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Figure 1 in First record of Planchonia stentae (Brain, 1920) (Hemiptera: Coccomorpha: Asterolecaniidae) on Asclepias curassavica Linnaeus, 1753 (Gentianales: Asclepiadaceae) in Mexico, with observations on parasitic Encyrtidae
Figure 1. Adult females of Planchonia stentae. A) Adult females (arrows) on the stem of Asclepias curassavica. B) Whole body. C) Anal ring (r) with setae (s), arched plate (a) and anal plate (asterisk). D) Lateral abdominal margin with 8-shaped pores in double or triple row (arrows). E) Multilocular pore bands (arrows), with locular plates of multilocular pores with fringed edges (insert); anal ring (r). Scales: B = 0.2 mm. C = 50 µm. D = 20 µm. E = 50 µm. Insert = 6.0 µm.
FIG. 1. — Orygmatobothrium schmittii n in A new Orygmatobothrium Diesing, 1863 (Eucestoda, Tetraphyllidea) parasite of Mustelus schmitti Springer, 1939 (Carcharhiniformes, Triakidae) from the southwestern Atlantic Ocean
FIG. 1. — Orygmatobothrium schmittii n. sp.; A, scolex; B, gravid proglottid (ventral view); C, cross section throught proglottid at level of ovary; D, detail of terminal genitalia with everted cirrus (lateral view); E, egg. Abbreviations: d, dorsal; v, ventral; o, ovary; od, oviduct; dod, dorsal osmorregulatory duct; vod, ventral osmoregulatory duct; mg, mehlis glands; va, vagin; vf, vitelline follicles; u, uterus. Scale bars: A-C, 0.2 mm; B, 1 mm; D, 0.05 mm; E, 0.01 mm.
FIG. 4 in Descriptions of two new species of the genus Tachygonetria Wedl, 1862 (Nematoda, Pharyngodonidae) and redescriptions of five species parasites of Palaearctic Testudinidae
FIG. 4. — Tachygonetria pusilla Seurat, 1918; A, cephalic end, en face of female; B, cephalic end, en face of male; C, caudal extremity, ventral view of male, D-F, scanning electron micrograph (SEM); D, cephalic end of female; E, cephalic end of male; F, caudal
FIG. 7. — Tachygonetria khallaayounei n in Descriptions of two new species of the genus Tachygonetria Wedl, 1862 (Nematoda, Pharyngodonidae) and redescriptions of five species parasites of Palaearctic Testudinidae
FIG. 7. — Tachygonetria khallaayounei n. sp.; A, cephalic end, en face of female; B, cephalic end, en face of male; C, caudal extremity, ventral view of male; D-F, scanning electron micrograph (SEM); D, cephalic end of female; E, cephalic end of male; F, caudal
FIG. 3 in Descriptions of two new species of the genus Tachygonetria Wedl, 1862 (Nematoda, Pharyngodonidae) and redescriptions of five species parasites of Palaearctic Testudinidae
FIG. 3. — Tachygonetria setosa Seurat, 1918; A, cephalic end, en face of female; B, cephalic end, en face of male; C, caudal extremity, ventral view of male; D-F, scanning electron micrograph (SEM); D, cephalic end of female; E, cephalic end of male; F, caudal
FIG. 6. — Tachygonetria marocana n in Descriptions of two new species of the genus Tachygonetria Wedl, 1862 (Nematoda, Pharyngodonidae) and redescriptions of five species parasites of Palaearctic Testudinidae
FIG. 6. — Tachygonetria marocana n. sp.; A, cephalic end, en face of female; B, cephalic end, en face of male; C, caudal extremity, ventral view of male; D-F, scanning electron micrograph (SEM); D, cephalic end of female; E, cephalic end of male; F, caudal
FIG. 5 in Descriptions of two new species of the genus Tachygonetria Wedl, 1862 (Nematoda, Pharyngodonidae) and redescriptions of five species parasites of Palaearctic Testudinidae
FIG. 5. — Tachygonetria palearcticus (Petter, 1966); A, cephalic end, en face of female; B, cephalic end, en face of male; C, caudal extremity, ventral view of male; D-F, scanning electron micrograph (SEM); D, cephalic end of female; E, cephalic end of male; F, caudal extremity of male, ventral view. Abbreviation: d, crescent-shaped cuticularized plates. Scale bars: A, B, 10 μm; C, 25 μm;
FIG. 1 in Descriptions of two new species of the genus Tachygonetria Wedl, 1862 (Nematoda, Pharyngodonidae) and redescriptions of five species parasites of Palaearctic Testudinidae
