The authors wish to acknowledge support from R21 AG067436 (to JP and SF), T32 AG057464 (to ET), and Rensselaer Polytechnic Institute School of Science Startup Funds (to SF).

The authors have nothing to disclose.

Microtubule networks are employed by myriad cell types to accomplish a wide range of tasks that are fundamentally mechanical in nature. In order to describe how cells function in both healthy and disease states, it is critical to understand how these micron-scale networks are organized and regulated by the nanometer-sized proteins that collectively build them. Biophysical tools such as optical tweezers are well suited to probing the mechanochemistry of key proteins at this scale. Reflecting the diversity of microtubule n...

Cells employ microtubule networks to perform a wide variety of mechanical tasks, ranging from vesicle transport1,2,3 to chromosome segregation during mitosis4,5,6. Many of the proteins that interact with microtubules, such as the molecular motor proteins kinesin and dynein, generate forces and are regulated by mechanical loads. To better understand how these critical molecules function, researchers have employed single-molecule biophysical methods, such as optical trapping and TIRF microscopy, to directly monitor critical parameters such as unloaded stepping rates, processivity, and force-velocity relationships for individual proteins. The most commonly used experimental geometry has been to attach motor proteins directly to trapping beads whose spherical geometry and size mimic vesicles undergoing motor-driven transport. Numerous kinesins, including kinesin-17,8,9, kinesin-210,11,12, kinesin-313,14,15,16 kinesin-517,18, kinesin-819,20, as well as dynein and dynein complexes21,22,23,24,25, have been studied with these methods.
In many cellular processes, however, motor and non-motor proteins use microtubules both as track and cargo26,27. Moreover, in these scenarios where microtubule filaments are crosslinked into higher-order bundles, these proteins function as ensembles rather than single units. For example, within dividing somatic cells, dense filament networks self-organize to build the mitotic spindle apparatus28,29,30. The interpolar spindle microtubule network is highly dynamic and is largely arranged with minus-ends pointing toward the spindle poles and plus-ends overlapping near the spindle equator. Filaments within the spindle are crosslinked by motor proteins such as kinesin-531,32,33, kinesin-1234,35,36, and kinesin-1437,38,39, or by non-motor proteins such as PRC140,41,42,43 or NuMA44,45,46. They frequently move or experience mechanical stress during processes such as poleward flux or while coordinating chromosome centering during metaphase or chromosome segregation during anaphase47,48,49,50,51,52. The integrity of the micron-scale spindle apparatus through mitosis, therefore, relies on a carefully regulated balance of pushing and pulling forces generated and sustained by this network of interacting filaments. However, the tools needed to probe this mechanical regulation and explain how protein ensembles work in concert to coordinate microtubule motions and produce the forces needed to properly assemble the spindle have only recently been developed, and we are just beginning to understand the biophysical rules that define dynamic microtubule networks.
