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Figs. 1–10 in Molecular phylogeny of Acerentomidae (Protura), with description of Acerentuloides bernardi sp. nov. from North America

Figs. 1–10. Acerentuloides bernardi sp. nov. 1. Head, right side; 2. pseudoculus with seta l3; 3. cephalic seta sd5; 4. maxillary palpus; 5. labial palpus; 6. maxillary gland; 7. comb; 8. female squama genitalis; 9. foretarsus, exterior view; 10. foretarsus, interior view. Arrows indicate pores (cp = clypeal pore, fr = frontal pore). Scale bars: 20 µm.

Figs. 24–34 in Molecular phylogeny of Acerentomidae (Protura), with description of Acerentuloides bernardi sp. nov. from North America

Figs. 24–34. Acerentuloides bernardi sp. nov. scanning electron microscopy photographs. 24. Habitus; 25. labial palp (apical tuf with 4 setae broken; s = labial sensillum); 26. foretarsus, interior view: sensilla t1, t3 and a'; 27. modified seta P2a on mesonotum; 28. modified seta A2 on prosternum; 29. modified seta M2 on prosternum; 30. modified seta A5 on tergite I and P1a on sternite I; 31. abdominal leg on segment III (sa = subapical, am = apical medial, al = apical lateral setae; apical lateral seta is broken); 32. hind margin of sternite VI; 33. hind margin of sternite VII; 34. sternites VIII–XII. Scale bars = 300 µm (Fig. 24), 4 µm (Figs. 25 and 27–29), 10 µm (Fig. 31), and 20 µm (Figs. 26, 30, and 32–34).

Figs. 35–40 in Molecular phylogeny of Acerentomidae (Protura), with description of Acerentuloides bernardi sp. nov. from North America

Figs. 35–40. Acerentulus confinis (Berlese, 1908), American specimen scanning electron microscopy photographs. 35. Habitus; 36. pseudoculus and cephalic setae sd4 and l3; 37. foretarsus, exterior view: sensilla t1, t3 and a'; 38. labial palpi with apical tuf of setae and basal sensillum (s); 39. sternite VIII; 40. modified seta P4 on metanotum. Scale bars = 300 µm (Fig. 35), 20 µm (Figs. 36 and 38), and 40 µm (Figs. 37, 39, and 40).

Figs. 16–23 in Molecular phylogeny of Acerentomidae (Protura), with description of Acerentuloides bernardi sp. nov. from North America

Figs. 16–23. Acerentuloides bernardi sp. nov. 16. Tergite I, right side; 17. tergite VI, right part; 18. tergite VII, right part; 19. tergite VIII; 20. sternite II; 21. abdominal leg of sternite II; 22. sternite VI; 23. sternites VII–IX. Arrows indicate pores (psm = tergal posterosubmedial, psl = tergal posterosublateral, spsm = sternal posterosubmedial, spm = sternal posteromedial). Scale bars: 20 µm.

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Fig. 1 in Pest and beneficial arthropods in a 'Tifon 85' bermudagrass field in north central Florida

Fig. 1. Mean number of major pest arthropods collected each sampling day (9 sweep net samples per sub-plot, 3 replicates) on a north central Florida farm in 'Tifon 85' bermudagrass grown for 35 d afer being cut to a stubble height of 8 or 15 cm. The sampling days were 5, 13, 18, and 27 Aug; 3, 9, 16, 23, and 30 Sep; and 7, 14, 22, and 29 Oct 2008. The grass was cut on 4 Sep and 9 Oct, between sample days 5 and 6 and sample days 10 and 11, respectively (bars below the x-axes).

Fig. 5 in Heartworm and seal louse: Trends in prevalence, characterisation of impact and transmission pathways in a unique parasite assembly on seals in the North and Baltic Sea

Fig. 5. Scanning electron microscopic images of E. horridus attached to seal skin and fur. A: E. horridus utilizing a hair follicle of harbour seal skin. B: E. horridus attached to seal hair with the head pointing towards seal skin. C: E. horridus with six claws attached to seal fur. D: Close up of an unattached claw of E. horridus. Asterisks positioned on nits of E. horridus. Sale bars: A 200 μm, B 400 μm, C 400 μm, D 100 μm.

Fig. 4 in Heartworm and seal louse: Trends in prevalence, characterisation of impact and transmission pathways in a unique parasite assembly on seals in the North and Baltic Sea

Fig. 4. Histological sections and staining of E. horridus revealing filarial stages in E. horridus. A: Filarial stages (arrowheads) in the pharynx. bar = 20 μm. B: Filarial stage (arrow) in the mouth region. bar = 40 μm. C: Filarial stage (arrowhead) in the intestine (in) surrounded by erythrocytes (e). bar = 15 μm. D: Filarial stage in the haemocoel (hc) of the abdomen of E. horridus (square). E: Close up of filarial stage. mp = mouthparts, mo = mouth, cu = cuticula. A–C: Haematoxylin - Eosin stain, D: Giemsa stain.

Fig. 3. A in Heartworm and seal louse: Trends in prevalence, characterisation of impact and transmission pathways in a unique parasite assembly on seals in the North and Baltic Sea

Fig. 3. A: Prevalence of A. spirocauda and E. horridus in harbour seals in the North and Baltic Sea from 1996 to 2021, data from 1996 to 2013 according to Lehnert et al. (2016). B: Prevalence of A. spirocauda and E. horridus in harbour seals during the seasons in the North and Baltic Sea from 2014 to 2021.