FIG. 1. — Tachygonetria numidica Seurat, 1918; A, cephalic end, en face of female; B, cephalic end, en face of male; C, caudal extremity, ventral view of male; D-F, scanning electron micrograph (SEM); D, cephalic end of female; E, cephalic end of male; F, caudal extremity of male, ventral view. Abbreviations: a, cephalic arc of broad line; b, superficial opaque lips; c, subjacent trans-
Protocol for transfection by microinjection into the eggs of the parasite vector snail Biomphalaria glabrata
<p><strong>1. Egg production</strong></p> <p>Place about 30 adult snails (10 mm diameter) into a 5.5-liter water tank. Place a piece of polystyrene of (3 x 3 cm) in each tank. There is the preferred support of <em>Biomphalaria glabrata</em> for laying its eggs. The snails are fed <em>ad libitum</em> with green lettuce leaves, they can also be fed with dry spirulina to boost reproduction. Maintain water at a temperature of 25 degrees Celsius.</p> <p><strong>2. Egg collection</strong></p> <p>Gently pick up several egg layers from the polystyrene with soft holding forceps and place the eggs into a petri dish with natural mineral water (e.g. Volvic) to prevent them from drying out.</p> <p>Start sorting the eggs under the stereoscopic microscope to choose only the gastrula stage and place them into another petri dish with natural mineral water.</p> <p> </p> <p><strong>3. Preparation of the transfection solution</strong></p> <p><strong>Material:</strong></p> <p>a. <em>in vivo</em> JetPEI transfection reagent</p> <p>b. 10% glucose solution</p> <p>c. 5% glucose solution</p> <p>d. Plasmids (dCas9-SunTag-BFP and scFv-DNMT3A-GFP)</p> <p>e. 0.2 ml microtubes</p> <p>f. P10 and P200 pipettes</p> <p>g. P10 and P200 pipette tips</p> <p>h. Permanent marker</p> <p>The glucose solution and the <em>in vivo</em> jetPEI transfection reagent are equilibrated at room temperature. </p> <p>Prepare 21 µl of each plasmid at a concentration of 78 and 88 ng / µl respectively (for a total volume of 42 µl =equals 3.5 µg of DNA) add the plasmid DNA to a 0.2 ml tube (labeled as Tube A) and mix with 21 µl of 10% glucose solution. </p> <p>In another microtube (labeled as Tube B), add 21 μl of 5% glucose solution and 1 μl of <em>in vivo</em> jetPEI. </p> <p> Prepare a third tube (labeled as Tube C) with 21 µl of 5% glucose solution and 0.5 µl of <em>in vivo</em> jetPEI to inject into embryos that will serve as controls. </p> <p>Leave the solutions at room temperature while you prepare the microinjection station. </p> <p> </p> <p> </p> <p><strong>4. Preparation of the micro-injection station</strong></p> <p><strong>Material:</strong></p> <p>a. Pre-pulled glass micropipettes (1mm diameter)</p> <p>b. Watch glass</p> <p>c. Modeling clay</p> <p>d. 35 mm and 90 mm petri dishes</p> <p>e. Mineral oil (M5904, SIGMA)</p> <p>f. Wash bottle with natural mineral water (Volvic)</p> <p>g. 0.2 ml microtubes</p> <p>h. 12-well cell culture plate</p> <p>i. Fine brush</p> <p>j. Phenol red solution</p> <p>k. Pasteur pipette or dropper</p> <p>l. Dissection forceps</p> <p>m. Soft holding forceps</p> <p>n. Snail eggs in the gastrula stage</p> <p>o. Drummond Scientific Nanoject III Programmable Nanoliter Injector</p> <p> </p> <p>Take a pre-pulled glass micropipette and cut it with a scalpel to have a ~ 0.2 mm tip slightly beveled if possible.</p> <p>Before attaching the micropipette to the programmable nanoliter injector, fill it with mineral oil. If this step is not done, the injector will not work properly. This can be done with a filling needle </p> <p>attached to a hamilton syringe of 10 microliters.