We thank the members of the Light Microscopy Facility, Max-Planck Institute of Experimental Medicine, for their expert advice and support.

1. Seeding of HeLa Cells
<ol>
	<li>Prepare 2 mL of 5 &#956;g/mL fibronectin solution in phosphate buffered saline (PBS) and add 450 &#956;L of it into each well of a 4 well chambered coverslip (#1.5). Incubate the slide for 15 min at 37 &#176;C and 5% CO2.</li>
	<li>Rinse asynchronously growing HeLa cells with Dulbecco&#8217;s Phosphate Buffered Saline (DPBS) and incubate with trypsin-EDTA (0.05%: 0.02%; w:v) for 5 min at 37 &#176;C. Stop the enzymatic reaction by the addition of Roswell Park Memorial Institute (RPMI) 1640 medium supplemented with 10% heat-inactivated fetal calf serum (FCS) at 3:1 (v:v) ratio of added trypsin-EDTA. 
	NOTE: HeLa cells were maintained in RPMI 1640 medium supplemented with 10% heat-inactivated FCS at 37 &#176;C and 5% CO2 and were routinely passaged once they reached 80&#8211;90% confluency as described above.</li>
	<li>Determine the cell concentration using a Neubauer chamber. Mix a 50 &#956;L aliquot of the cell suspension with trypan blue at 1:1 (v:v) ratio, resuspend, and transfer 10 &#956;L of the suspension into the chamber. Count only the trypan blue-negative cells inside of the four large squares (for details see Phelan et al.47). Derive the cell concentration from the counted cell number using the following formula: 
	<img alt="Equation 1" src="/files/ftp_upload/60478/60478eq1.jpg" /></li>
	<li>Pellet the cells by centrifugation at 300 x g for 2 min. Resuspend with fresh RPMI 1640 in order to obtain 1 x 106 cells/mL.</li>
	<li>Remove the fibronectin from the chambered coverslip, wash the wells twice with DPBS, and seed 50,000 cells per well.</li>
	<li>Return the chambered coverslip with the cells to the incubator and grow them for 24 h at 37 &#176;C and 5% CO2.</li>
</ol>
2. Expression of pEB3-tdTomato in HeLa Cells
<ol>
	<li>Prepare a 1.5 mL microcentrifuge tube. For each tube, dilute 2 &#956;g of pEB3-tdTomato48 with transfection buffer (synthetic product in aqueous solution) to a final volume of 396 &#956;L.</li>
	<li>Add 4 &#956;L of transfection reagent (non-lipidic, containing polyethylenimine) to the first tube, and vortex the mixture immediately for exactly 10 s.</li>
	<li>Briefly spin down the tube with a microcentrifuge and incubate at room temperature (RT) for 10 min.</li>
	<li>Remove the HeLa cells from the incubator. Dropwise, add 100 &#956;L of the transfection mixture to each well of a 4 well chambered coverslip, and return the cells to the incubator.</li>
	<li>After 4 h of incubation at 37 &#176;C and 5% CO2, supplement the cells with fresh growth medium and incubate for at least 24 h at 37 &#176;C and 5% CO2. 
	NOTE: It is necessary to optimize transfection conditions for each cell type. The expression levels need to be low enough to allow the identification of single MT growing ends. Alternatively, a cell line stably expressing EB3-tdTomato can be used in the experiments; this would reduce variability in expression levels of EB3-tdTomato between preparations and between cells from the same preparation49.</li>
</ol>
3. Synchronization and Live-cell Imaging of pEB3-tdTomato&#8211;expressing HeLa Cells
<ol>
	<li>Prepare a 2.5 &#956;M solution of dimethylenastron (DME) in phenol-red free Dulbecco&#39;s Modified Eagle Medium (DMEM) supplemented with 10% FCS and 2 mM L-glutamine or an alternative glutamine supply.</li>
	<li>Replace the growth medium in the chambered coverslip with 500 &#956;L of the growth medium containing 2.5 &#956;M DME and incubate the cells at 37 &#176;C and 5% CO2.</li>
	<li>After 3.5 h of incubation with DME, transfer the cells to the microscope, mount the chambered coverslip into an environmental chamber with dark panels for imaging at 37 &#176;C and 5% CO2, and further incubate until the total incubation time is 4 h. 
	NOTE: The maintenance of temperature at 37 &#176;C without fluctuation is crucial for the experiment.</li>
	<li>Perform the time-lapse imaging on an inverted microscope equipped with a 100x 1.49 N.A. oil immersion objective, a dual spinning disk confocal system, and a reliable autofocus system for continuous maintenance of the focal plane. Define the imaging parameters as follows. 
	NOTE: We use an Electron Multiplying Charge-Coupled Device camera (EM-CCD).
	<ol>
		<li>For EB3-tdTomato excitation, use a 561 nm laser line with 200 ms exposure time. Collect the emitted light through a quadruple bandpass (405, 488, 561, 640 nm) dichroic mirror and a 600/52 nm emission filter. 
		NOTE: Laser power can be adjusted for each imaged cell to prevent image saturation. In all time-lapse movies given here the laser power was set to 5.3 mW.</li>
		<li>Find a cell in prophase and focus in the Z-plane corresponding to the center of the monopolar mitotic spindle. Acquire images every 0.5 s over a total of 1 min with no binning and no illumination between the exposures.</li>
	</ol>
	</li>
</ol>
4. Analysis of the MT Dynamics Using U-Track v2.2.0
<ol>
	<li>To analyze the MT dynamics a numerical computing environment software is required (e.g., MATLAB). 
	NOTE: Basic understanding of the software is sufficient for the analysis. Comprehensive help material and tutorials are available on the developer&#8217;s website (https://uk.mathworks.com/products/&#173;matlab/&#173;getting-started.html).</li>
