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23 results for “centrohelids”
Fig. 3. Choanocystis symna spec. nov., strain CCAP 1597 in Two New Species of Centrohelid Heliozoans: Acanthocystis costata sp. nov. and Choanocystis symna sp. nov.
Fig. 3. Choanocystis symna spec. nov., strain CCAP 1597/1. (A) Scales of the air-dried specimen. Phase contrast. (B) Living cell. DIC. Contractile vacuoles (cv), kinetocysts (k), microtubule-organizing center (MTOC) and spine scales can be seen. (C) Living cell compressed with a cover-slip. Phase contrast. Nucleus (n) with a central nucleolus (nu) is visible. (D) Air-dried whole-mount of the cell. Type-slide NHMUK 2013.6.28.1. SEM. Spine scales and plate scales are visible. (E) The most common quantity of teeth on the tip of spine scale is four. SEM. (F) Spine scale with three teeth. SEM. (G) Tips of spine scales with five (left) and four (right) teeth. SEM. Scale bars: 5 μm (A–C), 10 μm (D), 1 μm (E–G).
Fig. 6. RAxML tree for 18S in Two New Species of Centrohelid Heliozoans: Acanthocystis costata sp. nov. and Choanocystis symna sp. nov.
Fig. 6. RAxML tree for 18S rRNA genes of 8 pterocystid heliozoans (1320 nucleotide positions). Only bootstrap values more than 50% are shown. New sequence is in bold.
Fig. 5. RAxML tree for 18S in Two New Species of Centrohelid Heliozoans: Acanthocystis costata sp. nov. and Choanocystis symna sp. nov.
Fig. 5. RAxML tree for 18S rRNA genes of 16 heliozoans from the genus Acanthocystis and Polyplacocystis ambigua as an outgroup (1520 nucleotide positions). Only bootstrap values more than 50% are shown. New sequence is in bold. Crossed branches were shortened fourfold.
Fig. 7 in Immunocytochemical Analysis of α-Tubulin Distribution Before and After Rapid Axopodial Contraction in the Centrohelid Raphidocystis contractilis
Fig. 7. (A, B) The distribution of α-tubulin in the cell after rapid axopodial contraction. (A) Low-magnification images of the cell after the induction of rapid axopodial contraction. A projection image of the whole cell constructed from 87 optical sections taken at a 0.5 µm interval (left) and the corresponding light micrograph (right). (B) High-magnification images of the same cell shown in (A). A projection image of the equatorial plane of the cell body constructed from 5 optical sections taken at a 0.5 µm interval (left), and the corresponding light micrograph (right). Note that the positive signals in the distal parts of the axopodia have a branched appearance (arrowheads). Scale bars: 20 μm (A), 10 μm (B).
Fig. 4 in Immunocytochemical Analysis of α-Tubulin Distribution Before and After Rapid Axopodial Contraction in the Centrohelid Raphidocystis contractilis
Fig. 4. (A–D) Evaluation of the maintenance of the original lengths of axopodia in R. contractilis using different combinations of fixative and buffer. Cells were fixed with (A) 4% paraformaldehyde in phosphate buffer, (B) 4% paraformaldehyde in PHEM, (C) 0.2% glutaraldehyde in phosphate buffer, or (D) 0.2% glutaraldehyde in PHEM. Note that only fixation with 0.2% glutaraldehyde in PHEM maintained the axopodial length. Scale bar: 20 µm.
Fig. 1 in Immunocytochemical Analysis of α-Tubulin Distribution Before and After Rapid Axopodial Contraction in the Centrohelid Raphidocystis contractilis
Fig. 1. (A, B) Rapid axopodial contraction induced by mechanical stimulation in R. contractilis. Images (A) before and (B) after rapid axopodial contraction. Arrowheads indicate kinetocysts. Note the synchronized retraction of all axopodia and the apparent increases in the widths of contracted axopodia relative to the features observed before the onset of axopodial contraction (an arrow). Scale bar: 20 µm (A, B).
Fig. 6 in Immunocytochemical Analysis of α-Tubulin Distribution Before and After Rapid Axopodial Contraction in the Centrohelid Raphidocystis contractilis
Fig. 6. Confocal analysis of α-tubulin-immunolabeled extended axopodia. Fine reconstructed projection image from 161 optical sections taken at a 0.05 µm (top) and the corresponding light micrograph (bottom). Note the regions of relatively low fluorescence (arrowheads) did not correspond to the location of kinetocysts (arrows). Scale bar: 10 μm.