The goal of this manuscript is to demonstrate the steps required to reconstitute crosslinked microtubule pairs in vitro, immobilize these bundles in a microscopy chamber that allows for simultaneous fluorescence visualization of both the microtubules and crosslinking proteins and nanoscale force measurement, and process these data robustly. We detail the steps needed to stably polymerize fluorescence-labeled microtubules, prepare microscope coverslips for attachment, prepare polystyrene beads for optical trapping experiments, and assemble crosslinked filament networks that preserve their in vivo functionality while allowing for direct biophysical manipulation.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>10W Ytterbium Fiber Laser, 1064nm</td>
			<td>IPG Photonics</td>
			<td>YLR-10-1064-LP</td>
			<td></td>
		</tr>
		<tr>
			<td>405/488/561/640nm Laser Quad Band Set for TIRF applications</td>
			<td>Chroma</td>
			<td>TRF89901v2</td>
			<td></td>
		</tr>
		<tr>
			<td>6x His Tag Antibody, Biotin Conjugate</td>
			<td>Invitrogen</td>
			<td>#MA1-21315-BTIN</td>
			<td></td>
		</tr>
		<tr>
			<td>Acetone, HPLC grade</td>
			<td>Fisher Scientific</td>
			<td>18-608-395</td>
			<td></td>
		</tr>
		<tr>
			<td>Alpha casein from bovine milk</td>
			<td>Sigma</td>
			<td>1002484390</td>
			<td></td>
		</tr>
		<tr>
			<td>ATP</td>
			<td>Fisher Scientific</td>
			<td>BP413-25</td>
			<td></td>
		</tr>
		<tr>
			<td>Benzonase</td>
			<td>Novagen</td>
			<td>70746-3</td>
			<td></td>
		</tr>
		<tr>
			<td>Biotin-PEG-SVA-5000</td>
			<td>Laysan Bio, Inc.</td>
			<td>NC0479433</td>
			<td></td>
		</tr>
		<tr>
			<td>BL21 (DE3) Rosetta Cells</td>
			<td>Millipore Sigma</td>
			<td>71-400-3</td>
			<td></td>
		</tr>
		<tr>
			<td>Catalase</td>
			<td>MP Biomedicals LLC</td>
			<td>190311</td>
			<td></td>
		</tr>
		<tr>
			<td>CFI Apo 100X/1.49NA oil immersion TIRF objective</td>
			<td>Nikon</td>
			<td>N/A</td>
			<td></td>
		</tr>
		<tr>
			<td>Chloramphenicol</td>
			<td>ACROS Organics</td>
			<td>227920250</td>
			<td></td>
		</tr>
		<tr>
			<td>Coverslip Mini-Rack, for 8 coverslips</td>
			<td>Fisher Scientific</td>
			<td>C14784</td>
			<td></td>
		</tr>
		<tr>
			<td>Delicate Task Wipers</td>
			<td>Kimberly-Clark</td>
			<td>34120</td>
			<td></td>
		</tr>
		<tr>
			<td>Dextrose Anhydrous</td>
			<td>Fisher Scientific</td>
			<td>BP3501</td>
			<td></td>
		</tr>
		<tr>
			<td>D-Sucrose</td>
			<td>Fisher Scientific</td>
			<td>BP220-1</td>
			<td></td>
		</tr>
		<tr>
			<td>DTT</td>
			<td>Fisher Scientific</td>