	<li>Download (https://github.com/DanuserLab/u-track) and install the open-source U-Track v2.2.0 software following the detailed instructions given in the &#34;Readme_u-track.pdf&#34; file50,51,52.</li>
	<li>Launch the numerical-analysis software and add U-Track v2.2.0 folder with subfolders into the software search path.</li>
	<li>From the command window call &#34;movieSelectorGUI&#34;. This opens a dialogue window from which the raw files generated by the image acquisition software at the microscope can be imported (Supplementary Figure 1, Figure 2, Figure 3, Figure 4). 
	NOTE: The U-Track software is compatible with other image data formats. It uses Bio-Formats, which recognizes different life science data formats53.</li>
	<li>The size of each image is read from the metadata automatically. Manually enter the numerical aperture of the objective (in this case 1.49) and the time interval (0.5 s) used for imaging (Supplementary Figure 1B). Additionally, information on the excitation wavelength, the fluorophore, and the exposure time can also be provided, but they are not critical for further analysis.</li>
	<li>Once all the images are loaded, save the entered time-lapse series as a movie list by selecting the &#34;Save As Movie List&#34;. On the right side of the dialogue window select the &#34;U-Track&#34; option and press &#34;Continue&#34; (Supplementary Figure 1C). 
	NOTE: The values are optimized for HeLa cells. If switching to a different cell line, the values should be defined again. Alternatively, use the settings recommended by the software developers. The detailed explanation of each of the parameters and how they should be defined can be found in the technical report provided with the previous version of the software, plusTipTracker50.</li>
	<li>From the pop-up window select &#34;Microtubule Plus-Ends&#34; and press &#34;Ok&#34; (Supplementary Figure 1C). The new dialogue window allows determining the parameters for the three steps of the analysis (Supplementary Figure 1D), which are detection, tracking, and track analysis.</li>
	<li>In step 1 choose &#34;Settings&#34; and from a drop-down menu select &#34;Comet Detection&#34; as a detection method (Supplementary Figure 2B).
	<ol>
		<li>From the new dialogue window define the parameters for the difference of Gaussians filter and the watershed segmentation as follows (Supplementary Figure 2C): Mask process to be used for the detection = None; Low-pass Gaussian standard deviation = 1 pixel; High-pass Gaussian standard deviation = 3 pixels; Minimum threshold = 3 standard deviations; Threshold step size = 0.25 standard deviations. Select &#34;Apply Settings to All Movies&#34; and &#34;Apply&#34;.</li>
	</ol>
	</li>
	<li>In step 2, the parameters for linking, gap closing, merging and splitting, and Kalman filter functions are defined in three steps as highlighted in pink, green, and blue, accordingly (Supplementary Figure 3B). For these steps, select the &#34;Microtubule Plus-end Dynamics&#34; and from the &#34;Setting&#34; option define the values as indicated in Supplementary Figure 3C&#8211;E, respectively.
	<ol>
		<li>For problems with dimensionality, choose &#34;2&#34; from the drop-down menu. Use Maximum Gap to Close = 5 frames; Minimum Length of Track Segments from First Step = 3 frames. As before, select &#34;Apply Settings to All Movies&#34; and click on &#34;Apply&#34;.</li>
	</ol>
	</li>
	<li>In step 3 of the analysis, the detected MT tracks are classified (Supplementary Figure 4). As a track analysis method, choose &#34;Microtubule Dynamics Classification&#34; and define the parameters through the &#34;Setting&#34; button as indicated in Supplementary Figure 4B,C. After that, choose the &#34;Apply Settings to All Movies&#34; box and click on &#34;Apply&#34;.</li>
	<li>Once all the parameters are defined, from the &#34;Control Panel&#8211;U-Track&#34; window (Supplementary Figure 1D) select the &#34;Apply Check/Uncheck to All Movies&#34; and &#34;Run All Movies&#34; boxes and press &#34;Run&#34;. This will initiate the MT analysis of the time-lapse series.</li>
	<li>Once the movie processing is completed, a message &#34;Your movie(s) have been processed successfully&#34; is displayed. Press &#34;Ok&#34;, then &#34;Save&#34;.</li>
	<li>Now it is safe to quit the numerical-analysis software. The results from the movie processing are stored in subfolder structures as m-files in the folder where the raw files are stored.</li>
</ol>
5. Statistical Analysis of the MT Dynamics
<ol>
	<li>Import the m-files into a preferred statistic analysis program. 
	NOTE: In our case, we first import the files in a standard spreadsheet to make them readable. The m-files contain statistical information (median, mean, and standard deviation) on different parameters (e.g., growth speed, MT dynamicity). The detailed list of the parameters is given in the technical report provided with the previous version of the software, plusTipTracker50,52. The generated m-files can also be imported into other data processing software.</li>
	<li>Choose the &#34;growth speed mean&#34; parameter and import it into a table for statistics and display. Enter the information on other parameters, (e.g., &#34;dynamicity&#34;) either in a new table or in a new column of the same grouped table and plot.</li>
</ol>

