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141 results for “microtubules”
Training and test dataset of STED images of microtubules in fixed cells
<p>Training and test dataset of microtubule used in the manuscript "Denoising diffusion models for high-resolution microscopy image restoration".</p>
PhasAGE Expert Seminar- Dissecting the contribution of motor-cargo adaptors to microtubule-based transport in neurons
<p>The PhasAGE Expert Seminars consist of a series of talks with speakers from PhasAGE partner’s institutions to promote a successful transfer of knowledge about PhasAGE topics – biomolecular phase separation, aging and age-related diseases.</p>
RMSD and Trp assays for "Structural analysis of a motor with increased mechanical output reveals new transitions in kinesin microtubule motility"
<p>This entry is for our manuscript, "Structural analysis of a motor with increased mechanical output reveals new transitions in kinesin microtubule motility" by Satoki Shibata*, Matthew Y. Wang*, Tsuyoshi Imasaki*, Hideki Shigematsu, Diego Ugarte La Torre, Yuanyuan Wei, Chacko Jobichen, Hajime Hagio, J. Sivaraman, Yuji Sugita, Sharyn A. Endow & Ryo Nitta.</p> <p>*Equal contribution</p> <p>Corresponding Authors: Tsuyoshi Imasaki*, Sharyn A. Endow, Ryo Nitta</p> <p>The deposited datasets are 1) beta-strand residue all-atom RMSD between kinesin-14 NcdY485K ADP and ADP + free Pi nucleotide states and 2) fluorimeter assays of NcdY485K intrinsic Trp fluorescence changes upon addition of free Pi . The data are presented in Fig. 6e, Fig. 7b,c and Supplementary Fig 10. Files are named for the figures in which the data are shown. A computer script for the RMSD analysis and methods for the fluorimeter assays and data analysis are included in the deposited files.</p>
Dataset of image processing - High-throughput characterization of cortical microtubule arrays response to anisotropic tensile stress
<p>The data set contains the analysis data files from the image analysis workflow developed to quantify cortical microtubules rearrangements in the case of tensile stress (<a href="https://github.com/VergerLab/MT_Angle2Ablation_Workflow">https://github.com/VergerLab/MT_Angle2Ablation_Workflow</a>), generated form a specific dataset (https://doi.org/10.5878/17te-jg54). The files include the intermediary images processed at each step of the image analysis workflow in imageJ, the log files produced by the imageJ macro describing the input and the output images and the text files containing the quantified values. </p>
Distinct States in an Active Liquid Crystal of Flagella and Kinesin-Powered Microtubules under Varying Kinesin Concentrations or Active Stress
<p>At a constant nematic elasticity (flagella concentration) and confinement (H=100μm), higher KSA (120nM) or motor concentrations induce robust active stress that disrupts nematic alignment, resulting in chaotic behavior. Conversely, lower KSA (<60nM) concentrations fail to overcome the nematic elastic background's alignment, leading to a unique state where active microtubule bundles phase-separate into lanes within a uniform flagella background. The confocal micrograph reveals this distinct phenomenon, showing uniform flagella distribution alongside microtubule segregation into lane-like structures within the channel's limited confinement.</p>
Distinct States in an Active Liquid Crystal of Flagella and Kinesin-Powered Microtubules under Varying Confinement
<p>At significant confinement levels, the potent active stress disrupts the alignment of the flagella, leading to a turbulent state in the composite liquid crystal. Conversely, minimal confinement enables the dominant nematic alignment to counteract the active stress, inducing the phase separation of microtubules within a uniform flagella background. The confocal micrograph unveils uniform flagella distribution across the medium, while the microtubules segregate into a distinct lane-like structure within the channel's narrowest confinement.</p>
Microtubules growing and shortening under constant force
<div class="abstract"> <div class="abstract-content selected"> <p>Kinetochores are macromolecular machines that couple chromosomes to dynamic microtubule tips during cell division, thereby generating force to segregate the chromosomes. Accurate segregation depends on selective stabilization of correct 'bi-oriented' kinetochore-microtubule attachments, which come under tension as the result of opposing forces exerted by microtubules. Tension is thought to stabilize these bi-oriented attachments indirectly, by suppressing the destabilizing activity of a kinase, Aurora B. However, a complete mechanistic understanding of the role of tension requires reconstitution of kinetochore-microtubule attachments for biochemical and biophysical analyses <em>in vitro</em>. Here we show that native kinetochore particles retaining the majority of kinetochore proteins can be purified from budding yeast and used to reconstitute dynamic microtubule attachments. Individual kinetochore particles maintain load-bearing associations with assembling and disassembling ends of single microtubules for >30 min, providing a close match to the persistent coupling seen<em> in vivo</em> between budding yeast kinetochores and single microtubules. Moreover, tension increases the lifetimes of the reconstituted attachments directly, through a catch bond-like mechanism that does not require Aurora B. On the basis of these findings, we propose that tension selectively stabilizes proper kinetochore-microtubule attachments in vivo through a combination of direct mechanical stabilization and tension-dependent phosphoregulation.