Fig. 5 in Immunocytochemical Analysis of α-Tubulin Distribution Before and After Rapid Axopodial Contraction in the Centrohelid Raphidocystis contractilis
Fig. 5. (A, B) The distribution of α-tubulin in the cell before rapid axopodial contraction. (A) Low-magnification images of a cell without the induction of rapid axopodial contraction. A projection image of the whole cell was constructed from 192 optical sections obtained at 0.5 µm intervals (left); the corresponding light micrograph is also shown (right). (B) High-magnification images of the same cell shown in (A). A projection image of the equatorial plane of the cell body constructed from 5 optical sections taken at 0.5 µm intervals, and the corresponding light micrograph. The asterisk indicates a centroplast. Note that positive signals were detected along the fully extended axopodia. Scale bars: 20 μm (A), 10 μm (B).
Fig. 3 in Immunocytochemical Analysis of α-Tubulin Distribution Before and After Rapid Axopodial Contraction in the Centrohelid Raphidocystis contractilis
Fig. 3. Schematic illustration of the experimental setup. The fixative is injected into the cell suspension via a micro flow-through chamber with a syringe pump. Next, the cells are fixed by injecting the fixative at a rate below the threshold required for inducing rapid contraction. The cells are then observed under a microscope.
Fig. 2 in Immunocytochemical Analysis of α-Tubulin Distribution Before and After Rapid Axopodial Contraction in the Centrohelid Raphidocystis contractilis
Fig. 2. (A, B) Fine structures associated with axopodial microtubules. (A) Centroplast in the center of the cell. (B) Cross-section of an axopodium in the peripheral region of the cell. Note that bundles of microtubules radiate from the centroplast (arrowheads) and that the axopodium comprises six microtubules. Scale bars: 500 nm (A), 100 nm (B).
Fig. 2 in New Freshwater Species of Centrohelids Acanthocystis lyra sp. nov. and Acanthocystis siemensmae sp. nov. (Haptista, Heliozoa, Centrohelea) from the South Urals, Russia
Fig. 2. SEM of Acanthocystis siemensmae sp. nov. A – General view of the scales. B – Long spine scales. C – Apices of long spine scales with ridges. D – Plate scale, long and short spine scale with basal plate. E – Apices of short spine scales with ridges. F – Ridges with teeth at the apex of short spine scale. G – Plate scale with marginal rim. Abbreviations: bp – basal plate; ct – central tooth; ls – long spine scale; lt – lateral tooth; mr – marginal rim; ps – plate scale; r – ridge; sh – shaft; ss – short spine scale; t – teeth. Scale bars: A – 10 µm; B, C, E, G – 1 µm; D – 2 µm, F – 0.5 µm.
Fig. 1 in New Freshwater Species of Centrohelids Acanthocystis lyra sp. nov. and Acanthocystis siemensmae sp. nov. (Haptista, Heliozoa, Centrohelea) from the South Urals, Russia
Fig. 1. SEM of Acanthocystis lyra sp. nov. A – General view of the scales. B – Long spine scale. C – Lyrate distal end of long spine scale with teeth on inner edge. D – Short spine scale with secondary bifurcation. E – Short spine scale with primary bifurcation and teeth. F – Short spine scales with primary and secondary bifurcations. A marginal rim is seen on basal plates of short and long spine scales. G – Plate scales ornamented with an axial thickening. Abbreviations: at – axial thickening; b1 – primary bifurcation; b2 – secondary bifurcation; bp – basal plate; br – branch of furca; mr1 – marginal rim on basal plate of short spine scale; mr2 – marginal rim on basal plate of long spine scale; sh – shaft; t – teeth. Scale bars: A – 10 µm; B, D, E, F, G – 1 µm; C – 0.5 µm.
Fig. 3 in New Freshwater Species of Centrohelids Acanthocystis lyra sp. nov. and Acanthocystis siemensmae sp. nov. (Haptista, Heliozoa, Centrohelea) from the South Urals, Russia
Fig. 3. Line drawings of scales of Acanthocystis lyra sp. nov. (A–C) and Acanthocystis siemensmae sp. nov. (D–F): A, D – long spine scales; B, E – short spine scales; C, F – plate scales.