The authors have nothing to disclose.

Here, we describe a modification of a method first established by Ertych et al.44. Along with several other modifications, we combine this technique of MT dynamics analysis with dual spinning disk confocal imaging. The use of the dual spinning disk improves the resolution of growing MTs while reducing phototoxicity36. We further reduce the photobleaching and laser light-induced damage of the cells by switching to a longer wavelength fluorescent reporter. The tdTomato fluore...

Microtubules (MTs) are highly dynamic structures found in virtually all eukaryotic cells and in some bacteria1. Together with actin and intermediate filaments, they sculpt the cytoskeleton2,3. Cell division4, molecule transport5, flagellar beating6, the sensation of the surrounding environment through primary cilium7, hearing (kinocilium)8,9, embryogenesis10,11,12, invasion and metastasis13,14, and even memory formation15,16,17,18, and many other processes primarily rely on MTs. Participation of MTs in all these events would be impossible without their remarkable ability to rapidly switch between growth (polymerization) and shrinkage (depolymerization). This property is described as dynamic instability19. MT dynamicity is altered in many pathological conditions20,21,22. Hence, determining the nature of this property can help to understand disease mechanisms and subsequently their treatment.
A long list of methods has been developed for MT dynamics analysis, most of which are based on imaging techniques23. Initially, wide field light microscopes were used for observing the formation of tubulin polymers in vitro24. The discovery of end-binding (EB)-proteins that collect at MT plus-ends and the development of methods to fluorescently label proteins made it possible to observe the behavior of MTs directly in living cells with wide field and confocal fluorescence microscopes25,26,27. One EB-protein is end-binding protein 3 (EB3)28; by overexpressing and tracking EB3 fused to a fluorescent protein, MT plus-end assembly rates can be determined29,30.
Confocal laser scanning fluorescence microscopy (CLSM) is frequently used to follow MT dynamics. However, this imaging technique poses a high risk of phototoxicity and photobleaching, two undesirable processes for live cell and dim sample imaging31. In order to obtain a better signal-to-noise ratio, the laser power and the exposure duration should be high enough while not damaging the samples, and this requires sacrificing resolution in exchange for speed. A suitable alternative to CLSM is spinning disk microscopy32. This imaging modality is based on the use of a Nipkow disk33, which consists of a moving disk bearing an array of pinholes, and works equivalently to many CLS microscopes imaging the same sample simultaneously34. Therefore, the light from the laser will illuminate several regions in the sample simultaneously but retain the confocal nature. The Nipkow disk, therefore, allows obtaining images similar to CLSM but faster and using less laser power. The Nipkow disk was further improved by Yokogawa Electric, which introduced a second disk with an array of microlenses on it that individually direct light into a respective pinhole, further reducing phototoxicity and photobleaching35. Thus, spinning disk laser scanning microscopy became a method of choice for live cell imaging, and it makes it possible to obtain images with high signal-to-noise ratio at a high speed31,36, which is crucial for resolving signals such as those from the fast-growing MT ends.
MT dynamics differ temporarily. For example, the mitotic MTs are more dynamic than the interphase ones37,38. Similarly, differences in the growth rate and shrinkage have been observed even within the same cell cycle phase, such as mitosis39,40. Therefore, to avoid false data collection, the measurement of MT dynamics should be limited to a narrow time-window during the cell cycle. For example, measurement of MT dynamics in prometaphase can be achieved by treating the cells with dimethyl&#173;enastron (DME), a monastrol analogue that inhibits the motor kinesin Eg541 and prevents the formation of the bipolar mitotic spindle42. Inhibition of cells at prometaphase with Eg5 inhibitor DME and other monastrol derivatives does not affect the MT dynamics43,44,45, which makes DME a useful tool for studying MT dynamics both in fixed and live cells44.
Here we combine the method of MT dynamics analysis in prometaphase cells described by Ertych et al.44 with dual spinning disk imaging. This method allows measurement of the MT dynamics in prometaphase cells collected from a single focal plane with a higher imaging rate, yet without photobleaching and minimal phototoxicity. Furthermore, as a fluorescent reporter, we use tandem dimer Tomato fluorescent protein (tdTomato) which has improved brightness and photostability in comparison to the green fluorescent protein (EGFP) and is excited with lower energy light46. Therefore, tdTomato requires less laser power for excitation and is less phototoxic. Altogether, we further improve the method by reducing the phototoxicity and improving the resolution and postprocessing required for the MT dynamics analysis. Additionally, we create a basis for future modifications of the method by combining it with other synchronization techniques.