</p> </div> </div>
Paired microtubules growing with a shared load
<p>During mitosis, kinetochore-attached microtubules form bundles (k-fibers) in which many filaments grow and shorten in near-perfect unison to align and segregate each chromosome. However, individual microtubules grow at intrinsically variable rates, which must be tightly regulated for a k-fiber to behave as a single unit. This exquisite coordination might be achieved biochemically, via selective binding of polymerases and depolymerases, or mechanically, because k-fiber microtubules are coupled through a shared load that influences their growth. Here, we use a novel dual laser trap assay to show that microtubule pairs growing <em>in vitro</em> are coordinated by mechanical coupling. Kinetic analyses show that microtubule growth is interrupted by stochastic, force-dependent pauses and indicate persistent heterogeneity in growth speed during non-pauses. A simple model incorporating both force-dependent pausing and persistent growth speed heterogeneity explains the measured coordination of microtubule pairs without any free fit parameters. Our findings illustrate how microtubule growth may be synchronized during mitosis and provide a basis for modeling k-fiber bundles with three or more microtubules, as found in many eukaryotes.</p>
Structural basis for Parkinson's Disease-linked LRRK2's binding to microtubules
<p>This dataset includes all of the tabular data used in the figures in the article. Original article is available at: https://doi.org/10.1101/2022.01.21.477284</p> <p>Leucine Rich Repeat Kinase 2 (<em>LRRK2</em>) is one of the most commonly mutated genes in familial Parkinson’s Disease (PD). Under some circumstances, LRRK2 co-localizes with microtubules in cells, an association enhanced by PD mutations. We report a cryo-electron microscopy structure of the catalytic half of LRRK2, containing its kinase, which is in a closed conformation, and GTPase domains, bound to microtubules. We also report a structure of the catalytic half of LRRK1, which is closely related to LRRK2, but is not linked to PD. LRRK1’s structure is similar to LRRK2, but LRRK1 does not interact with microtubules. Guided by these structures, we identify amino acids in LRRK2’s GTPase domain that mediate microtubule binding; mutating them disrupts microtubule binding in vitro and in cells, without affecting LRRK2’s kinase activity. Our results have implications for the design of therapeutic LRRK2 kinase inhibitors.</p>
Microtubule retrograde flow retains neuronal polarization in a fluctuating state
<p>In developing vertebrate neurons, a neurite is formed by more than a hundred microtubules. While individual microtubules are dynamic, the microtubule array has been regarded as stationary. Using live-cell imaging of neurons in culture or in brain slices, combined with photoconversion techniques and pharmacological manipulations, we uncovered that the microtubule array flows retrogradely within neurites to the soma. This flow drives cycles of microtubule density, a hallmark of the fluctuating state before axon formation, thereby inhibiting neurite growth. The motor protein dynein fuels this process. Shortly after axon formation, microtubule retrograde flow slows down in the axon, reducing microtubule density cycles and enabling axon extension. Thus, keeping neurites short is an active process. Microtubule retrograde flow is a novel type of cytoskeletal dynamics, which changes the hitherto axon-centric view of neuronal polarization.</p>
Figures 27–30 in Cell structure and microtubule organisation during gametogenesis of Ulva mutabilis Føyn (Chlorophyta)
Figures 27–30: Tubulin immunofluorescence in released gametes of Ulva mUtabilis. (27) Released gamete bearing two equal flagella. Note that cortical microtubule bundles (tubulin fluorescence) extend from basal body towards the posterior. Scale bar: 20 µm. (28) Differential interference contrast microscopy-image of gamete in Figure 27. Scale bar: 20 µm. (29 and 30) Two gametes in which (apart from the two flagella) an intensely fluorescent microtubule bundle starting from flagellar basal body is directed to cytoplasm (arrows). Scale bar: 20 µm.
