We are grateful to the B. Ladoux and R.-M. M&#232;ge lab for the use of their UV-cleaner equipment, and J. Heuvingh and 0. du Roure for the initial training we received on preparing molds on silicon wafers and providing tips on microfluidics. We acknowledge funding from European Research Council Grant StG-679116 (to A.J.) and Agence Nationale de la Recherche Grants Muscactin and Conformin (to G.R.-L.).

<ol>
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The authors declare no conflicts of interest.

Compared to standard single-filament methods where actin filaments are anchored to the surface by multiple points along their length or maintained close to it by a crowding agent such as methylcellulose, microfluidics offers a number of advantages. As interactions with the surface are minimal, the artificial pauses these interactions can induce during both elongation and depolymerization are avoided. The filaments are aligned by the flow, parallel to each other, easing their monitoring and the measurement of their length...

The assembly and disassembly of actin filaments and actin filament networks are controlled by several biochemical reactions and depend on the mechanical context. In order to gain insight into these complex mechanisms, it is invaluable to be able to observe individual reactions on individual filaments (in sufficiently large numbers). Over the past decades, the observation of dynamic actin filaments in real-time, mostly using total internal reflection fluorescence (TIRF) microscopy, has emerged as a key technique and has provided an impressive list of results that could not have been obtained with bulk solution biochemical assays1.
To achieve this, one needs to maintain fluorescently labeled actin filaments close to the surface of the microscope coverslip while exposing them to solutions of actin-binding proteins (ABPs), which can also be fluorescently labeled. Doing so provides a means to monitor events taking place on individual filaments in well-controlled biochemical conditions, and thus quantify reaction rates. However, a number of specific limitations should be considered. Artificially maintaining filaments close to the surface, often thanks to multiple anchoring points or by using a crowding agent such as methylcellulose, can alter their behavior (e.g., causing pauses in their polymerization and depolymerization2). Tracking the contour of each filament can be challenging, particularly if new filaments or filament fragments accumulate in the field of view over time. The reactions take place in a finite volume where the concentration of actin monomers and ABPs can vary over time, potentially making it difficult to derive accurate rate constants. Finally, renewing or changing the solution of ABPs is difficult to achieve in less than 30 s and will often lead to inhomogeneous protein content in the sample.
A bit over 10 years ago, inspired by what was already done to study individual Deoxyribonucleic Acid (DNA) strands3, we introduced a new technique based on microfluidics to observe and manipulate individual actin filaments4. It allows one to circumvent the aforementioned limitations of classical single-filament techniques. In these microfluidics assays, actin filaments are grown from spectrin-actin seeds adsorbed on the coverslip. Filaments are thus anchored by one end only to the bottom of the microfluidic chamber and fluctuate above the surface without sticking. Filaments align with the flow of incoming solutions, thereby easing the monitoring of their contour length and maintaining them in a shallow region above the coverslip where TIRF can be used. Different solutions are simultaneously flowed into the chamber without mixing, and the filaments can be exposed to them sequentially and rapidly.
Here, we propose a series of basic protocols to set up single-actin-filament microfluidics assays in the lab. Coverslips and microfluidics chambers can be prepared in advance (in half a day), and the experiment itself, where several biochemical conditions can be tested, is done in less than a day.