<table><tbody><tr><td>Dimethylenastron</td><td>Merck</td><td>324622</td><td></td></tr><tr><td>DMEM w/o phenol red</td><td>Gibco</td><td>31053-28</td><td></td></tr><tr><td>DPBS</td><td>Gibco</td><td>14190-094</td><td></td></tr><tr><td>Fetal bovine serum</td><td>Biochrom</td><td>S0415</td><td></td></tr><tr><td>Fibronectin Bovine Plasma</td><td>Merck</td><td>F4759</td><td>Sterile powder</td></tr><tr><td>GlutaMAX</td><td>Gibco</td><td>35050-038</td><td>Stable glutamine substitutive</td></tr><tr><td>jetPRIME</td><td>Polyplus</td><td>114-15</td><td></td></tr><tr><td>EB3-TdTomato</td><td>Addgene</td><td>plasmid #50708</td><td></td></tr><tr><td>RPMI 1640</td><td>Gibco</td><td>61870-010</td><td></td></tr><tr><td>Trypan Blue</td><td>Merck</td><td>T8154-20ML</td><td></td></tr><tr><td>Trypsin/EDTA solution</td><td>Biochrom</td><td>L2143</td><td>0.05%/0.02 % w/o calcium and magnesium</td></tr><tr><td>&amp;micro;-slide</td><td>Ibidi</td><td>80426</td><td>4-well slide with #1.5 coverslip</td></tr><tr><td>Eclipse Ti Inverted microscope</td><td>Nikon</td><td>NA</td><td></td></tr><tr><td>Objective</td><td>Nikon</td><td>MRD01991</td><td>CFI Apo TIRF 100xC Oil</td></tr><tr><td>ACAL Laser Excahnger</td><td>Nikon</td><td></td><td>Laser box. 405, 458, 488, 514, 561 and 647 nm</td></tr><tr><td>Spinning disk module</td><td>Andor</td><td>CSU-W</td><td></td></tr><tr><td>Camera</td><td>Andor</td><td>iXon Ultra 888</td><td></td></tr><tr><td>Environmental Chamber</td><td>Okolab</td><td></td><td>Dark chamber equipped with CO&lt;sub&gt;2&lt;/sub&gt; supply, temperature control and humidifier</td></tr><tr><td>HeLa Cells</td><td>DSMZ</td><td>ACC-57</td><td></td></tr><tr><td>NIS Elements v4</td><td>Nikon</td><td></td><td>Spinning disk microscope. Acquisition Software</td></tr><tr><td>MATLAB</td><td>Mathworks</td><td></td><td>Computing environment</td></tr><tr><td>Prism 8</td><td>GraphPad</td><td></td><td>Statistical analysis and display software</td></tr></tbody></table>

measurement of microtubule dynamics by spinning disk microscopy in monopolar mitotic spindles

We describe a modification of an established method to determine microtubule dynamics in living cells. The protocol is based on the expression of a genetically encoded marker for the positive ends of microtubules (EB3 labelled with tdTomato fluorescent protein) and high-speed, high-resolution, live-cell imaging using spinning disk confocal microscopy. Cell cycle synchronization and increased density of microtubules are achieved by inhibiting centrosomal separation in mitotic cells, and analysis of growth is performed using open-source U-Track software. The use of a bright and red-shifted fluorescent protein, in combination with the lower laser power and reduced exposure time required for spinning disk microscopy reduce phototoxicity and the probability of light-induced artifacts. This allows for imaging a larger number of cells in the same preparation while maintaining the cells in a growth medium under standard culture conditions. Because the analysis is performed in a supervised automatic fashion, the results are statistically robust and reproducible.