Figures 5–8 in Cell structure and microtubule organisation during gametogenesis of Ulva mutabilis Føyn (Chlorophyta)
Figures 5–8: Transmission electron micrographs of cells of Ulva mUtabilis 48 h after the induction of gametogenesis. (5) The cell is polarised with most of the cytoplasm and organelles gathered at the tip area. The external cell wall has formed a projection towards the apical region. The cell wall appears differentiated at this area (N: nucleus). Scale bar: 2 µm. (6) Detail of the apical wall of the previous image, showing the loose structure of the wall and the formation of amorphous dark-stained material. Scale bar: 0.35 µm. (7) More advanced stage of a wall projection. The dark-stained material covers the projection, while its central part appears as a plug, consisting of amorphous material. The internal wall layer under the tip is considerably thickened and pushes the plug out (asterisk). Scale bar: 4 µm. (8) Cross-section of a cell with a plug-like cap. The dark-stained material is extended covering the cap. Scale bar: 0.33 µm.
Figures 9–14 in Cell structure and microtubule organisation during gametogenesis of Ulva mutabilis Føyn (Chlorophyta)
Figures 9–14: Immunolocalization of tubulin in developing gametangia of Ulva mUtabilis. (9) Interphase cell before induction of gametogenesis. The microtubules are mainly peripherally arranged in more or less parallel bundles. Scale bar: 20 µm. (10) Early stage, 36–48 h after induction of gametogenesis. Intensely fluorescent microtubule bundles traverse the cortical cytoplasm converging on a particular area (arrow localises the conical cell projection). Scale bar: 20 µm. (11) More advanced stage, 48–60 h after induction of gametogenesis, showing microtubules starting from a pointed site at basal part of gametangium. Scale bar: 20 µm. (12) Αdvanced stage, 48–60 h after induction of gametogenesis, showing microtubules organised in a basketlike configuration, leaving a circular opening at the top (arrow). Scale bar: 20 µm. (13) Gametangium, 60–72 h after induction of gametogenesis. The microtubule-free area at the top appears broader than before (arrow). Scale bar: 20 µm. (14) Hoechst 33258 staining of the nucleus of the cell in Figure 12. Scale bar: 20 µm.
Figures 2–4 in Cell structure and microtubule organisation during gametogenesis of Ulva mutabilis Føyn (Chlorophyta)
Figures 2–4: Vegetative cells of Ulva mUtabilis before induction of gametogenesis. (2) Light micrograph of a semithin section showing a unilayered thallus. Note the difference in staining between external and internal cell wall layer. Scale bar: 20 µm. (3) Transmission electron micrograph of a thallus cell before induction of gametogenesis. The nucleus (N) has a relatively central position and the chloroplast (CHL) occupies a large part of the cytoplasm (CYT). Note the thick external wall. Scale bar: 3.6 µm. (4) Detail of external wall of a cell before induction of gametogenesis. Wall shows strict stratification, i.e. a thick internal amorphous layer, a median region with multiple fibrillar layers and an external dark-stained region (see brackets). Scale bar: 1 µm.
Figure 1 in Cell structure and microtubule organisation during gametogenesis of Ulva mutabilis Føyn (Chlorophyta)
Figure 1: Time course of induced gametogenesis and discharge of gametangia from Ulva mUtabilis in relation to the swarming inhibitor (SWI) synthesis and action (summary of new and previous results). Gametogenesis was induced in Ulva mUtabilis [mutant slender sl-G(mt +)]. The "determination phase" and "differentiation phase" are defined as described by Stratmann et al. (1996). The "swarming phase" is the time period when gamete release can be induced by light or a medium change, or may occur later spontaneously. (A) Regular vegetative G1 cell cycle phase in which gametogenesis can be induced by the removal of sporulation inhibitors SI-1 and SI-2 from the medium. (B) Regular vegetative S phase in which the genome is replicated normally or, after induction of gametogenesis, the period of SWI synthesis and excretion. (C) Next G1 phase after induction of gametogenesis. (D) Next S phase after induction of gametogenesis and accumulation of starch granules. (E) Time of irreversible commitment to gametangium differentiation. (F) Period of progamete formation, chloroplast reorientation and papilla initiation. (G) Period of progamete multiplication to 16 cells, and papilla maturation. (H) Period of gamete and pore cap maturation. (I) Period when the exit pores are open and when gamete release can be induced by light and/or depletion of SWI in the medium. (J) Period when the gametangia become insensitive to SWI and gamete release may occur spontaneously and asynchronously. Figure is from Wichard and Oertel (2010) with permission of © Wiley 2010. Images show the transformation of blade cells into gametangia. Microscopic images for stages a, f, h and i are shown (scale bars: 25 µm).