			<td>BP172-25</td>
			<td></td>
		</tr>
		<tr>
			<td>Ecoline Immersion Thermostat E100 with 003 Bath</td>
			<td>LAUDA-Brinkmann</td>
			<td>27709</td>
			<td></td>
		</tr>
		<tr>
			<td>EDTA</td>
			<td>Fisher Scientific</td>
			<td>BP118-500</td>
			<td></td>
		</tr>
		<tr>
			<td>EGTA</td>
			<td>Millipore Corporation</td>
			<td>32462-25GM</td>
			<td></td>
		</tr>
		<tr>
			<td>FIJI / Image J</td>
			<td>https://fiji.sc/</td>
			<td>N/A</td>
			<td></td>
		</tr>
		<tr>
			<td>Frosted Microscope Slides</td>
			<td>Corning</td>
			<td>12-553-10</td>
			<td>75mmx25mm, with thickness of 0.9-1.1mm</td>
		</tr>
		<tr>
			<td>Glucose Oxidase</td>
			<td>MP Biomedicals LLC</td>
			<td>195196</td>
			<td>Type VII, without added oxygen</td>
		</tr>
		<tr>
			<td>GMPCPP</td>
			<td>Jena Biosciences</td>
			<td>JBS-NU-405S</td>
			<td>Can be stored for several months at -20 &#176;C and up to a year at -80 &#176;C</td>
		</tr>
		<tr>
			<td>Gold Seal-Cover Glass</td>
			<td>Thermo Scientific</td>
			<td>3405</td>
			<td></td>
		</tr>
		<tr>
			<td>HEPES</td>
			<td>Fisher Scientific</td>
			<td>BP310-500</td>
			<td></td>
		</tr>
		<tr>
			<td>Imidazole</td>
			<td>Fisher Scientific</td>
			<td>03196-500</td>
			<td></td>
		</tr>
		<tr>
			<td>IPTG</td>
			<td>Fisher Scientific</td>
			<td>BP1755-10</td>
			<td></td>
		</tr>
		<tr>
			<td>Laboratory dessicator</td>
			<td>Bel-Art</td>
			<td>999320237</td>
			<td>190mm plate size</td>
		</tr>
		<tr>
			<td>Kanamycin Sulfate</td>
			<td>Fischer Scientific</td>
			<td>BP906-5</td>
			<td></td>
		</tr>
		<tr>
			<td>KIF5A K439 (aa:1-439)-6His</td>
			<td>Gilbert Lab, RPI</td>
			<td>N/A</td>
			<td>doi.org/10.1074/jbc.RA118.002182</td>
		</tr>
		<tr>
			<td>Kimwipe</td>
			<td>Kimberley Clark</td>
			<td>Z188956</td>
			<td>lint-free tissue</td>
		</tr>
		<tr>
			<td>Immersion Oil, Type B</td>
			<td>Cargille</td>
			<td>16484</td>
			<td></td>
		</tr>
		<tr>
			<td>Lens Tissue</td>
			<td>ThorLabs</td>
			<td>MC-5</td>
			<td></td>
		</tr>
		<tr>
			<td>LuNA Laser launch (4 channel: 405, 488, 561, 640nm)</td>
			<td>Nikon</td>
			<td>N/A</td>
			<td></td>
		</tr>
		<tr>
			<td>Lysozyme</td>
			<td>MP Biomedicals LLC</td>
			<td>100834</td>
			<td></td>
		</tr>
		<tr>
			<td>Magnesium Acetate Tetrahydrate</td>
			<td>Fisher Scientific</td>
			<td>BP215-500</td>
			<td></td>
		</tr>
		<tr>
			<td>Microfuge 18</td>
			<td>Beckman Coulter</td>
			<td>367160</td>
			<td></td>
		</tr>
		<tr>
			<td>MPEG-SVA MW-5000</td>
			<td>Laysan Bio, Inc.</td>
			<td>NC0107576</td>
			<td></td>
		</tr>
		<tr>
			<td>Neutravadin</td>
			<td>Invitrogen</td>
			<td>PI31000</td>
			<td></td>
		</tr>
		<tr>
			<td>Nikon Ti-E inverted microscope</td>
			<td>Nikon</td>
			<td>N/A</td>
			<td>Nikon LuN4 Laser</td>
		</tr>
		<tr>
			<td>Ni-NTA Resin</td>
			<td>Thermo Scientific</td>
			<td>88221</td>
			<td></td>
		</tr>
		<tr>
			<td>Oligonucleotide - CACCTATTCTGAGTTTGCGCGA 