Following the given protocol outlined in Figure 1A, the pEB3-tdTomato plasmid was transiently expressed in asynchronously growing HeLa cells. The cells were synchronized 48 h after the transfection at prometaphase through DME treatment (Figure 1B). This step ensured that the measurement of MT dynamics was always done at the same phase of the cell cycle. The time-lapse movies were further processed and analyzed with U-Track v2.2.0 as described in its supplementar...

Watch this Scientific Journal Video about Measurement of Microtubule Dynamics by Spinning Disk Microscopy in Monopolar Mitotic Spindles at JoVE.com

Measurement of Microtubule Dynamics by Spinning Disk Microscopy in Monopolar Mitotic Spindles

Here we present a robust and detailed method of microtubule dynamics analysis in cells synchronized in prometaphase using live-cell spinning disk confocal microscopy and MATLAB-based image processing.

We describe a modification of an established method to determine microtubule dynamics in living cells. The protocol is based on the expression of a ...

measurement-microtubule-dynamics-spinning-disk-microscopy-monopolar

Universidad de Cartagena UDEC

Research

JoVE Journal

Biology

6.1K Views.  Max-Planck Institute of Experimental Medicine. Here we present a robust and detailed method of microtubule dynamics analysis in cells synchronized in prometaphase using live-cell spinning disk confocal microscopy and MATLAB-based image processing.

Video: Measurement of Microtubule Dynamics by Spinning Disk Microscopy in Monopolar Mitotic Spindles

Simultaneous Visualization of the Dynamics of Crosslinked and Single Microtubules In Vitro by TIRF Microscopy

Microtubules are polymers of &#945;&#946;-tubulin heterodimers that organize into distinct structures in cells. Microtubule-based architectures and networks often contain subsets of microtubule arrays that differ in their dynamic properties. For example, in dividing cells, stable bundles of crosslinked microtubules coexist in close proximity to dynamic non-crosslinked microtubules. TIRF-microscopy-based in vitro reconstitution studies enable the simultaneous visualization of the dynamics of these different microtubule arrays. In this assay, an imaging chamber is assembled with surface-immobilized microtubules, which are either present as single filaments or organized into crosslinked&#160;bundles. Introduction of tubulin, nucleotides, and protein regulators allows direct visualization of associated proteins and of dynamic properties of single and crosslinked microtubules. Furthermore, changes that occur as dynamic single microtubules organize into bundles can be monitored in real-time. The method described here allows for a systematic evaluation of the activity and localization of individual proteins, as well as synergistic effects of protein regulators on two different microtubule subsets under identical experimental conditions, thereby providing mechanistic insights that are inaccessible by other methods.

Simultaneous Visualization of the Dynamics of Crosslinked and Single Microtubules In Vitro by TIRF Microscopy

Here, a TIRF microscopy-based in vitro reconstitution assay is presented to simultaneously quantify and compare the dynamics of two microtubule populations. A method is described to simultaneously view the collective activity of multiple microtubule-associated proteins on crosslinked microtubule bundles and single microtubules.

Microtubules are polymers of &#945;&#946;-tubulin heterodimers that organize into distinct structures in cells. Microtubule-based architectures and ...

Biochemistry

Microtubule Plus-End Dynamics Visualization in Huntington's Disease Model based on Human Primary Skin Fibroblasts

Transfection with a fluorescently labeled marker protein of interest in combination with time-lapse video microscopy is a classic method of studying the dynamic properties of the cytoskeleton. This protocol offers a technique for human primary fibroblast transfection, which can be difficult because of the specifics of primary cell cultivation conditions. Additionally, cytoskeleton dynamic property maintenance requires a low level of transfection to obtain a good signal-to-noise ratio without causing microtubule stabilization. It is important to take measures to protect the cells from light-induced stress and fluorescent dye fading. In the course of our work, we tested different transfection methods and protocols as well as different vectors to select the best combination of conditions suitable for human primary fibroblast studies. We analyzed the resulting time-lapse videos and calculated microtubule dynamics using ImageJ. The dynamics of microtubules' plus-ends in the different cell parts are not similar, so we divided the analysis into subgroups - the centrosome region, the lamella, and the tail of fibroblasts. Notably, this protocol can be used for in vitro analysis of cytoskeleton dynamics in patient samples, enabling the next step towards understanding the dynamics of the various disease development.

This protocol is dedicated to the microtubule plus-end visualization by EB3 protein transfection to study their dynamic properties in primary cell culture. The protocol was implemented on human primary skin fibroblasts obtained from Huntington's disease patients.

Transfection with a fluorescently labeled marker protein of interest in combination with time-lapse video microscopy is a classic method of studying the ...