Fig. 5 in Cortical Localization of α- and γ-Tubulin and the Assembly of Cortical Microtubule Cytoskeleton in Hypotrichous Ciliate Euplotes eurystomus
Fig. 5. (A–D) Projections of the optical sections passing through the ventral surface of E. eurystomus cells in mitotic phase decorated with FLUTAX. Scale: 20 μm. (A) The AZM primordium appeared at the bottom-left of old AZM in the early stage of morphogenesis. (AZMP: adoral zone of membranelles primordium; UMP: undulating membrane primordium; arrow shows the UMP originated from the right side of AZMP). (B) The pattern of FVTC primordia was 3-3-3-2-2 from left to right, shown in a box (FVTCP: frontal–ventral–transverse cirri primordium; arrows show the blastema of ALM and PLM originating from the base of FVTCPs). (C) The AZM and UM of proter inherited the olds (white arrow), and those of opisthe were generated from the AZMP (black arrows show the primordium of the left marginal cirri). (D) Some old ciliary organelles still existed until the separation of the proter and opisthe (white arrows show the neonatal ALM and PLM; black arrows show the residual ALM; some old cirri are shown in the circle).
Fig. 3 in Cortical Localization of α- and γ-Tubulin and the Assembly of Cortical Microtubule Cytoskeleton in Hypotrichous Ciliate Euplotes eurystomus
Fig. 3. (A–C) Projection of optical section passing through the ventral surface of E. eurystomus cells decorated with the FLUTAX. Scale: 20 μm. (A) All the ventral ciliary organelles, and especially their base-associated microtubules, were strongly and clearly stained by FLUTAX (ALM: anterior longitudinal microtubules; PLM: posterior longitudinal microtubules; TM: transverse microtubules; PM: peripheral microtubules). (B) The microtubules associated with the adoral ciliary organelles were also marked by FLUTAX (SMB: small membranelle bracket; hollow arrow shows the microtubule fasolculus extending from the base of SMB; white arrow shows the microtubules in the adoral rib; black arrow shows the UM ciliary base–associated microtubules). (C) The FLUTAX decorated the short microtubular bundles between the bases of TCs (arrow).
Fig. 4 in Cortical Localization of α- and γ-Tubulin and the Assembly of Cortical Microtubule Cytoskeleton in Hypotrichous Ciliate Euplotes eurystomus
Fig. 4. (A–D) Projections of the optical sections passing through the dorsal cortex of E. eurystomus cells decorated with FLUTAX. Scale: 20 μm. (A) The leptos microtubular nets covered the dorsal cortex (arrow). (B) The longitudinal microtubules in the dorsal cortex (arrow). (C) The oblique microtubules in the dorsal cortex (arrow). (D) The dargyrome was faintly stained by the FLUTAX.
Fig. 1 in Cortical Localization of α- and γ-Tubulin and the Assembly of Cortical Microtubule Cytoskeleton in Hypotrichous Ciliate Euplotes eurystomus
Fig. 1. The permutation pattern of ciliary organelles in the ventral and dorsal of E. eurystomus cells (A, B). (A) The ventral ciliary pattern of E.eurystomus, (B) the dorsal ciliary pattern of E.eurystomus. (AZM: adoral zone of membranelles, OC: oral cirri; UM: undulating membranes, FVC: frontal–ventral cirri, TC: transverse cirri, CC: caudal cirri, LMC: left marginal cirri DK: dorsal kineties).
Figure 3 in The function of ciliopathy protein FOP on cilia and cortical microtubule cytoskeleton in Euplotes amieti
Figure 3. Protein loss and motility changes after RNAi. (A) Relative expression of basal body-associated protein BBS8. **P <0.01 vs the blank group, n = 3. (B) Relative expression of cilia assembly-related protein IFT88. **P <0.01 vs the blank group, n = 3. (C) Relative expression of basal body protein γ-tubulin. (D) The swimming tracks of Euplotes amieti during RNAi. The swimming speeds (n = 20). The FOPRNAi is the interference group, The Blank is the blank group (E) The swimming speeds of Euplotes amieti during RNAi (n = 20).
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Allen Brain Atlas
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Annotated Behaviour and Observability Dataset (ABODe)
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