			GAACTTTCAAAGGC</td>
			<td>IDT</td>
			<td>N/A</td>
			<td></td>
		</tr>
		<tr>
			<td>Oligonucleotide - GCCTTTGAAAGTTCTCGCGCAA 
			ACTCAGAATAGGTG</td>
			<td>IDT</td>
			<td>N/A</td>
			<td></td>
		</tr>
		<tr>
			<td>Open-top thickwall polycarbonate tube, 0.2 mL, 7 mm x 22 mm</td>
			<td>Beckman Coulter</td>
			<td>343755</td>
			<td></td>
		</tr>
		<tr>
			<td>Optima-TLX Ultracentrifuge</td>
			<td>Beckman Coulter</td>
			<td>361544</td>
			<td></td>
		</tr>
		<tr>
			<td>Paclitaxel (Taxol equivalent)</td>
			<td>Thermo Fisher Scientific</td>
			<td>P3456</td>
			<td></td>
		</tr>
		<tr>
			<td>PIPES</td>
			<td>ACROS Organics</td>
			<td>172615000</td>
			<td></td>
		</tr>
		<tr>
			<td>PMSF</td>
			<td>Millipore</td>
			<td>7110-5GM</td>
			<td></td>
		</tr>
		<tr>
			<td>Porcine Tubulin, biotin label</td>
			<td>Cytoskeleton, Inc.</td>
			<td>T333P</td>
			<td></td>
		</tr>
		<tr>
			<td>Porcine Tubulin, HiLyte 647 Fluor</td>
			<td>Cytoskeleton, Inc.</td>
			<td>TL670M</td>
			<td>far red labelled</td>
		</tr>
		<tr>
			<td>Porcine Tubulin, Rhodamine</td>
			<td>Cytoskeleton, Inc.</td>
			<td>TL590M</td>
			<td></td>
		</tr>
		<tr>
			<td>Porcine Tubulin, Tubulin Protein</td>
			<td>Cytoskeleton, Inc.</td>
			<td>T240</td>
			<td></td>
		</tr>
		<tr>
			<td>Potassium Acetate</td>
			<td>Fisher Scientific</td>
			<td>BP364-500</td>
			<td></td>
		</tr>
		<tr>
			<td>Prime 95B sCMOS camera</td>
			<td>Photometric</td>
			<td>N/A</td>
			<td></td>
		</tr>
		<tr>
			<td>Quadrant Detector Sensor Head</td>
			<td>ThorLabs</td>
			<td>PDQ80A</td>
			<td></td>
		</tr>
		<tr>
			<td>Quikchange Lightning Kit</td>
			<td>Agilent Technologies</td>
			<td>210518</td>
			<td></td>
		</tr>
		<tr>
			<td>Sodium Bicarbonate</td>
			<td>Fisher Scientific</td>
			<td>S233-500</td>
			<td></td>
		</tr>
		<tr>
			<td>Sodium Phosphate Dibasic Anhydrous</td>
			<td>Fisher Scientific</td>
			<td>BP332-500</td>
			<td></td>
		</tr>
		<tr>
			<td>Square Cover Glasses</td>
			<td>Corning</td>
			<td>12-553-450</td>
			<td>18 mm x 18 mm, with thickness of 0.13-0.17 mm</td>
		</tr>
		<tr>
			<td>Streptavidin Microspheres</td>
			<td>Polysciences Inc.</td>
			<td>24162-1</td>
			<td></td>
		</tr>
		<tr>
			<td>Superose-6 Column</td>
			<td>GE Healthcare</td>
			<td>29-0915--96</td>
			<td></td>
		</tr>
		<tr>
			<td>TCEP</td>
			<td>Thermo Scientific</td>
			<td>77720</td>
			<td></td>
		</tr>
		<tr>
			<td>TLA-100 Fixed-Angle Rotor</td>
			<td>Beckman Coulter</td>
			<td>343840</td>
			<td></td>
		</tr>
		<tr>
			<td>Ultrasonic Cleaner (Sonicator)</td>
			<td>Vevor</td>
			<td>JPS-08A(DD)</td>
			<td>304 stainless steel, 40 kHz frequency, 60 W power</td>
		</tr>
		<tr>
			<td>Vectabond APTES solution</td>
			<td>Vector Laboratories</td>
			<td>SP-1800-7</td>
			<td></td>
		</tr>
		<tr>
			<td>Windex Powerized Glass Cleaner with Ammonia-D</td>
			<td>S.C. Johnson</td>
			<td>SJN695237</td>
			<td></td>
		</tr>
	</tbody>
</table>