Fig. 8 in Centrohelid Heliozoans (Centroplasthelida Febvre-Chevalier et Febvre, 1984) from Different Types of Freshwater Bodies in the Middle Russian Forest-steppe
Fig. 8. Dendrogram showing the Bray-Curtis similarity (%) of studied microbiotopes by species diversity of centrohelids. Abbreviations: p.d.s – plant debris and silt; pt – peat from mire's lakes; sn – sand; sph – Sphagnum from hollows; wt – water column.
Fig. 1 in Centrohelid Heliozoans (Centroplasthelida Febvre-Chevalier et Febvre, 1984) from Different Types of Freshwater Bodies in the Middle Russian Forest-steppe
Fig. 1. Morphology of living cells of some observed centrohelid species (DIC): A – Acanthocystis nichollsi; B – A. pectinata (from plant debris and silt of Usman' River); C – A. turfacea (from plant debris and silt of Lake Chistoye); D – Raphidocystis symmetrica (from plant debris and silt of Lake Chistoye); E – Choanocystis aculeata (from sand of Lake Maklok); F – Raphidiophrys capitata (from sand of Lake Maklok). Abbreviations: ax – axopodia; pr – protoplast; sc – scales. Scale bar: 20 µm.
Fig. 7 in Centrohelid Heliozoans (Centroplasthelida Febvre-Chevalier et Febvre, 1984) from Different Types of Freshwater Bodies in the Middle Russian Forest-steppe
Fig. 7. Morphology of observed scales of unidentified species (TEM): A–C – Heterophrys-like organism (A – from plant debris and silt of Usman' River; B, C – from plant debris and silt of Lake Krugloye; A – whole cell; B, C – spicules), D–F – unidentified Centrohelea sp. (from water column of Lake Vosmerka; D – scales of the single cell; E, F – plate scales). Abbreviations: sp – spicules. Other abbreviation as in Fig. 1. Scale bars: A, D – 10 µm; B, C, E, F – 1 µm.
Fig. 6 in Centrohelid Heliozoans (Centroplasthelida Febvre-Chevalier et Febvre, 1984) from Different Types of Freshwater Bodies in the Middle Russian Forest-steppe
Fig. 6. Morphology of observed scales of Raineriophrys genera (A–C, E, F, I–L – TEM; D, G, H – SEM): A–D – R. echinata (A–C – from peat of Sphagnum bog Klukvennoye-2; D – from Sphagnum of Sphagnum bog Klukvennoye-3), E–H – R. erinaceoides (E, F – from water column of Lake Vosmerka; G, H – from plant debris and silt of Lake Cherepashye), I–L – R. fortesca (I – Vosmerka, J–L – Cherepashye). E, I – scales of the single cell; A–D, F–H, J–L – spine and plate scales. Abbreviations as in Fig. 2, 5. Scale bars: A, E, I – 10 µm; B–D, F – 5 µm; G, H, J–L – 1 µm.
Fig. 9 in Centrohelid Heliozoans (Centroplasthelida Febvre-Chevalier et Febvre, 1984) from Different Types of Freshwater Bodies in the Middle Russian Forest-steppe
Fig. 9. Dendrogram showing the Bray-Curtis similarity (%) of studied water bodies by species diversity of centrohelids.
Fig. 3 in Puzzle-like Cyst Wall in Centrohelid Heliozoans Raphidiophrys heterophryoidea and Raineriophrys erinaceoides
Fig. 3. Raphidiophrys heterophryoidea. Light microscopic images of the cyst (A, B) and excysted organism (C, D, E). Differential interference contrast (A, B, D) and phase contrast (C, E). A – intact cyst; B – a cyst, crushed with a cover slip; C – excysted individual, compressed with a cover slip; D – excysted individual: view on the cell surface and axopodia; E – excysted individual in a Petri dish. Scale bars: 10 μm.
Fig. 2 in Puzzle-like Cyst Wall in Centrohelid Heliozoans Raphidiophrys heterophryoidea and Raineriophrys erinaceoides
Fig. 2. Raineriophrys erinaceoides. Scanning electron micrographs of air-dried excysted individuals. A – general view; B–G, I, J – individual cyst scales; H – flexible and undersized trophic scales. Scale bars: 2 μm.
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