Find research datasets worth reusing

Fig. 4 in First record and description of actinospore stages (raabeia, triactinomyxon, and aurantiactinomyxon types) of fish parasitic myxozoans from Malaysia

Fig. 4. Maximum likelihood phylogenetic tree based on small subunit 18S ribosomal DNA sequences of raabeia type and related species. The tree is rooted to the Chloromyxum cristatum and Chloromyxum fluviatile as the outgroup, with bootstrap support ≥70% indicated at nodes. GenBank accession numbers are given in parentheses followed by the name of the parasite species. Actinosporean examined in the present study are in bold. The scale bar indicates the number of expected substitutions per site.

opencc-by-4.0Dec 2023View details →

Fig. 1. Liver showing a in First report of infection with metacestode stages of Echinococcus multilocularis in a kulan (Equus hemionus kulan) from Slovakia

Fig. 1. Liver showing a single nodular encapsulated mass of about 6 cm in diameter. The mainly solid cut surface reveals several small cystic areas (arrows).

opencc-by-4.0Dec 2023View details →

Fig. 3 in First report of infection with metacestode stages of Echinococcus multilocularis in a kulan (Equus hemionus kulan) from Slovakia

Fig. 3. Phylogenetic analysis of the herein obtained sequence (Donkey_Slovakia) and exemplary sequences from GenBank (e.g., OK268248.1; OP277525.1) using the Neighbor-Joining method, bootstraps consensus tree inferred from 1000 replicates. A sequence of E. granulosus was used as an outgroup (OR166778.1).

opencc-by-4.0Dec 2023View details →

Fig. 2 in First report of infection with metacestode stages of Echinococcus multilocularis in a kulan (Equus hemionus kulan) from Slovakia

Fig. 2. Photomicrograph of the hepatic nodular lesion showing a pyogranulomatous to necrotizing inflammation with intralesional fragments of amorphous band-like eosinophil and strongly PAS-positive (inset) structures, consistent with the laminated layer of the larval stage of Echinococcus multilocularis. Hematoxylin-eosin stain and Periodic acid-Schiff stain (inset), respectively.

Fig. 7 in Quatrefoil light traps for free-swimming stages of cymothoid parasitic isopods and seasonal variation in their species compositions in the Seto Inland Sea, Japan

Fig. 7. Number of Mothocya parvostis collected at tidal levels: low tide, 1/3 tide, 2/3 tide, and high tide during the three days of sampling.

Fig. 6 in Quatrefoil light traps for free-swimming stages of cymothoid parasitic isopods and seasonal variation in their species compositions in the Seto Inland Sea, Japan

Fig. 6. Temporal variation in water temperature from October 2020 to December 2021. The gap in data is due to faulty logging equipment.

Fig. 8 in Quatrefoil light traps for free-swimming stages of cymothoid parasitic isopods and seasonal variation in their species compositions in the Seto Inland Sea, Japan

Fig. 8. Number of Mothocya parvostis collected on each sampling date (solid line) and tidal levels (broken line) from November 15 (new moon) to December 15 (new moon).

Fig. 5 in Quatrefoil light traps for free-swimming stages of cymothoid parasitic isopods and seasonal variation in their species compositions in the Seto Inland Sea, Japan

Fig. 5. Number of cymothoid juveniles collected in each month from October 2020 to December 2021. Dot bars (red) indicate Mothocya parvostis and diagonal right pattern bars (blue) indicate Ceratothoa verrucosa. (For interpretation of the references to colour in this figure legend, the reader is referred to the Web version of this article.)

Fig. 4 in Quatrefoil light traps for free-swimming stages of cymothoid parasitic isopods and seasonal variation in their species compositions in the Seto Inland Sea, Japan

Fig. 4. Number of cymothoid mancae collected in each month from October 2020 to December 2021. Dot bars (red) indicate Mothocya parvostis, diagonal right pattern bars (blue) indicate Ceratothoa verrucosa, diagonal left pattern bars (green) indicate Ceratothoa carinata. (For interpretation of the references to colour in this figure legend, the reader is referred to the Web version of this article.)

Fig. 3 in Quatrefoil light traps for free-swimming stages of cymothoid parasitic isopods and seasonal variation in their species compositions in the Seto Inland Sea, Japan

Fig. 3. Dorsal views of cymothoid free-swimming stages collected by the light trap. (a) and (d): Mothocya parvostis, (b) and (e): Ceratothoa verrucosa, (c): Ceratothoa carinata. (a)–(c): mancae, (d) and (e): juveniles. Scale bars indicate (a)–(c): 1 mm, (d) and (e): 3 mm.