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.

The preparation of microtubule bundles suitable for biophysical analysis is considered successful if several of the key criteria are met. First, imaging in three colors should reveal two aligned microtubules with a concentration of crosslinking protein preferentially decorating the overlap region (Figure 5B,C and Figure 6B). Ideally, the distance between the overlap edge and the free end of the rhodamine microtubule should be at...

Watch this Scientific Journal Video about Directly Measuring Forces Within Reconstituted Active Microtubule Bundles at JoVE.com

Directly Measuring Forces Within Reconstituted Active Microtubule Bundles

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.

Microtubule networks are employed in cells to accomplish a wide range of tasks, ranging from acting as tracks for vesicle transport to working as ...

directly-measuring-forces-within-reconstituted-active-microtubule

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Biology

1.5K Views.  Rensselaer Polytechnic Institute. 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.

Video: Directly Measuring Forces Within Reconstituted Active Microtubule Bundles

Microfabricated Post-Array-Detectors (mPADs): an Approach to Isolate Mechanical Forces

In this video, we will present our approach to measure cellular traction forces using a microfabricated array of posts.  Traction forces are generated through myosin-actin interactions and play an important role in our physiology.  During development, they enable cells to move from one location to the next in order to form the early structures of tissue.  Traction forces help in the healing processes.  They are necessary for the proper closure of wounds or the migration and crawling of leukocytes through our body.  These same forces can be detrimental to our health in the case of cancer metastasis or vascular growth towards a tumor.  The most common method by which to study cells in vitro has been to use a glass or polystyrene dish.  However, the rigidity of the substrates makes it impossible to physically measure cell traction forces, and there are relatively few methods to study traction forces.  Our lab has developed a technique to overcome these limitations.  The method is based on a vertical array of flexible cantilevers, the stiffness and size scale of which are such that individual cells spread across many cantilevers and deflect them in the process.  The pillars we use are 3 &mu;m in diameter, 10 &mu;m tall, and are configured in a regular array with 9 &mu;m center-to-center spacing.  But these physical dimensions can be readily varied to accommodate a variety of studies.  We start with a silicon master, but the final posts are made out of silicone rubber called poly (dimethyl siloxane), or PDMS.  We can measure the deflections under a microscope and calculate the magnitude and direction of traction forces required to produce the observed deflections.  We call these substrates microfabricated post-array-detectors, or mPADs.  Here, we will show you how we fabricate and use the mPADs to assess modulations of cellular contractility.

Microfabricated Post-Array-Detectors mPADs: an Approach to Isolate Mechanical Forces

In this video, we demonstrate how to fabricate and utilize microfabricated post array detectors (mPADs) to assess modulations of cellular contractility.

In this video, we will present our approach to measure cellular traction forces using a microfabricated array of posts.  Traction forces are generated ...

Chromatographic Purification of Highly Active Yeast Ribosomes

Eukaryotic ribosomes are much more labile as compared to their eubacterial and archael counterparts, thus posing a significant challenge to researchers. Particularly troublesome is the fact that lysis of cells releases a large number of proteases and nucleases which can degrade ribosomes. Thus, it is important to separate ribosomes from these enzymes as quickly as possible. Unfortunately, conventional differential ultracentrifugation methods leaves ribosomes exposed to these enzymes for unacceptably long periods of time, impacting their structural integrity and functionality. To address this problem, we utilize a chromatographic method using a cysteine charged Sulfolink resin. This simple and rapid application significantly reduces co-purifying proteolytic and nucleolytic activities, producing high yields of intact, highly biochemically active yeast ribosomes. We suggest that this method should also be applicable to mammalian ribosomes. The simplicity of the method, and the enhanced purity and activity of chromatographically purified ribosome represents a significant technical advancement for the study of eukaryotic ribosomes.

Contamination of preparations of eukaryotic ribosomes purified by traditional methods by co-purifying nucleases and proteases negatively impacts on downstream biochemical and structural analyses. A rapid and simple chromatographic purification method is used to solve this problem using yeast ribosomes as a model system.

Eukaryotic ribosomes are much more labile as compared to their eubacterial and archael counterparts, thus posing a significant challenge to researchers.  ...

Production of Xenopus tropicalis Egg Extracts to Identify Microtubule-associated RNAs

Many organisms localize mRNAs to specific subcellular destinations to spatially and temporally control gene expression. Recent studies have demonstrated that the majority of the transcriptome is localized to a nonrandom position in cells and embryos. One approach to identify localized mRNAs is to biochemically purify a cellular structure of interest and to identify all associated transcripts. Using recently developed high-throughput sequencing technologies it is now straightforward to identify all RNAs associated with a subcellular structure. To facilitate transcript identification it is necessary to work with an organism with a fully sequenced genome. One attractive system for the biochemical purification of subcellular structures are egg extracts produced from the frog Xenopus laevis. However, X. laevis currently does not have a fully sequenced genome, which hampers transcript identification. In this article we describe a method to produce egg extracts from a related frog, X. tropicalis, that has a fully sequenced genome. We provide details for microtubule polymerization, purification and transcript isolation. While this article describes a specific method for identification of microtubule-associated transcripts, we believe that it will be easily applied to other subcellular structures and will provide a powerful method for identification of localized RNAs.

