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FIG. 2. — Oocyste d in Description d'une nouvelle espèce d'Eimeria (Coccidia, Eimeridea) chez le lapin de garenne Oryctolagus cuniculus en France

FIG. 2. — Oocyste d' Eimeria roobroucki n. sp. (holotype), n° d'enregistrement: P301LV. Échelle: 10 µm.

Рис. 2. Ооцисты кокциΔий роΔа Eimeria, Isospora, Octosporella моΔифицированные, световая микроскопия (размеры увеΛичены в 400 раз, 1 ΔеΛение равно 10 мкм). A — ооцисты Eimeria изоΛированные из Testudo graeca из Апшеронской попуΛяции; B — ооцисты Isospora изоΛированные из Testudo graeca из Апшеронской попуΛяции; С — ооцисты Isospora изоΛированные из Teniadactylus caspius из Апшеронской попуΛяции; D — ооцисты Octosporella изоΛированные из Paralaudakia caucasia из Гобустанской попуΛяции. Автор: С. О. МамеΔова Fig. 2. Eimeria, Isospora, Octosporella oocysts (Magnification 1000 x, each segment corresponds to 10 μm): A — oocysts Eimeria found in Testudo graeca from Absheron population; B — oocysts Isospora found in Testudo graeca from Absheron population; C — oocysts Isospora found in Teniadactylus caspius from Absheron population; D — oocysts Octosporella found in Paralaudakia caucasia from Gobustan population. Author: S. O. Mamedova in Intestinal coccidia (Apicomplexa: Coccidia) in reptiles of Azerbaijan and anthropogenic influences on their prevalence

Рис. 2. Ооцисты кокциΔий роΔа Eimeria, Isospora, Octosporella моΔифицированные, световая микроскопия (размеры увеΛичены в 400 раз, 1 ΔеΛение равно 10 мкм). A — ооцисты Eimeria изоΛированные из Testudo graeca из Апшеронской попуΛяции; B — ооцисты Isospora изоΛированные из Testudo graeca из Апшеронской попуΛяции; С — ооцисты Isospora изоΛированные из Teniadactylus caspius из Апшеронской попуΛяции; D — ооцисты Octosporella изоΛированные из Paralaudakia caucasia из Гобустанской попуΛяции. Автор: С. О. МамеΔова Fig. 2. Eimeria, Isospora, Octosporella oocysts (Magnification 1000 x, each segment corresponds to 10 μm): A — oocysts Eimeria found in Testudo graeca from Absheron population; B — oocysts Isospora found in Testudo graeca from Absheron population; C — oocysts Isospora found in Teniadactylus caspius from Absheron population; D — oocysts Octosporella found in Paralaudakia caucasia from Gobustan population. Author: S. O. Mamedova

Рис. 1. Ооцисты криптоспориΔий моΔифицированные, световая микроскопия по метоΔу ЦиΛя — НиΛьсена (Henriksen, Pohlenz 1981) (размеры увеΛичены в 1000 раз, 1 ΔеΛение равно 10 мкм): A — ооцисты изоΛированные из Testudo graeca из Апшеронской попуΛяции; B — ооцисты изоΛированные из Paralaudakia caucasia из Гобустанской попуΛяции; С — ооцисты изоΛированные из Eremias arguta; D — ооцисты изоΛированные из Natrix tessellata из Апшеронской попуΛяции. Автор: С. О. МамеΔова Fig. 1. Cryptosporidium oocysts stained with carbol-fucsin (Henriksen, Pohlenz 1981) (Magnification 1000 x, each segment corresponds to 10 μm): A — oocysts found in Testudo graeca from Absheron population; B — oocysts found in Paralaudakia caucasia from Gobustan population; C — oocysts found in Eremias argute; D — oocysts found in Natrix tessellata from Absheron population. Author: S. O. Mamedova in Intestinal coccidia (Apicomplexa: Coccidia) in reptiles of Azerbaijan and anthropogenic influences on their prevalence

Рис. 1. Ооцисты криптоспориΔий моΔифицированные, световая микроскопия по метоΔу ЦиΛя — НиΛьсена (Henriksen, Pohlenz 1981) (размеры увеΛичены в 1000 раз, 1 ΔеΛение равно 10 мкм): A — ооцисты изоΛированные из Testudo graeca из Апшеронской попуΛяции; B — ооцисты изоΛированные из Paralaudakia caucasia из Гобустанской попуΛяции; С — ооцисты изоΛированные из Eremias arguta; D — ооцисты изоΛированные из Natrix tessellata из Апшеронской попуΛяции. Автор: С. О. МамеΔова Fig. 1. Cryptosporidium oocysts stained with carbol-fucsin (Henriksen, Pohlenz 1981) (Magnification 1000 x, each segment corresponds to 10 μm): A — oocysts found in Testudo graeca from Absheron population; B — oocysts found in Paralaudakia caucasia from Gobustan population; C — oocysts found in Eremias argute; D — oocysts found in Natrix tessellata from Absheron population. Author: S. O. Mamedova

Fig. 4 in Nematode-coccidia parasite co-infections in African buffalo: Epidemiology and associations with host condition and pregnancy

Fig. 4. Predicted mean and standard error body condition scores show associations with infection presence and season, with co-infected buffalo in much lower condition in the early wet season (Table S2). Coccidia infection status is represented with C– and C+; nematode infection status is represented with N– and N+.