<table><tbody><tr><td>&amp;beta;-Casein</td><td>Merck</td><td>C6905</td><td>Used at 8 mg/mL</td></tr><tr><td>Biopsy punch (with plunger)</td><td>Ted Pella</td><td>15115-2</td><td>ID 0.75 mm, OD 1.07 mm</td></tr><tr><td>Biotin-BSA</td><td>Merck</td><td>A8549</td><td>Used at 1 mg/mL</td></tr><tr><td>BSA</td><td>Merck</td><td>A8022</td><td>Used at 50 mg/mL</td></tr><tr><td>Coverslip Mini-Rack&lt;br/&gt; Teflon holder</td><td>Invitrogen</td><td>C14784</td><td>for 8 coverslips</td></tr><tr><td>Coverslips 22x40mm&lt;br/&gt; Thickness #1.5</td><td>Menzel Gl&amp;auml;ser</td><td>631-1370</td><td></td></tr><tr><td>DABCO</td><td>Merck</td><td>D27802</td><td>component in f-buffer</td></tr><tr><td>DTT</td><td>Euromedex</td><td>EU0006-D</td><td>component in f-buffer</td></tr><tr><td>Ester NHS Alexa Fluor 488</td><td>Invitrogen</td><td>A20000</td><td>Fluorophore for actin labeling on Lys328.</td></tr><tr><td>EZ-Link Sulfo-NHS-Biotin</td><td>Thermo Scientific</td><td>21338</td><td>To biotinylate actin on Lys328</td></tr><tr><td>Hellmanex III</td><td>Hellma</td><td>9-307-011-4-507</td><td>Glass cleaning detergent</td></tr><tr><td>ImageJ</td><td>NIH</td><td>N/A</td><td>open source software</td></tr><tr><td>Laboport</td><td>KNF</td><td>811kn.18</td><td>vacuum pump (ultimate vacuum: 240 mbar)</td></tr><tr><td>Magic invisible tape</td><td>Scotch</td><td>7100024666</td><td>standard transparent office tape</td></tr><tr><td>Micrewtube</td><td>Simport</td><td>T341-6T</td><td>2 mL microfluidic reservoir tubes</td></tr><tr><td>Microfluidic device Part 1: Flow Unit S</td><td>Fluigent</td><td>FLU-S-D-PCKB</td><td>Flowmeter</td></tr><tr><td>Microfluidic device Part 2: Fluiwell-4C-2 mL</td><td>Fluigent</td><td>14002001PCK</td><td>Reservoir holder</td></tr><tr><td>Microfluidic device Part 3: MFCS-EZ</td><td>Fluigent</td><td>EZ-11000001&lt;br/&gt; EZ-00345001</td><td>Pressure controller</td></tr><tr><td>Model 42 - UVO-Cleaner</td><td>Jelight Inc.</td><td>42-220</td><td>Ultraviolet cleaner</td></tr><tr><td>N6-(6-Aminohexyl)-ATP-ATTO-488</td><td>Jena Bioscience</td><td>NU-805-488</td><td>ATP-ATTO used to label actin</td></tr><tr><td>neutravidin</td><td>Thermo Scientific</td><td>31000</td><td></td></tr><tr><td>PLL-PEG</td><td>SuSoS</td><td>PLL(20)-g[3.5]- PEG(2)</td><td>Use at 1 mg/mL in PBS.</td></tr><tr><td>Polydimethylsiloxane (PDMS) Sylgard 184 Silicon Elastomer</td><td>Dow Corning</td><td>1673921</td><td>Contains PDMS base and curing agent</td></tr><tr><td>Polyetheretherketone (PEEK) tubing</td><td>Merck</td><td>Z226661</td><td>&amp;ldquo;Blue&amp;rdquo; : I.D. = 0.25 mm</td></tr><tr><td>Safety blow gun</td><td>Coilhose Pneumatics</td><td>700-S</td><td>filtered air</td></tr><tr><td>Silicon tubing</td><td>VWR</td><td>228-0701P</td><td>connect PEEK to coupler</td></tr><tr><td>Stainless steel catheter coupler</td><td>Prime Bioscience</td><td>SC22/15</td><td>Inserted into PDMS inlets and outlet to connect to PEEK tubing</td></tr><tr><td>Thermoplastic film</td><td>Sigma Aldrich</td><td>PM996</td><td>Standard &quot;parafilm&quot;</td></tr><tr><td>Ultrapure ethanol</td><td>VWR</td><td>64-17-5</td><td></td></tr><tr><td>Ultrasonic cleaning bath</td><td>VWR</td><td>USC200TH</td><td>To accomodate 1 L beakers</td></tr><tr><td>Vacuum dessicator</td><td>SP Bel-Art</td><td>F42022-0000</td><td>to degas the PDMS or solutions</td></tr></tbody></table>

using microfluidics and fluorescence microscopy to study the assembly dynamics of single actin filaments and bundles

In order to decipher the complex molecular mechanisms that regulate the assembly and disassembly of actin filaments, it is a great asset to monitor individual reactions live in well-controlled conditions. To do so, live single-filament experiments have emerged over the past 20 years, mostly using total internal reflection fluorescence (TIRF) microscopy, and have provided a trove of key results. In 2011, in order to further expand the possibilities of these experiments and to avoid recurring problematic artifacts, we introduced simple microfluidics in these assays. This study details our basic protocol, where individual actin filaments are anchored by one end to the passivated coverslip surface, align with the flow, and can be successively exposed to different protein solutions. We also present the protocols for specific applications and explain how controlled mechanical forces can be applied, thanks to the viscous drag of the flowing solution. We highlight the technical caveats of these experiments and briefly present possible developments based on this technique. These protocols and explanations, along with today&#39;s availability of easy-to-use microfluidics equipment, should allow non-specialists to implement this assay in their labs.

