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Fig. 4 in Integrated morphological and molecular characterization of the fish parasitic nematode Rhabdochona (Rhabdochona) gendrei Campana-Rouget, 1961 infecting Labeobarbus altianalis (Boulenger, 1900) in Kenya

Fig. 4. Rhabdochona (Rhabdochona) gendrei Campana-Rouget (1961) from Labeobarbus altianalis (Boulenger, 1900), scanning electron micrographs of a female. A (deirids shown by white circle), B, C – cephalic end dorsolateral and anterior views; D– lateral view of the excretory pore (as shown by white arrow); E – detail of vulva sub-ventral view; F, G – fully mature (larvated) eggs dissected out of the nematode body; H – tail tip (white arrows shows openings with a papilla); Abbreviations: a – amphids; c – submedian cephalic papilla; pt – anterior prostomal teeth.

Fig. 3 in Integrated morphological and molecular characterization of the fish parasitic nematode Rhabdochona (Rhabdochona) gendrei Campana-Rouget, 1961 infecting Labeobarbus altianalis (Boulenger, 1900) in Kenya

Fig. 3. Rhabdochona (Rhabdochona) gendrei Campana-Rouget (1961) from Labeobarbus altianalis (Boulenger, 1900), light micrographs of an adult male. A – left (longer) and right (shorter) spicules (dorsolateral view); B– left (longer) and right (shorter – showing the boat-like shape) spicules (ventrolateral view); black and white arrows indicate left and right spicules respectively.

Fig. 1. A in Integrated morphological and molecular characterization of the fish parasitic nematode Rhabdochona (Rhabdochona) gendrei Campana-Rouget, 1961 infecting Labeobarbus altianalis (Boulenger, 1900) in Kenya

Fig. 1. A map of River Nyando, in Lake Victoria Basin, showing the study locations at Koru and Ahero.

Fig. 5 in Integrated morphological and molecular characterization of the fish parasitic nematode Rhabdochona (Rhabdochona) gendrei Campana-Rouget, 1961 infecting Labeobarbus altianalis (Boulenger, 1900) in Kenya

Fig. 5. Phylograms of the 28S rRNA gene region based on the A– maximum likelihood and B – Bayesian Inference analyses. Bootstrap and posterior probability support values are presented along branch nodes. The branch length was reduced to two (//) and three (///) times the scale bar.

Fig. 2 in Integrated morphological and molecular characterization of the fish parasitic nematode Rhabdochona (Rhabdochona) gendrei Campana-Rouget, 1961 infecting Labeobarbus altianalis (Boulenger, 1900) in Kenya

Fig. 2. Rhabdochona (Rhabdochona) gendrei Campana-Rouget (1961) from Labeobarbus altianalis (Boulenger, 1900), scanning electron micrographs of the male. A – cephalic end, subapical view; B – deirid; (circled); C, D – posterior end of male, ventrolateral view (white stars and white circles indicate a pair of lateral pre- and postanal papillae, respectively); E – right spicule; F – excised right spicule, following enzymatic digestion. Abbreviations: a – amphid; 1–11, and 1–9 – pairs of sub-ventral preanal papillae (in C); 1–5 – pairs of postanal papillae (in D).

Fig. 1 in Working towards a conservation plan for fish parasites: Cyprinid parasites from the south African cape fold freshwater ecoregion as a case study

Fig. 1. Map illustrating localities where the five cyprinid hosts were collected in the Cape Fold ecoregion in the Western Cape, South Africa.

Fig. 7 in Working towards a conservation plan for fish parasites: Cyprinid parasites from the south African cape fold freshwater ecoregion as a case study

Fig. 7. Rarefaction/extrapolation curve estimating the diversity of parasites as a function of sampling effort for three of the five hosts collected in the OlifantsDoorn River System, Western Cape Province, South Africa. Shaded area represents the 95% confidence interval obtained using the bootstrap method based on 100 repetitions. Created using iNEXT Online (Chao et al., 2016).

Fig. 4 in Working towards a conservation plan for fish parasites: Cyprinid parasites from the south African cape fold freshwater ecoregion as a case study

Fig. 4. Pseudobarbus calidus (Barnard, 1938) (max. length: 125 mm) (A). Sclerites of Paradiplozoon sp. from the gills (B). Acanthocephala from the body cavity, whole specimen (C) and hooks on proboscis (top left insert). Larval Contracaecum sp. from the body cavity, anterior (D) and posterior (E) ends, lateral view. Scale bars: 100 μm (B, C, D, E).