</p> <p>When the micropipette is filled with oil, it must be fixed on the injector. For this it is necessary to: </p> <p>Slide the chuck and collet onto the glass micropipette, then slide the black O-ring with the seal onto the wire plunger </p> <p>With the micropipette attached to the injector, press the [EMPTY] icon until the plunger is fully extended. This step can be done with the footswitch by pressing once [EMPTY] then [STOP] and then proceeding [EMPTY] with the foot switch. A single beep is emitted when the plunger is fully extended.</p> <p>Fill the micropipette with 3 µl of the control solution or the transfection solution by placing the glass micropipette tip in a 0.2 ml tube with the solution to be injected and pressing the [FILL] icon. It is desirable to fill it at a slow rate, by pressing the [FILL] icon for a few seconds, then the [STOP] icon to allow the sample to equilibrate before pressing again the '[FILL] icon.</p> <p>Note: The piston continues to extend or retract until the [STOP] icon is pressed, or until the fully extended or fully retracted position is reached.</p> <p> </p> <p>5. Microinjection</p> <p>Place a watch glass into a 35mm petri dish and secure it on one side with modeling clay to form a slope. Use soft handling forceps to transfer an egg mass and lay it on the slope side of the watch glass so that the egg mass is in a sloping position.</p> <p>Remove excess water from the eggs with absorbent paper. Rehydrate if necessary with a fine brush to improve the visibility of the embryos. To inject the sample, return to the operating mode screen by pressing the [EXIT] icon, then select the injection mode by pressing the [INJECT] icon. Set the injection volume to 30nL and the flow rate to 20nL per second using the icons [+] and [-] respectively. Press the [INJECT] icon to inject the sample. </p> <p>Inject 30nL of the microinjection solution into each egg. Place the microinjected egg masses in a 12-well cell culture plate and note with a marker whether they were microinjected with the control solution or with the solution containing the plasmids.</p> <p>We colored the injection solution with red phenol to facilitate the visibility in this video.</p> <p><strong>Monitor the expression of the plasmids</strong></p> <p>Monitor the plasmids expression 72 h after microinjection in a contrast / fluorescent microscope or in a fluorescent stereo microscope. Then sort the fluorescent snails and perform a second micro-injection with a solution containing 10 µl of single guide RNA (at a concentration of 2ng / µl), add 0.5 µl of <em>in vivo</em> jetPEI reagent and 10 µl of 5% glucose solution. 3 days after the second microinjection, collect the hatched snails in a 1.5 ml tube containing 25 µl of lysis buffer for DNA and RNA purification.</p> <p>In this photo produced under a confocal microscope we washed a veliger larva in PBS solution, then we fixed it with 4% paraformaldehyde solution and then we placed it in a slide with two drops of the Dako fluorescence mounting medium. </p> <p>96 after the transfection we can observe the expression of the green fluorescent protein, the blue fluorescent protein and the co-localization of both proteins. </p> <p>This protocol is used to perform DNA methylation changes in a target gene. This transfection protocol can be used with other plasmids, with small interfering RNAs, or with messenger RNAs.</p> <p>Produced at IHPE (http://ihpe.univ-perp.fr)</p>
Coevolutionary theory of hosts and parasites
<p>Host and parasite evolution are closely intertwined, with selection for adaptations and counter-adaptations forming a coevolutionary feedback loop. Coevolutionary dynamics are often difficult to intuit due to these feedbacks and are hard to demonstrate empirically in most systems. Theoretical models have therefore played a crucial role in shaping our understanding of host-parasite coevolution. Theoretical models vary widely in their assumptions, approaches and aims, and such variety makes it difficult, especially for non-theoreticians and those new to the field, to: (1) understand how model approaches relate to one another; (2) identify key modelling assumptions; (3) determine how model