Production of Xenopus tropicalis Egg Extracts to Identify Microtubule-associated RNAs

We describe the collection of unfertilized Xenopus tropicalis eggs and production of a meiosis II-arrested egg extract. This egg extract can be used to purify microtubules and microtubule-associated RNAs.

Many  organisms localize mRNAs to specific subcellular destinations to spatially and  temporally control gene expression. Recent studies have demonstrated ...

Robust 3D DNA FISH Using Directly Labeled Probes

3D DNA FISH has become a major tool for analyzing three-dimensional organization of the nucleus, and several variations of the technique have been published. In this article we describe a protocol which has been optimized for robustness, reproducibility, and ease of use. Brightly fluorescent directly labeled probes are generated by nick-translation with amino-allyldUTP followed by chemical coupling of the dye. 3D DNA FISH is performed using a freeze-thaw step for cell permeabilization and a heating step for simultaneous denaturation of probe and nuclear DNA. The protocol is applicable to a range of cell types and a variety of probes (BACs, plasmids, fosmids, or Whole Chromosome Paints) and allows for high-throughput automated imaging. With this method we routinely investigate nuclear localization of up to three chromosomal regions.

We describe a robust and versatile protocol for analyzing nuclear architecture by 3D DNA FISH using directly labeled fluorescent probes.

3D DNA FISH has become a major tool for  analyzing three-dimensional organization of the nucleus, and several variations  of the technique have been ...

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 ...

Using plusTipTracker Software to Measure Microtubule Dynamics in Xenopus laevis Growth Cones

Microtubule (MT) plus-end-tracking proteins (+TIPs) localize to the growing plus-ends of MTs and regulate MT dynamics1,2. One of the most well-known and widely-utilized +TIPs for analyzing MT dynamics is the End-Binding protein, EB1, which binds all growing MT plus-ends, and thus, is a marker for MT polymerization1. Many studies of EB1 behavior within growth cones have used time-consuming and biased computer-assisted, hand-tracking methods to analyze individual MTs1-3. Our approach is to quantify global parameters of MT dynamics using the software package, plusTipTracker4, following the acquisition of high-resolution, live images of tagged EB1 in cultured embryonic growth cones5. This software is a MATLAB-based, open-source, user-friendly package that combines automated detection, tracking, visualization, and analysis for movies of fluorescently-labeled +TIPs. Here, we present the protocol for using plusTipTracker for the analysis of fluorescently-labeled +TIP comets in cultured Xenopus laevis growth cones. However, this software can also be used to characterize MT dynamics in various cell types6-8.

Using plusTipTracker Software to Measure Microtubule Dynamics in Xenopus laevis Growth Cones

The MATLAB-based, open source software package, plusTipTracker, can be used to analyze image series of fluorescently-labeled +TIPs to quantify microtubule dynamics.

Microtubule (MT) plus-end-tracking proteins (+TIPs) localize to the growing plus-ends of MTs and regulate MT dynamics1,2. One of the most well-known 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 ...

A Multi-hole Cryovial Eliminates Freezing Artifacts when Muscle Tissues are Directly Immersed in Liquid Nitrogen