Fig. 6 in Working towards a conservation plan for fish parasites: Cyprinid parasites from the south African cape fold freshwater ecoregion as a case study

Fig. 6. Sedercypris erubescens (Skelton, 1974) (max. length: 120 mm) (A). Larval Contracaecum sp. from the body cavity, anterior (B) and posterior (C) ends, lateral view. Scale bars: 100 μm (B, C).

Fig. 5 in Working towards a conservation plan for fish parasites: Cyprinid parasites from the south African cape fold freshwater ecoregion as a case study

Fig. 5. Pseudobarbus phlegethon (Barnard, 1938) (max. length: 65 mm) (A); Acanthogyrus sp. found from the body cavity (B). Scale bar: 500 μm.

Fig. 3 in Working towards a conservation plan for fish parasites: Cyprinid parasites from the south African cape fold freshwater ecoregion as a case study

Fig. 3. Labeobarbus seeberi (Gilchrist et Thompson, 1913) (max. length: 270 mm) (A); Myxobolus sp. (B) and Dactylogyrus sp. from the gills of L. seeberi, hamuli and marginal hooks (C), male copulatory complex (D) and vagina (E). Lateral view of Rhabdochona sp. 2 from the intestine, anterior end of female (F) and male (G), posterior end of male (H); metacercariae of Diplostomidae (I) from black cysts on skin. Scale bars: 10 μm (B); 50 μm (D, E); 100 μm (C, F, G, H, I).

Fig. 2 in Working towards a conservation plan for fish parasites: Cyprinid parasites from the south African cape fold freshwater ecoregion as a case study

Fig. 2. Cheilobarbus serra (Peters, 1864) (max. length: 350 mm) (A); adult Paradiplozoon sp. (B) and sclerites in attachment clamps (C, D) found on the gills; hamuli of Gyrodactylus sp. (E) and marginal hooks (F), and a pre-metamorphic stage of the copepod belonging to the Lernaeidae (G), both from the gills. Anterior (H) and posterior (I) ends of Rhabdochona sp. 1 (lateral view) from the intestine; (J) whole specimen of the Caryophyllidea. Scale bars: 10 μm (E, F); 100 μm (C, D, G, H, I); 500 μm (B); 1000 μm (J).

Fig. 2 in A young parasite in an old fish host: A new genus for proteocephalid tapeworms (Cestoda) of bowfin (Amia calva) (Holostei: Amiiformes), and a revised list of its cestodes

Fig. 2. Laruella perplexa (La Rue, 1911) n. comb. from Amia calva. SEM micrographs. A – frontal view of scolex. B, D – subapical view of scolex; note tear-shaped inner rim of suckers. C – apical view of scolex; note apical pit. E – capiliform filitriches on scolex (apical region anterior to suckers – indicated by letter E in Fig. 2B).

Fig. 4 in A young parasite in an old fish host: A new genus for proteocephalid tapeworms (Cestoda) of bowfin (Amia calva) (Holostei: Amiiformes), and a revised list of its cestodes

Fig. 4. Laruella perplexa (La Rue, 1911) n. comb. from Amia calva, Illinois and Mississippi (A–D) and Proteocephalus ambloplitis (Leidy, 1887) from Micropterus dolomieu (E, F). A – terminal genitalia with uterine diverticula near anterior part of proglottids (MHNG-PLAT-0063348), dorsal view; vitelline follicles are not illustrated. B, C – terminal genitalia, frontal section and ventral view of paratype (USNM 1348679). D – posterolateral end of proglottid (MHNG-PLAT-0063348); note band of posterior (median) vitelline follicles bent inwards. E – proximal part of vaginal canal, dorsal view; note numerous loops. F – cirrus-sac, dorsal view; note large, thick-waled vaginal sphincter and strongly convoluted internal sperm duct. Abbreviations: CI - cirrus; CS - cirrus sac; DOC – dorsal osmoregulatory canal; EG – egg; MG – Mehlis' gland; MVF – median vitelline follicles; OV – ovary; SR – seminal receptacle; TE – testes; UD – uterine diverticula; VA – vagina; VD – vas deferens; VF – vitelline follicles; VOC – ventral osmoregulatory canal; VS – vaginal sphincter.