Bioengineering

Quantifying Cytoskeleton Dynamics Using Differential Dynamic Microscopy

Cells can crawl, self-heal, and tune their stiffness due to their remarkably dynamic cytoskeleton. As such, reconstituting networks of cytoskeletal biopolymers may lead to a host of active and adaptable materials. However, engineering such materials with precisely tuned properties requires measuring how the dynamics depend on the network composition and synthesis methods. Quantifying such dynamics is challenged by variations across the time, space, and formulation space of composite networks. The protocol here describes how the Fourier analysis technique, differential dynamic microscopy (DDM), can quantify the dynamics of biopolymer networks and is particularly well suited for studies of cytoskeleton networks. DDM works on time sequences of images acquired using a range of microscopy modalities, including laser-scanning confocal, widefield fluorescence, and brightfield imaging. From such image sequences, one can extract characteristic decorrelation times of density fluctuations across a span of wave vectors. A user-friendly, open-source Python package to perform DDM analysis is also developed. With this package, one can measure the dynamics of labeled cytoskeleton components or of embedded tracer particles, as demonstrated here with data of intermediate filament (vimentin) networks and active actin-microtubule networks. Users with no prior programming or image processing experience will be able to perform DDM using this software package and associated documentation.

Differential dynamic microscopy (DDM) combines features of dynamic light scattering and microscopy. Here, the process of using DDM to characterize reconstituted cytoskeleton networks by quantifying the subdiffusive and caged dynamics of particles in vimentin networks and the ballistic motion of active myosin-driven actin-microtubule composites is presented.

Cells can crawl, self-heal, and tune their stiffness due to their remarkably dynamic cytoskeleton. As such, reconstituting networks of cytoskeletal ...

3D Particle Tracking for Noninvasive In Vivo Analysis of Synaptic Microtubule Dynamics in Dendrites and Neuromuscular Junctions of Drosophila

Microtubules (MTs) play critical roles in neuronal development, but many questions remain about the molecular mechanisms of their regulation and function. Furthermore, despite progress in understanding postsynaptic MTs, much less is known about the contributions of presynaptic MTs to neuronal morphogenesis. In particular, studies of in vivo MT dynamics in Drosophila sensory dendrites yielded significant insights into polymer-level behavior. However, the technical and analytical challenges associated with live imaging of the fly neuromuscular junction (NMJ) have limited comparable studies of presynaptic MT dynamics. Moreover, while there are many highly effective software strategies for automated analysis of MT dynamics in vitro and ex vivo, in vivo data often necessitate significant operator input or entirely manual analysis due to inherently inferior signal-to-noise ratio in images and complex cellular morphology. &#160;To address this, this study optimized a new software platform for automated and unbiased in vivo particle detection. Multiparametric analysis of live time-lapse confocal images of EB1-GFP labeled MTs was performed in both dendrites and the NMJ of Drosophila larvae and found striking differences in MT behaviors. MT dynamics were furthermore analyzed following knockdown of the MT-associated protein (MAP) dTACC, a key regulator of Drosophila synapse development, and identified statistically significant changes in MT dynamics compared to wild type. These results demonstrate that this novel strategy for the automated multiparametric analysis of both pre- and postsynaptic MT dynamics at the polymer-level significantly reduces human-in-the-loop criteria. The study furthermore shows the utility of this method in detecting distinct MT behaviors upon dTACC-knockdown, indicating a possible future application for functional screens of factors that regulate MT dynamics in vivo. Future applications of this method may also focus on elucidating cell type and/or compartment-specific MT behaviors, and multicolor correlative imaging of EB1-GFP with other cellular and subcellular markers of interest.&#160;

This study presents a noninvasive intravital neuronal imaging strategy combined with a new software strategy to achieve automated, unbiased tracking and analysis of in vivo microtubule (MT) plus-end dynamics in the sensory dendrites and the neuromuscular junctions of Drosophila.

Microtubules (MTs) play critical roles in neuronal development, but many questions remain about the molecular mechanisms of their regulation and function. ...

Neuroscience

Preparation of Segmented Microtubules to Study Motions Driven by the Disassembling Microtubule Ends

Microtubule depolymerization can provide force to transport different protein complexes and protein-coated beads in vitro. The underlying mechanisms are thought to play a vital role in the microtubule-dependent chromosome motions during cell division, but the relevant proteins and their exact roles are ill-defined. Thus, there is a growing need to develop assays with which to study such motility in vitro using purified components and defined biochemical milieu. Microtubules, however, are inherently unstable polymers; their switching between growth and shortening is stochastic and difficult to control. The protocols we describe here take advantage of the segmented microtubules that are made with the photoablatable stabilizing caps. Depolymerization of such segmented microtubules can be triggered with high temporal and spatial resolution, thereby assisting studies of motility at the disassembling microtubule ends. This technique can be used to carry out a quantitative analysis of the number of molecules in the fluorescently-labeled protein complexes, which move processively with dynamic microtubule ends. To optimize a signal-to-noise ratio in this and other quantitative fluorescent assays, coverslips should be treated to reduce nonspecific absorption of soluble fluorescently-labeled proteins. Detailed protocols are provided to take into account the unevenness of fluorescent illumination, and determine the intensity of a single fluorophore using equidistant Gaussian fit. Finally, we describe the use of segmented microtubules to study microtubule-dependent motions of the protein-coated microbeads, providing insights into the ability of different motor and nonmotor proteins to couple microtubule depolymerization to processive cargo motion.