Fig. 2 in Heartworm and seal louse: Trends in prevalence, characterisation of impact and transmission pathways in a unique parasite assembly on seals in the North and Baltic Sea

Fig. 2. Sampling routine of E. horridus infected seal skin for histological and bacteriological examinations. A: Mild E. horridus infection of a harbour seal yearling, asterisk pointing at E. horridus. B: Close up of E. horridus C: Removing of E. horridus D: Cutting and removing of the infected skin with a sterile forceps for further investigations. Scale bars: A-D 1 cm.

Fig. 1 in Heartworm and seal louse: Trends in prevalence, characterisation of impact and transmission pathways in a unique parasite assembly on seals in the North and Baltic Sea

Fig. 1. Levels of infection with E. horridus in P. vitulina. A: Mild E. horridus infection of a harbour seal yearling, asterisk pointing at E. horridus B: Close up of E. horridus in the head area of a harbour seal C: Severe E. horridus infection of a harbour seal D: Close up of severe E. horridus infection. Scale bars: A-D 1 cm.

Fig. 6. Ectoparasite-host interactions for A in Ectoparasitism on deep-sea fishes in the western North Atlantic: In situ observations from ROV surveys

Fig. 6. Ectoparasite-host interactions for A) Antimora rostrata, B) Nezumia bairdii, and C) Synaphobranchus spp. across the region for which the presence or absence of ectoparasites could be confirmed. Circle Size = Number of confirmed ectoparasite-host interactions 10 m —2. Square = No ectoparasites observed. Colors denote habitats.

Fig. 2 in Ectoparasitism on deep-sea fishes in the western North Atlantic: In situ observations from ROV surveys

Fig. 2. Example images of ectoparasites infecting various host species. A) Antimora rostrata with a lernaeopodid copepod (1059 m, Alvin Canyon); B) Synaphobranchus sp. with a sphyriid copepod (820 m, open slope); C) Nezumia bairdii with a sphyriid copepod parasitized by eight leeches (1035 m, Phoenix Canyon; D), Black fish, Diaphus sp. with an unknown siphonostomatid copepod (1130 m, cold seep) attached behind dorsal fin; E) N. bairdii with aegid isopod (780 m, open slope); F) Hoplostethus mediterraneus with a cymothoid isopod (744 m, Nygren Canyon); G) Amblyraja radiata with aegid isopod (1010 m, Alvin Canyon); and H) Cottunculus thomsonii with gnathiids (1210 m, Oceanographer Canyon).

Fig. 3 in Ectoparasitism on deep-sea fishes in the western North Atlantic: In situ observations from ROV surveys

Fig. 3. Percentage of ectoparasites infecting three common demersal fishes. Unconfirmed observations are not included.

Fig. 1 in Ectoparasitism on deep-sea fishes in the western North Atlantic: In situ observations from ROV surveys

Fig. 1. Locations of 43 ROV dives conducted along the northeastern U.S. continental margin and New England Seamount Chain. Circle Size = Number of ectoparasite-host interactions per dive. Square = No ectoparasites observed. Colors denote habitats.

Fig. 5 in Ectoparasitism on deep-sea fishes in the western North Atlantic: In situ observations from ROV surveys

Fig. 5. Mean abundance (±SE) of A) Antimora rostrata, B) Nezumia bairdii, and C) Synaphobranchus spp. per habitat (black bars). Mean abundance (±SE) of ectoparasitehost interactions (grey bars) is also included. n = number of dives during which the species was present.

Fig. 4 in Ectoparasitism on deep-sea fishes in the western North Atlantic: In situ observations from ROV surveys

Fig. 4. Mean abundance (±SE) of A) Antimora rostrata, B) Nezumia bairdii, and C) Synaphobranchus spp. per depth zone (black bars). Mean abundance (±SE) of ectoparasite-host interactions (grey bars) also included. n = number of dives during which the species was present.

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Рис. 6. МоΔеΛирование экоΛогических ниш коΛораΔского жука ΔΛя ΔаΛьневосточного, европейского и североамериканского ареаΛов метоΔом метрического Δвухмерного шкаΛирования с применением коэффициента Жаккара Fig. 6. Models of ecological niches of the Colorado potato beetle for the Far Eastern, European, and North-American habitats (metric multidimensional scaling, Jaccard index) in Comparative characterization of the ecology of native (Henosepilachna vigintioctomaculata) and invasive (Leptinoatrsa decemlineata) species under the conditions of the monsoon climate in the southern part of the Russian Far East

Рис. 6. МоΔеΛирование экоΛогических ниш коΛораΔского жука ΔΛя ΔаΛьневосточного, европейского и североамериканского ареаΛов метоΔом метрического Δвухмерного шкаΛирования с применением коэффициента Жаккара Fig. 6. Models of ecological niches of the Colorado potato beetle for the Far Eastern, European, and North-American habitats (metric multidimensional scaling, Jaccard index)

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Fig. 5 in Prevalence and molecular characterisation of Acanthocephala in pinnipedia of the North and Baltic Seas

Fig. 5. Maximum likelihood tree based on COI sequences using the Jones-TaylorThornton (JTT) model. The log likelihood is −2479.70. The percentage of trees based on 1000 bootstrap replicates in which the associated taxa clustered together is shown next to the branches. GenBank accession numbers of analysed amino acid sequences are listed in Table 2.

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Fig. 1 in Prevalence and molecular characterisation of Acanthocephala in pinnipedia of the North and Baltic Seas

Fig. 1. Prevalence of acanthocephalan infections in harbour and grey seals from the German North and Baltic Seas between 1996 and 2012. Notations indicate the Acanthocephala positive and total number of examined Phoca vitulina (Pv) and Halichoerus grypus (Hg). Connecting lines indicate statistically significant differences between annual prevalences after Holm–Bonferroni correction (P ≤ 0.001).