Name: directly measuring forces within reconstituted active microtubule bundles
Uploaded: 2022-05-10T15:00:00
Description: Watch this Scientific Journal Video about Directly Measuring Forces Within Reconstituted Active Microtubule Bundles at JoVE.com

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).

1. Preparation of microtubules
NOTE: When employing GFP-labeled crosslinking proteins, red (e.g., rhodamine) and far-red (e.g., biotinylated HiLyte647, referred to as biotinylated far red in the rest of the text) organic fluorophore labeling of the microtubules works well. Minimal crosstalk between all three channels can be achieved during imaging by using a high-quality quad band total internal reflection fluorescence (TIRF) filter.
<ol>
	<li>Prepare GMPCPP microtubule seed...

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.

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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 ...

Our protocol allows researchers to build cytoskeletal motifs out of purified components and directly measure the forces these networks generate in order to understand how these mechanical components of the cell function in both healthy and disease states. This method allows us to quantify forces and correlate these numbers directly to keep parameters of the proteins involved, including protein concentration and density. Parameters that are generally impossible to acquire within the cell.This method can provide insights into any type of cytoskeleton network, such as those involved in mitosis or neural development. It can readily be adapted to study networks involved in muscle contraction or cell migration by simply swapping out this specific proteins being used. The trickiest aspect of this method is the coordination of the different technologies, which include a single-beam optical trap, and a TIRF microscope.Additionally, single-molecule experiments require careful preparation of surfaces, such as the cover slip, so preparing high-quality samples is paramount. Start with constructing a humidity chamber by placing a folded-up, lint-free tissue dampened with ultrapure water at the bottom of an empty pipette tip box. Introduce all the reagents by pipetting the liquid at one end of the channel and wicking the liquid out at the opposite end.Using a 20-microliter-angled pipette tip, slowly flow in 10 microliters of 0.5 milligrams per milliliter streptavidin or NeutrAvidin. Incubate the slide in the humidity chamber for 4 minutes with the cover slip facing down. Flush out the chamber using 20 microliters of a 1x BRB80 using a lint-free tissue to draw out liquids.After repeating flowing reagents and incubation one more time, flush step with 10 microliters of 0.5 milligrams per milliliter casein blocking solution. Flow 10 microliters of diluted biotinylated-far-red microtubules into the chamber, then flush the chamber with 20 microliters of 1x BRB80. After flowing the reagent and incubating as demonstrated previously, flush the step with 10 microliters of an appropriate concentration of non-motor protein to be studied, then flow in 10 microliters of rhodamine microtubules and repeat the incubation and flushing.Next, combine 1 microliter of rigor kinesin-coated beads and 19 microliters of reaction buffer. Seal both edges of the chamber with clear nail polish and wait until the polish dries. Employ an inverted microscope instrument that has total internal reflection fluorescence imaging capabilities, and into which a single-beam optical trap has been constructed.Add 1 drop of immersion oil onto the cover slip of the sample chamber. With the cover slip side of the sample chamber facing downward, place the sample to be imaged on the microscope platform. Slowly bring the objective up so that there is contact between both the cover slip and the objective through the oil.Adjust the objective height so that the cover slip surface of the sample chamber is in focus. Record single-frame images of the samples in all three channels. For the experiments here, use 100 to 200 milliseconds of exposure.Adjust the laser power to achieve a good signal-to-noise ratio from the samples while minimizing photo bleaching over 2 to 5 minutes when the individual bundle is assayed. The frequency of microtubule bundles per field of view should be approximately three to four microtubule bundles per 100 microliters by 100 microliters field of view per chamber. Observe the density of beads in the sample using a bright field and the 100x objective on the microscope.Employ a concentration of beads such that the beads are a minimum of 10 micrometers from one another on average, and ensure that they are free from any kind of surface confinement or clustering. Ensure that the biotinylated far-red microtubules are firmly attached to the surface of the cover slip and that the rhodamine with no visible Brownian motion. Ensure that the rhodamine microtubules are attached to biotinylated microtubules via the cross-linking map and are visibly fluctuating at their free ends.Identify a suitable microtubule bundle that contains only two microtubules, one biotinylated with full surface attachment and one non-biotinylated microtubule that is partly free in solution and is aligned on either the x or y-axis. Use the optical trap to capture a free bead demonstrating Brownian motion. Once the bead has been trapped, carefully move the bead over to the selected microtubule bundle ensuring no other beads are pulled into the trap in the process.Ensure that the bead is several microns away from the overlap region containing cross-linking proteins. Carefully lower the bead in the z-axis until it makes contact with the edge of the free segment of the rhodamine microtubule. Pull up gently and move perpendicular to the microtubule axis to check for attachment by observing the rhodamine microtubule bending and following the bead position.Carefully realign the rhodamine microtubule along the microtubule axis of the surface-attached microtubule by moving the nano-positioning stage and set up the parameters for automated pulling of the microtubule bundle. Set the direction along the parallel axis of the microtubules. Set the desired pull speed and desired pull time depending on overlap length.Begin recording fluorescence data in the three channels at a rate of 1 to 2 frames per second. Use the optimized the laser powers and exposure times. Initiate automated stage motion and record trapping data from the position-sensitive detector stage and total internal reflection fluorescence camera stage.Deactivate the traps to release the bead and repeat the experiment as desired. Ensembles of the essential mitotic motor protein kinesin-5 can regulate microtubule sliding by generating both pushing and braking forces that scale linearly with overlap length. It was found that the C-terminal tail domain is required for efficient cross-linking and force generation.The full-length protein is able to generate sustained forces that plateau when all motor proteins reach their individual stall force. However, the kinesin-5 motor lacking its C-terminal tail generates maximum forces that are nearly fivefold smaller in magnitude. Ensembles of the non-motor mitotic protein PRC1 operate as a mechanical dash pod to resist microtubule sliding in anaphase spindle mid-zone.The resistive forces do not depend on the length of overlap regions between microtubule pairs or the local crosslinker density, but they do strongly depend on the total concentration of engaged PRC1 molecules. This method can be adapted to study diverse biological systems such as microbial organization and neurons by using alternate proteins. It can also readily be expanded to analyze actin-based network geometries to study muscle contraction or cell motility.This technique lets one study unique cell total geometries, where filaments are both track and cargo and are organized by small ensembles of proteins. These geometries better mimic what's happening in cells during numerous biological processes.

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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 ...

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 ...

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

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 ...

Protrusion Force Microscopy: A Method to Quantify Forces Developed by Cell Protrusions

In numerous biological contexts, animal cells need to interact physically with their environment by developing mechanical forces. Among these, traction ...