assumptions relate to biological systems and (4) reconcile the results of different models with contrasting assumptions. In this review, we identify important model features, highlight key results and predictions and describe how these pertain to model assumptions. We carry out a literature survey of theoretical studies published since the 1950s (n=219 papers) to support our analysis. We identify two particularly important features of models that tend to have a significant qualitative impact on the outcome of host-parasite coevolution: population dynamics and the genetic basis of infection. We also highlight the importance of other modelling features, such as stochasticity and whether time proceeds continuously or in discrete steps, that have received less attention but can drastically alter coevolutionary dynamics. We finish by summarising recent developments in the field, specifically the trend towards greater model complexity, and discuss likely future directions for research.</p>
Data for: Termite nest evolution fostered social parasitism by termitophilous rove beetles
<p>Colonies of social insects contain large amounts of resources often exploited by specialized social parasites. While some termite species host numerous parasitic arthropod species, called termitophiles, others host none. The reason for this large variability remains unknown. Here we report that the evolution of termitophily in rove beetles is linked to termite nesting strategies. We compared one-piece nesters, whose entire colony life is completed within a single wood piece, to foraging species, which exploit multiple physically separated food sources. Our epidemiological model predicts that characteristics related to foraging (e.g., extended colony longevity and frequent interactions with other colonies) increase the probability of parasitism by termitophiles. We tested our prediction using literature data. We found that foraging species are more likely to host termitophilous rove beetles than one-piece nesters: 99.6% of known termitophilous species were associated with foraging termites, while 0.4% were associated with one-piece nesters. Notably, the few one-piece nesting species hosting termitophiles were those having foraging potential and access to soil. Our phylogenetic analyses confirmed that termitophily primarily evolved with foraging termites. These results highlight that the evolution of complex termite societies fostered social parasitism, explaining why some species have more social parasites than others.</p>
Photographs of parasites encapsulated in Cepaea nemoralis shells (appendix to the paper "Morph-dependent effect of nematode infection on host movement in the land snail *Cepaea nemoralis* (Mollusca, Gastropoda)")
<p>A set of four photographs taken of shell fragments of Cepaea nemoralis (grove snail), taken in the course of the project leading to the manuscript titled: "Morph-dependent effect of nematode infection on host movement in the land snail *Cepaea nemoralis* (Mollusca, Gastropoda)". Each photograph shows a parasite encapsulated/trapped in the shell by the snail:</p> <p>AcaRX3_3.tif: a mite (presumably Riccardoella sp.)<br> NemaBX7_1.tif, NemaBX7_3.tif, NemaRK5_2.tif: unidentified nematodes (note in the latter image, the clearly visible brown-band on yellow background pattern of the shell).</p> <p>A scale bar (0.25 mm) is overlaid on each image</p> <p> </p>
Fig. 3. A in Metazoan Parasites Infecting Xiphias Gladius From The Eastern Coast Of Algeria (Sw Mediterranean Sea)
Fig. 3. A — Pennella instructa (Wilson, 1917) attached to the host's body with Conchoderma virgatum (red star); B — section of P. instructa surrounded by connective tissue; C — general morphology of parasite P. instructa (female); D — cystic forms found into the musculature: n — neck; lh — lateral horns; t — trunk; p — plumes; (ct) — connective tissue and (pi) — P. instructa. Scale bars:, C, D = 2 cm; B— x4 = 300 Μm.