Fig. 2 in Quatrefoil light traps for free-swimming stages of cymothoid parasitic isopods and seasonal variation in their species compositions in the Seto Inland Sea, Japan

Fig. 2. Map showing location of the Seto Inland Sea and sampling site, where light trap sampling was performed.

Fig. 1 in Quatrefoil light traps for free-swimming stages of cymothoid parasitic isopods and seasonal variation in their species compositions in the Seto Inland Sea, Japan

Fig. 1. The quatrefoil light trap using in this study. (a): front view, (b): bottom view without net, (c): Light traps in use underwater. A: 15 W LED fishing light, B: Net to collect organisms in trap (0.5 mm mesh).

Fig. 6 in New brachiopods from the Lower-Middle Ordovician (Billingen-Volkhov stages) of the East Baltic

Fig. 6. Bivariate plots of 51 measured specimens of Leoniorthis robusta. Solid rhombi represent specimens from Putilovo mud mound, open squares represent specimens from Popovka River, triangles represent specimens from Suhkrumägi road cut.

Fig. 5 in New brachiopods from the Lower-Middle Ordovician (Billingen-Volkhov stages) of the East Baltic

Fig. 5. Leoniorthis robusta gen. et. sp. nov. A–F. Specimens from Putilovo mud mound. G–J. Specimens from Popovka River. A. Holotype, PMU In 110, dorsal exterior (A1) and interior (A2). B. PMU In 111, ventral exterior. C. PMU In 145, ventral interior. D. PMU In 144, ventral interior. E. PMU In 120, dorsal interior (E1); enlargement of cardinal region (E2). F. PMU In 153, oblique view of hingeline and dorsal valve of conjoined specimen. G. PMU In 121, dorsal exterior. H. PMU In 122, dorsal interior. I. PMU In 125, ventral exterior. J. PMU In 123, ventral interior. Scale bars 1 mm.

Fig. 2 in New brachiopods from the Lower-Middle Ordovician (Billingen-Volkhov stages) of the East Baltic

Fig. 2. The standard section in Putilovo Quarry compared with the mud mound section (after Tolmacheva, Fedorov, and Egerquist in press). Levels with occurrences of Neumania paucicostata are marked with squares; levels with Leoniorthis robusta are marked with circles.

Fig. 4 in New brachiopods from the Lower-Middle Ordovician (Billingen-Volkhov stages) of the East Baltic

Fig. 4. Neumania paucicostata sp. nov. Specimens from Putilovo Quarry. A. Holotype PMU In 108; ventral valve interior (A1), enlargement of the spondylium showing the muscle attachment area (A2). B. PMU In 113, oblique internal view of ventral valve showing the U−shaped spondylium. Note that this specimen does not possess rounded muscle scars as in the holotype. C. PMU In 112, enlargement of spondylium on broken ventral valve, slightly oblique view showing theroughmuscleattachmentarea. D.PMUIn109,ventralviewofconjoinedspecimen. E.ExteriorofspecimenPMUIn228indifferentviews. F.PMUIn107, dorsal interior and exterior. G. PMU In 224; oblique posterior view of cardinalia (G1), dorsal valve interior (G2). Scale bars 1 mm.

Fig. 3. Neumania paucicostata. A. Dorsal valve interior showing the anteriorlylobedmusclescars,drawnfromspecimenPMUIn107. B in New brachiopods from the Lower-Middle Ordovician (Billingen-Volkhov stages) of the East Baltic

Fig. 3. Neumania paucicostata. A. Dorsal valve interior showing the anteriorlylobedmusclescars,drawnfromspecimenPMUIn107. B.Lateralview of ventral valve drawn from specimen PMU In 108 (holotype).

Fig. 3 in Location and elimination of Anisakis simplex third stage larvae in Atlantic herring Clupea harengus L

Fig. 3. The association of nematode larvae with musculature and body cavity organs in herring (unprocessed).

Fig. 2. A in Location and elimination of Anisakis simplex third stage larvae in Atlantic herring Clupea harengus L

Fig. 2. A. The correlation between total body length of herring (unprocessed) and the number of nematode larvae in each fish. B. The correlation between the total body weight of herring (unprocessed) and the number of nematode larvae in each fish.

Fig. 1 in Location and elimination of Anisakis simplex third stage larvae in Atlantic herring Clupea harengus L

Fig. 1. Left side of a herring showing division of the musculature into epaxial, hypaxial and caudal segments. The lower part of the hypaxial part (processed hypaxial) is removed during industrial processing. The right side is segmented similarly. Following processing the right and left musculature segments are still kept connected by dorsal connective tissue.