Microtubules are inherently unstable polymers, and their switching between growth and shortening is stochastic and difficult to control. Here we describe protocols using segmented microtubules with photoablatable stabilizing caps. Depolymerization of segmented microtubules can be triggered with high temporal and spatial resolution, thereby assisting analysis of motions with the disassembling microtubule ends.

Microtubule depolymerization can provide force to transport different protein complexes and protein-coated beads in vitro. The underlying mechanisms are ...

High-resolution Time-lapse Imaging and Automated Analysis of Microtubule Dynamics in Living Human Umbilical Vein Endothelial Cells

The physiological process by which new vasculature forms from existing vasculature requires specific signaling events that trigger morphological changes within individual endothelial cells (ECs). These processes are critical for homeostatic maintenance such as wound healing, and are also crucial in promoting tumor growth and metastasis. EC morphology is defined by the organization of the cytoskeleton, a tightly regulated system of actin and microtubule (MT) dynamics that is known to control EC branching, polarity and directional migration, essential components of angiogenesis. To study MT dynamics, we used high-resolution fluorescence microscopy coupled with computational image analysis of fluorescently-labeled MT plus-ends to investigate MT growth dynamics and the regulation of EC branching morphology and directional migration. Time-lapse imaging of living Human Umbilical Vein Endothelial Cells (HUVECs) was performed following transfection with fluorescently-labeled MT End Binding protein 3 (EB3) and Mitotic Centromere Associated Kinesin (MCAK)-specific cDNA constructs to evaluate effects on MT dynamics. PlusTipTracker software was used to track EB3-labeled MT plus ends in order to measure MT growth speeds and MT growth lifetimes in time-lapse images. This methodology allows for the study of MT dynamics and the identification of how localized regulation of MT dynamics within sub-cellular regions contributes to the angiogenic processes of EC branching and migration.

Protocols for Human Umbilical Vein Endothelial Cell (HUVEC) culture, transient transfection of fluorescently-labeled markers of microtubule growth, live-cell imaging and automated analysis of interphase microtubule growth dynamics are detailed.

The physiological process by which new vasculature forms from existing vasculature requires specific signaling events that trigger morphological changes ...

Reconstitution of Basic Mitotic Spindles in Spherical Emulsion Droplets

Mitotic spindle assembly, positioning and orientation depend on the combined forces generated by microtubule dynamics, microtubule motor proteins and cross-linkers. Growing microtubules can generate pushing forces, while depolymerizing microtubules can convert the energy from microtubule shrinkage into pulling forces, when attached, for example, to cortical dynein or chromosomes. In addition, motor proteins and diffusible cross-linkers within the spindle contribute to spindle architecture by connecting and sliding anti-parallel microtubules. In vivo, it has proven difficult to unravel the relative contribution of individual players to the overall balance of forces. Here we present the methods that we recently developed in our efforts to reconstitute basic mitotic spindles bottom-up in vitro. Using microfluidic techniques, centrosomes and tubulin are encapsulated in water-in-oil emulsion droplets, leading to the formation of geometrically confined (double) microtubule asters. By additionally introducing cortically anchored dynein, plus-end directed microtubule motors and diffusible cross-linkers, this system is used to reconstitute spindle-like structures. The methods presented here provide a starting point for reconstitution of more complete mitotic spindles, allowing for a detailed study of the contribution of each individual component, and for obtaining an integrated quantitative view of the force-balance within the mitotic spindle.

The assembly and positioning of the mitotic spindle depend on the combined forces generated by microtubule dynamics, motor proteins and cross-linkers. Here we present our recently developed methods in which the geometrical confinement of spherical emulsion droplets is used for the bottom-up reconstitution of basic mitotic spindles.

Mitotic spindle assembly, positioning and orientation depend on the combined forces generated by microtubule dynamics, microtubule motor proteins and ...

High-resolution Imaging and Analysis of Individual Astral Microtubule Dynamics in Budding Yeast

Dynamic microtubules are fundamental to many cellular processes, and accurate measurements of microtubule dynamics can provide insight into how cells regulate these processes and how genetic mutations impact regulation. The quantification of microtubule dynamics in metazoan models has a number of associated challenges, including a high microtubule density and limitations on genetic manipulations. In contrast, the budding yeast model offers advantages that overcome these challenges. This protocol describes a method to measure the dynamics of single microtubules in living yeast cells. Cells expressing fluorescently tagged tubulin are adhered to assembled slide chambers, allowing for stable time-lapse image acquisition. A detailed guide for high-speed, four-dimensional image acquisition is also provided, as well as a protocol for quantifying the properties of dynamic microtubules in confocal image stacks. This method, combined with conventional yeast genetics, provides an approach that is uniquely suited for quantitatively assessing the effects of microtubule regulators or mutations that alter the activity of tubulin subunits.