Fig. 5 in Nematode-coccidia parasite co-infections in African buffalo: Epidemiology and associations with host condition and pregnancy

Fig. 5. Season and co-infection differences in nematode aggregation. (a) Aggregation patterns in calves (b) and non-calves. (c) In non-calves, the distribution of nematode parasites in the late wet season shows that k is not significantly different in coccidia positive vs. negative buffalo. (d) In the early wet season coccidia positive buffalo have a truncated distribution, resulting in significantly reduced aggregation. Arrows indicate nematode intensity values in the tail of the distribution of coccidia negative buffalo. Coccidia infection status is represented with C– and C+.

Fig. 3 in Nematode-coccidia parasite co-infections in African buffalo: Epidemiology and associations with host condition and pregnancy

Fig. 3. Patterns of parasite egg/oocyst counts with co-infection for (a) nematodes and (b) coccidia. (c), the mean nematode intensity in calves is higher in early wet season than in the late wet season independent of co-infection with coccidia. (d) Co-infection with coccidia alters the seasonal patterns of nematode intensity in non-calf buffalo (>1 year, juvenile through senescent). Calf vs. non-calf division is based on model paramters (Table 1).

Fig. 2 in Nematode-coccidia parasite co-infections in African buffalo: Epidemiology and associations with host condition and pregnancy

Fig. 2. Age specific patterns of parasite prevalence with co-infection. (a) Prevalence of nematodes is higher in buffalo co-infected with coccidia (C+) compared to coccidia negative buffalo (C–) in all age categories (N = 33, 318, 166, 272, 162 for calf, juvenile, subadult, adult and senescent C– buffalo; N = 58, 237, 55, 54, 20 for C+ buffalo). (b) Prevalence of coccidia is higher in buffalo co-infected with nematodes (N+) compared to nematode negative buffalo (N–) in calf, juvenile, subadult, and senescent buffalo but not adult buffalo (N = 13, 107, 92, 144, 56 for calf, juvenile, subadult, adult and senescent N– buffalo; N = 38, 448, 129, 208, 100 for N+ buffalo).

Fig. 1 in Nematode-coccidia parasite co-infections in African buffalo: Epidemiology and associations with host condition and pregnancy

Fig. 1. Age, sex and seasonal patterns of infection. Both parasites had the highest (a) prevalence (sample size for calf, juvenile, subadult, adult, and senescent respectively: N = 91, 555, 221, 326, 182) and (b) mean intensity in calves and juveniles (nematode N = 78, 448, 129, 208, 100; coccidia N = 58, 237, 55, 60, 14). (c) Males had lower estimated nematode prevalence and (d) higher estimated coccidia intensity compared to female buffalo. (e) The estimated nematode prevalence, coccidia prevalence, and (f) mean coccidia intensity were all increased in the early wet season compared to the late wet season. ‡Indicates significant differences at p <0.05.

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Figure 3 in An ultrastructural study on the merogonic stages of Goussia senegalensis (Faye, 1988) Diouf and Toguebaye, 1993 (Apicomplexa, Coccidia) from the liver of Pagellus bellottii (Pisces, Teleostei)*

Figure 3. Meront showing the limiting membranes of merozoites (Lm). Er = endoplasmic reticulum, Hc = host cell cytoplasm, HcN = host cell nucleus, I = invagination, Mi = mitochondrion, and N = nucleus. Scale: 2.3 µm.

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Figure 2. Advanced meront showing a in An ultrastructural study on the merogonic stages of Goussia senegalensis (Faye, 1988) Diouf and Toguebaye, 1993 (Apicomplexa, Coccidia) from the liver of Pagellus bellottii (Pisces, Teleostei)*

Figure 2. Advanced meront showing a nucleus (N). Db = dense body, Er = endoplasmic reticulum, and HcN = host cell nucleus. Scale: 2.6 µm.

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Fig. 4 in Detection and quantification of house mouse Eimeria at the species level - Challenges and solutions for the assessment of coccidia in wildlife

Fig. 4. qPCR detection of intracellular stages of Eimeria in cecum and ileum from Mus musculus. -Delta Ct value (CtEimeria - CtMouse) from each tissue for 164 mice are plotted on the graph. The dotted line indicate the threshold of −5, values above the line are considered positive for the corresponding tissue. Circles represent negative samples, triangles indicate samples with Eimeria species identification and colors correspond to the Eimeria species identified in those samples. (For interpretation of the references to color in this figure legend, the reader is referred to the Web version of this article.)