Studies on skeletal muscle physiology face the technical challenge of appropriately processing the specimens to obtain sections with clearly visible cytoplasmic compartments. Another hurdle is the tight apposition of myofibers to the surrounding tissues. Because the process of tissue fixation and paraffin embedding leads to the shrinkage of muscle fibers, freezing is an optimal means of hardening muscle tissue for sectioning. However, a commonly encountered issue, the formation of ice crystals, occurs during the preparation of frozen sections because of the high water content of muscle. The protocol presented here first describes a simple and efficient method for properly freezing muscle tissues by immersing them in liquid nitrogen. The problem with using liquid nitrogen alone is that it causes the formation of a nitrogen gas barrier next to the tissue, which acts as an insulator and inhibits the cooling of the tissues. To avoid this "vapor blanket" effect, a new cryovial was designed to increase the speed of liquid flow around the tissue surface. This was achieved by punching a total of 14 inlet holes in the wall of the vial. According to bubble dynamics, a higher rate of liquid flow results in smaller bubbles and fewer chances to form a gas barrier. When liquid nitrogen flows into the cryovial through the inlet holes, the flow velocity around the tissue is fast enough to eliminate the gas barrier. Compared to the method of freezing muscle tissues using pre-chilled isopentane, this protocol is simpler and more efficient and can be used to freeze muscle in a throughput manner. Furthermore, this method is optimal for institutions that do not have access to isopentane, which is extremely flammable at room temperature.

This protocol describes a procedure to freeze muscle tissues by plunging them directly into liquid nitrogen. This protocol also highlights a new cryovial that can avoid the &#34;blanket effect&#34; of nitrogen gas when liquid nitrogen contacts the tissue surface of a specimen.

Studies on skeletal muscle physiology face the technical challenge of appropriately processing the specimens to obtain sections with clearly visible ...

Experimental Protocol for Using Drosophila As an Invertebrate Model System for Toxicity Testing in the Laboratory

Emergent properties and external factors (population-level and ecosystem-level interactions, in particular) play important roles in mediating ecologically-important endpoints, though they are rarely considered in toxicological studies. D. melanogaster is emerging as a toxicology model for the behavioral, neurological, and genetic impacts of toxicants, to name a few. More importantly, species in the genus Drosophila can be utilized as a model system for an integrative framework approach to incorporate emergent properties and answer ecologically-relevant questions in&#160;toxicology research. The aim of this paper is to provide a protocol for exposing species in the genus Drosophila to pollutants to be used as a model system for a range of phenotypic outputs and ecologically-relevant questions. More specifically, this protocol can be used to 1) link multiple biological levels of organization and understand the impact of toxicants on both individual- and population-level fitness; 2) test the impact of toxicants at different stages of developmental exposure; 3) test multigenerational and evolutionary implications of pollutants; and 4) test multiple contaminants and stressors simultaneously.

Experimental Protocol for Using Drosophila As an Invertebrate Model System for Toxicity Testing in the Laboratory

In this paper, we provide a detailed protocol for exposing species in the genus Drosophila to pollutants with the goal of studying the impact of exposure on a range of phenotypic outputs at different developmental stages and for more than one generation.

Emergent properties and external factors (population-level and ecosystem-level interactions, in particular) play important roles in mediating ...

Micron-scale Phenotyping Techniques of Maize Vascular Bundles Based on X-ray Microcomputed Tomography

It is necessary to accurately quantify the anatomical structures of maize materials based on high-throughput image analysis techniques. Here, we provide a 'sample preparation protocol' for maize materials (i.e., stem, leaf, and root) suitable for ordinary microcomputed tomography (micro-CT) scanning. Based on high-resolution CT images of maize stem, leaf, and root, we describe two protocols for the phenotypic analysis of vascular bundles: (1) based on the CT image of maize stem and leaf, we developed a specific image analysis pipeline to automatically extract 31 and 33 phenotypic traits of vascular bundles; (2) based on the CT image series of maize root, we set up an image processing scheme for the three-dimensional (3-D) segmentation of metaxylem vessels, and extracted two-dimensional (2-D) and 3-D phenotypic traits, such as volume, surface area of metaxylem vessels, etc. Compared with traditional manual measurement of vascular bundles of maize materials, the proposed protocols significantly improve the efficiency and accuracy of micron-scale phenotypic quantification.

We provide a novel method to improve the X-ray absorption contrast of maize tissue suitable for ordinary microcomputed tomography scanning. Based on CT images, we introduce a set of image-processing workflows for different maize materials to effectively extract microscopic phenotypes of vascular bundles of maize.

It is necessary to accurately quantify the anatomical structures of maize materials based on high-throughput image analysis techniques. Here, we provide a ...