - mouse 8. C: Hymenolepidid egg cropped without changing dimensions from the same image as B, for comparison - mouse 8. D: Anoplocephalid tapeworm egg - mouse X (faeces from a cleat). E: Anoplocephalid onchosphere released after squashing an egg under a cover slip - mouse X (faeces from a cleat). F: Tapeworm from mouse 8 (scolex not intact). G: hymenolepidid tapeworm egg - mouse X (faeces from a cleat). H: Hexacanth onchosphere surrounded by an onchospheral membrane and inner zone of the embryophore, which has swollen, having been released from the egg shell by squashing under a cover slip - mouse 9. I: Hymenolepidid tapeworm egg - mouse 7. J: Hexacanth onchosphere surrounded by an intact onchospheral membrane and inner and outer zones of the embryophore, being released from a cracked egg shell by squashing under a cover slip - mouse 7. K: Hymenolepidid tapeworm egg - mouse 8. L: Hexacanth onchosphere surrounded by an onchospheral membrane and inner zone of the embryophore, which has swollen, having been released from the egg shell by squashing under a cover slip - mouse 8.

Fig. 1. Urinary parasites, light microscopy. A in Urinary capillariosis in a free-ranging Marsican brown bear (Ursus arctos marsicanus)

Fig. 1. Urinary parasites, light microscopy. A) Mature capillariid egg, 40X, B) Adult female, vulvar appendix, 40X, C) Adult female, anterior extremity, 40X, D) Adult female, posterior extremity, 40X.

Fig. 2 in Metazoan parasites of California sea lions (Zalophus californianus): A new data and review

Fig. 2. Prevalence and proportion of separate species in the gastrointestinal helminth community of California sea lions (Zalophus californianus). Abbreviations of the genera: A – Anisakis, An – Andracantha, Ap – Apophallus, C – Contracaecum, Co – Corynosoma, P – Pseudoterranova, Pa – Parafilaroides, Pr – Profilicollis, Z – Zalophotrema.