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Fig. 1 in Detection and quantification of house mouse Eimeria at the species level - Challenges and solutions for the assessment of coccidia in wildlife

Fig. 1. Geographical localization of house mice (Mus musculus) collected for this study and comparison of diagnostic methods for Eimeria. A) Localization from the 378 mice included in the present study, colors indicate the Eimeria species identified for each. B) Venn diagram showing the overlap between detection methods and successful genotyping identification of the isolates. (For interpretation of the references to color in this figure legend, the reader is referred to the Web version of this article.)

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Fig. 3 in Detection and quantification of house mouse Eimeria at the species level - Challenges and solutions for the assessment of coccidia in wildlife

Fig. 3. Morphological and morphometrical characteristics of Eimeria oocyst isolated from Mus musculus. a) Photomicrographs at 1000x amplification of Eimeria oocyst from the three species isolated from Mus musculus (red = E. falciformis; green = E. ferrisi and yellow = E. vermiformis). Length/Width ratio from b) oocyst and c) sporocysts corresponding to each species (E. falciformis n = 31; E. ferrisi n = 127 and E. vermiformis n = 35). Mean ± 95% Confidence Interval is plotted. * Represent significant difference (Tukey HSD, p <0.05). (For interpretation of the references to color in this figure legend, the reader is referred to the Web version of this article.)

Fig. 4 in First record of an infection by tissue cyst-forming coccidia in wild vizcachas (Lagostomus maximus, Rodentia) of Argentina

Fig. 4. Morphological structure of the cyst wall. Cortical brain photomicrograph shows a thin cyst wall (A). A detailed view shows that the cyst wall is made up of host-nucleated cells indicated by black arrowheads (B). In contrast, tissue cysts seen in organs other than the nervous system show a thicker cystic wall. Such is the case for those tissue cysts observed in the mammary (C–D), adrenal (E) and spleen (F) glands. The most notorious differential feature is the thick collagen outer layer in cysts outside the nervous system (white arrowheads). Bz: bradyzoites. A–B: Giemsa staining; C–D: Masson's trichrome staining. E–F: hematoxilin-eosin staining.

Fig. 2 in First record of an infection by tissue cyst-forming coccidia in wild vizcachas (Lagostomus maximus, Rodentia) of Argentina

Fig. 2. Cysts localized in adult brain and in an embryonic pituitary gland of plains vizcachas. Representative photomicrographs of histological sections of adult brains of infected vizcachas. Black arrowhead indicates a solitary tissue cyst in the striatum (A). A detailed view shows signs of inflammatory cells infiltration (arrow) (B). Silver impregnation staining allows visualization of the wall of a cortical tissue cyst (white arrowhead)(C). Embryonic pituitary exhibited multiple of cysts per section (black arrowheads) (D). Arrows indicate the radial arrangement of bradyzoites in the cyst wall in a detailed view of the embryo pituitary (E). A–B: Giemsa staining. C: silver impregnation technique. D–E: hematoxilin-eosin staining.

Fig. 1 in First record of an infection by tissue cyst-forming coccidia in wild vizcachas (Lagostomus maximus, Rodentia) of Argentina

Fig. 1. Tissue cysts in various organs of plains vizcachas. Representative photomicrographs of histological sections of ovary (A–B), uterus (C), mammary gland (D), adrenal (E), spleen (F), and suspension of cysts recovered from the superficial fascia (G), all of them collected from infected adult vizcachas. Suspension of cysts recovered from amniotic sac of an infected embryo (H). Note that while the ovaries, mammary glands, superficial fascia and fluid from amniotic sac show a high density of cysts, the rest of the organs tend to show only one or very few cysts per section. White arrowheads indicate corpora lutea and black arrowheads indicate tissue cysts. A, B, C, E and F: hematoxilin-eosin staining. D: Masson's trichrome staining.

Fig. 3 in First record of an infection by tissue cyst-forming coccidia in wild vizcachas (Lagostomus maximus, Rodentia) of Argentina

Fig. 3. Morphology of the tissue cyst recorded in plains vizcachas. Mammary gland photomicrograph illustrating a tissue cyst with a thick wall made up of a collagen fibers outer layer and the inner layer consisting of nucleated host cells enclosing thousands of spindle-shaped bradyzoites (Bz). Inset: the bradyzoites showed a conoidal (anterior) and a nonconoidal (posterior) end where the nucleus is located, and they exhibit cluster arrangements where they are ordered parallel to each other. Masson's trichrome staining.