Budding yeast is an advantageous model for studying microtubule dynamics in vivo due to its powerful genetics and the simplicity of its microtubule cytoskeleton. The following protocol describes how to transform and culture yeast cells, acquire confocal microscopy images, and quantitatively analyze microtubule dynamics in living yeast cells.

Dynamic microtubules are fundamental to many cellular processes, and accurate measurements of microtubule dynamics can provide insight into how cells ...

Live Cell Imaging to Assess the Dynamics of Metaphase Timing and Cell Fate Following Mitotic Spindle Perturbations

Live cell time-lapse imaging is an important tool in cell biology that provides insight into cellular processes that might otherwise be overlooked, misunderstood, or misinterpreted by the fixed-cell analysis. While the fixed cell imaging and analysis is robust and sufficient to observe cellular steady-state, it can be limited in defining a temporal order of events at the cellular level and is ill-equipped to assess the transient nature of dynamic processes including mitotic progression. In contrast, live cell imaging is an eloquent tool that can be used to observe cellular processes at the single-cell level over time and has the capacity to capture the dynamics of processes that would otherwise be poorly represented in fixed cell imaging. Here we describe an approach to generate cells carrying fluorescently labeled markers of chromatin and microtubules and their use in live cell imaging approaches to monitor metaphase chromosome alignment and mitotic exit. We describe imaging-based techniques to assess the dynamics of spindle formation and mitotic progression, including the identification of cells at various stages in mitosis, identification&#160;and tracking of mitotic defects, and analysis of spindle dynamics and mitotic cell fate following the treatment with mitotic inhibitors.

Here we present a protocol to assess the dynamics of spindle formation and mitotic progression. Our application of time-lapse imaging enables the user to identify cells at various stages of mitosis, track and identify mitotic defects, and analyze spindle dynamics and mitotic cell fate upon exposure to anti-mitotic drugs.

Live cell time-lapse imaging is an important tool in cell biology that provides insight into cellular processes that might otherwise be overlooked, ...

Directly Measuring Forces Within Reconstituted Active Microtubule Bundles

Microtubule networks are employed in cells to accomplish a wide range of tasks, ranging from acting as tracks for vesicle transport to working as specialized arrays during mitosis to regulate chromosome segregation. Proteins that interact with microtubules include motors such as kinesins and dynein, which can generate active forces and directional motion, as well as non-motor proteins that crosslink filaments into higher-order networks or regulate filament dynamics. To date, biophysical studies of microtubule-associated proteins have overwhelmingly focused on the role of single motor proteins needed for vesicle transport, and significant progress has been made in elucidating the force-generating properties and mechanochemical regulation of kinesins and dyneins. However, for processes in which microtubules act both as cargo and track, such as during filament sliding within the mitotic spindle, much less is understood about the biophysical regulation of ensembles of the crosslinking proteins involved. Here, we detail our methodology for directly probing force generation and response within crosslinked microtubule minimal networks reconstituted from purified microtubules and mitotic proteins. Microtubule pairs are crosslinked by proteins of interest, one microtubule is immobilized to a microscope coverslip, and the second microtubule is manipulated by an optical trap. Simultaneous total internal reflection fluorescence microscopy allows for multichannel visualization of all the components of this microtubule network as the filaments slide apart to generate force. We also demonstrate how these techniques can be used to probe pushing forces exerted by kinesin-5 ensembles and how viscous braking forces arise between sliding microtubule pairs crosslinked by the mitotic MAP PRC1. These assays provide insights into the mechanisms of spindle assembly and function and can be more broadly adapted to study dense microtubule network mechanics in diverse contexts, such as the axon and dendrites of neurons and polar epithelial cells.

Here, we present a protocol for reconstituting microtubule bundles in vitro and directly quantifying the forces exerted within them using simultaneous optical trapping and total internal reflection fluorescence microscopy. This assay allows for nanoscale-level measurement of the forces and displacements generated by protein ensembles within active microtubule networks.

Measurement of Microtubule Dynamics by Spinning Disk Microscopy in Monopolar Mitotic Spindles

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Reconstitution of Basic Mitotic Spindles in Spherical Emulsion Droplets

High-resolution Imaging and Analysis of Individual Astral Microtubule Dynamics in Budding Yeast

Live Cell Imaging to Assess the Dynamics of Metaphase Timing and Cell Fate Following Mitotic Spindle Perturbations

Directly Measuring Forces Within Reconstituted Active Microtubule Bundles