Fig. 1 in A young parasite in an old fish host: A new genus for proteocephalid tapeworms (Cestoda) of bowfin (Amia calva) (Holostei: Amiiformes), and a revised list of its cestodes

Fig. 1. Laruella perplexa (La Rue, 1911) n. comb. from Amia calva (A, B, C, E, G, H) and Proteocephalus ambloplitis (Leidy, 1887) from Micropterus dolomieu (D, F). A, D – anterior part of body with first proglottids. B, C, E, F – scolex, frontal view; B – holotype (USNM 1347286); note narrow sphincters around sucker opening in C, E). G – cross section through scolex; note sphincters on margin of sucker openings; paratype (USNM 1348679). Abbreviations: AO - apical organ; EG – egg; ILM – inner longitudinal musculature; OC – osmoregulatory canals; SS – sucker sphincter; SU – sucker; TE - testes; TG - tegument; UT – uterus; VF – vitelline follicles.

Fig. 3 in A young parasite in an old fish host: A new genus for proteocephalid tapeworms (Cestoda) of bowfin (Amia calva) (Holostei: Amiiformes), and a revised list of its cestodes

Fig. 3. Laruella perplexa (La Rue, 1911) n. comb. from Amia calva, Mississippi, USA (MHNG-PLAT-0063348). A – mature proglottid, ventral view. B – pregravid proglottid, ventral view. Abbreviations: CS - cirrus sac; DOC – dorsal osmoregulatory canal; EG – egg; GP – genital pore; MG – Mehlis' gland; MVF – median vitelline follicles; OC – oocapt; OV – ovary; SR – seminal receptacle; SU – sucker; TE – testes; UD – uterine diverticula; VA – vagina; VD – vas deferens; VF – vitelline follicles; VOC – ventral osmoregulatory canal; VS – vaginal sphincter.

Fig. 1 in First step towards understanding the specific identity of fish muscle parasites of the genus Sarcotaces (Copepoda: Philichthyidae)-New species and first molecular ID in the genus

Fig. 1. Photographs documenting copepod (Sarcotaces izawai sp. nov.) infection of fish (Mora moro); (A) Parasite-induced black-stained void in the body of host fish, (B) parasite gall in the muscles of host fish, (C) The same with myomeres removed, (D) Composite microphotograph of the male parasite, lateral view. Scale bars: A–C = 30 mm, D = 0.5 mm. Photos A–C: by Karolina Po´łtorak.

Fig. 4 in First step towards understanding the specific identity of fish muscle parasites of the genus Sarcotaces (Copepoda: Philichthyidae)-New species and first molecular ID in the genus

Fig. 4. SEM micrographs of Sarcotaces izawai sp. nov.; male; (A) Cephalon, dorsal, (B) Cephalothorax, ventral, (C) Cephalon, anterior view, (D) Antennule, dorsal, (E) Antennule, ventral, (F) Antenna, ventral.

Fig. 3 in First step towards understanding the specific identity of fish muscle parasites of the genus Sarcotaces (Copepoda: Philichthyidae)-New species and first molecular ID in the genus

Fig. 3. Line drawings of Sarcotaces izawai sp. nov.; male; (A–E) Caudal rami of other male specimens, ventral; (F) Antennule, ventral; (G) Antenna, ventral; (H) Mandible (Mdb), and maxillae (Mx), ventral; above—protuberances of supramandibular ridge; left mandible omitted), (I) First leg (right side), ventral, (J) Second leg (left side), ventral; Scale bars: A–E = 0.1 mm, F–J = 0.01 mm.

Fig. 5 in First step towards understanding the specific identity of fish muscle parasites of the genus Sarcotaces (Copepoda: Philichthyidae)-New species and first molecular ID in the genus

Fig. 5. SEM micrographs of Sarcotaces izawai sp. nov.; male; (A) Antenna, mandible and maxillule, ventral, (B) Mandibular claw and maxillule, ventral, (C) Maxillae, ventral, (D) Exopod of first thoracopod (left side), (E) Caudal rami of other male specimens, ventral, (F) Caudal ramus of another male specimens, ventral.