Fig. 4 in Metazoan Parasites Infecting Xiphias Gladius From The Eastern Coast Of Algeria (Sw Mediterranean Sea)
Fig. 4. General morphology of the species Hysterothylacium aduncum (Rudolphi, 1802) observed under an optical microscope. A — anterior part showing lips (l); nerve ring (nr); oesophagus (e); intestinal caecum (ic); ventriculus (v); ventricular appendix (va) and intestine (i), lateral view. B — middle part showing oesophagus (e); intestinal caecum (ic); ventriculus (v); ventricular appendix (va) and intestine (i), lateral view. C — dorsal labium showing labia (lb) lateral view. D — posterior end showing spinous tail (sp); lateral view. Scale bars: A — x10 = 100 µm; B, C, D — x40 = 50 µm.
Fig. 2. A, B in Metazoan Parasites Infecting Xiphias Gladius From The Eastern Coast Of Algeria (Sw Mediterranean Sea)
Fig. 2. A, B — Tristoma coccineum Cuvier, 1817 and Tristoma integrum Diesing, 1850 (see red circles) attached to gills of X. gladius: C — ventral view and D — dorsal view, of General morphology of the species Tristoma coccineum Cuvier, 1817; E — ventral view and F — dorsal view of General morphology of the species Tristoma integrum Diesing, 1850 observed under binocular magnifying glass: P — parasite, hap — haptor, alv — alveoli, pp — papillae. Scale bars: A, B — 2 cm; C, D, E, F — 2 m.
Fig. 1 in Review Of The Helminths Parasitic In Rare Aquatic Birds In Ukraine
Fig. 1. Localities of the material collection: — based on collection and literature; — based on collection; — based on literature.
Fig. 3 in Egg Batches Parasitism Of Processionary Moth, Thaumetopoea Pityocampa (Lepidoptera, Thaumetopoeidae), From Two Atlas Cedar Ecotypes In Algeria
Fig. 3. Distribution of the number of egg rows in relation to the twig diameter in Chréa (A), and Ouled Yagoub (B).
Fig. 1 in Egg Batches Parasitism Of Processionary Moth, Thaumetopoea Pityocampa (Lepidoptera, Thaumetopoeidae), From Two Atlas Cedar Ecotypes In Algeria
Fig. 1. Eggs batches of Thaumetopoea pityocampa: A — cylindrical form; B — egg batches in thick twigs; C — types of eggs.
Fig. 2 in Helminths Of Antarctic Rockcod Notothenia Coriiceps (Perciformes, Nototheniidae) From The Akademik Vernadsky Station Area (Argentine Islands, West Antarctica): New Data On The Parasite Community
Fig. 2. Average number of cysts of Corynosoma spp. in the body cavity of Notothenia coriiceps of five size groups.
Fig. 1 in Helminths Of Antarctic Rockcod Notothenia Coriiceps (Perciformes, Nototheniidae) From The Akademik Vernadsky Station Area (Argentine Islands, West Antarctica): New Data On The Parasite Community
Fig. 1. Prevalence (in %) and mean intensity of helminth species found in Notothenia coriiceps in the waters surrounding the Ukrainian Antarctic station "Akademik Vernadsky" in 2014–2015.
ScienceDex guides
Understand access before you commit
These curated guides explain access requirements, typical timelines, costs, and reuse considerations for widely used research datasets.
Allen Brain Atlas
Allen Brain Atlas is an Allen Institute collection of brain map atlases, datasets, APIs, and analysis tools covering mouse, human, and non-human primate brain resources.
Annotated Behaviour and Observability Dataset (ABODe)
ABODe is a University of Edinburgh DataShare dataset for behavior classification in group-housed mice using home-cage video, identities, bounding boxes, ground-plate positions, and annotator labels.
DANDI Archive for NWB datasets
DANDI is a BRAIN Initiative archive for publishing and sharing neurophysiology data, including electrophysiology, optophysiology, and behavioral data packaged as NWB and related standards.
International Brain Laboratory public data
The International Brain Laboratory public data releases expose standardized mouse decision-making experiments, including Neuropixels recordings, widefield calcium imaging, behavior, and session metadata accessed through the ONE API.
OpenNeuro
OpenNeuro is a free, open platform for sharing neuroimaging datasets, with public search, dataset pages, and download paths for web, S3, DataLad, and the OpenNeuro CLI.