For all the experiments described above, fluorescently labeled actin filaments should be clearly visible, with good contrast, indicative of low background fluorescence from the surface (Figure 4, see Supplementary File 1 for troubleshooting of common issues). Actin filaments should also not stick to the surface: when the dominant flow rate is low, the actin filaments&#39; lateral fluctuations should be perceptible when observing them live and allow one to clearly determine t...

Watch this Scientific Journal Video about Using Microfluidics and Fluorescence Microscopy to Study the Assembly Dynamics of Single Actin Filaments and Bundles at JoVE.com

Using Microfluidics and Fluorescence Microscopy to Study the Assembly Dynamics of Single Actin Filaments and Bundles

We present protocols for simple actin filament microfluidic assays, in combination with fluorescence microscopy, that allow one to accurately monitor individual actin filaments in real-time while sequentially exposing them to different protein solutions.

In order to decipher the complex molecular mechanisms that regulate the assembly and disassembly of actin filaments, it is a great asset to monitor ...

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2.5K Views.  Institut Jacques Monod. We present protocols for simple actin filament microfluidic assays, in combination with fluorescence microscopy, that allow one to accurately monitor individual actin filaments in real-time while sequentially exposing them to different protein solutions.

Video: Using Microfluidics and Fluorescence Microscopy to Study the Assembly Dynamics of Single Actin Filaments and Bundles

Monitoring Actin Disassembly with Time-lapse Microscopy

Okay, so what I'd like to do today is to show you how to construct a flow chamber, a fusion chamber for imaging on an upright microscope. What then I then do is I create two spaces using film. So what I do is I take the para film, I cut it into small strips.Next I'm going to, this will basically form the channels for the, the walls, and I dry them with a tilted air. I put the cover slip on top of the para, And here's the finished. Then I heat it on a heat block to melt the param and basically create a seal.So what this is, is the, the cover glass here, and then on top of the cover glass is two layers of phim as you see right here. And then on top of this, again, there's a, it's a 22 by two two mil meter cover slip. This one right here, which I pressed against the param to fall the seal.Okay, so the liquid goes in in one direction here and I can suck it out with a filter paper here in this direction. And I'll show you how to do that in a bit. So what I like to do is I like to put it in a humidified chamber, which is very simple.Take two kind of elevated strips and have a filter of paper underneath. And to keep it noisy as I, I put the cover, cover slip on the flow chamber on. Now the way you introduce a solution to the chamber is that you just put the pipet tip at the entrance of the chamber and you just split.Now what I do is I incubate the chamber with the lid closed for 10 minutes. What I usually do to suck the liquid out from the other side is to use it, get a filter paper and cut it into thin strips like this. And I take one of these thin strips and I simultaneously apply the solution into the entrance while putting the filter paper at the exit to suck the exchange, the solution.So you'll see the liquid flowing through the solution. And then I wait for five minutes for the mid close. Now what I'm ready to do is to actually put that the actin it and the actin, I actually have to polymerize, I polymerize inside the flow chamber and it's just the same procedures before.So because like this, and then now that the action is polymerized, I wash out all the un excess un polymerized action using a large volume of just buffer. So here I have about a hundred microliters and it's a few reaction chamber volumes worth of buffer. Just, you know, when the filter paper becomes saturated, I have to change the direction.Okay?And so I'm ready to do imaging with this. So perfusion chamber with actin attached to it. Now I take the slide, mount it under the microscope, and since I'm imaging with an oil objective, I'm gonna put some immersion oil on the top surface of the cover, grass touch, such so assess solution or protein or anything on acting inside.So what I do is I can actually exchange the solution side while I'm doing the time lapse imaging. As such.

Biology

Visualizing Actin and Microtubule Coupling Dynamics In Vitro by Total Internal Reflection Fluorescence (TIRF) Microscopy

Traditionally, the actin and microtubule cytoskeletons have been studied as separate entities, restricted to specific cellular regions or processes, and regulated by different suites of binding proteins unique for each polymer. Many studies now demonstrate that the dynamics of both cytoskeletal polymers are intertwined and that this crosstalk is required for most cellular behaviors. A number of proteins involved in actin-microtubule interactions have already been identified (i.e., Tau, MACF, GAS, formins, and more) and are well characterized with regard to either actin or microtubules alone. However, relatively few studies showed assays of actin-microtubule coordination with dynamic versions of both polymers. This may occlude emergent linking mechanisms between actin and microtubules. Here, a total internal reflection fluorescence (TIRF) microscopy-based in vitro reconstitution technique permits the visualization of both actin and microtubule dynamics from the one biochemical reaction. This technique preserves the polymerization dynamics of either actin filament or microtubules individually or in the presence of the other polymer. Commercially available Tau protein is used to demonstrate how actin-microtubule behaviors change in the presence of a classic cytoskeletal crosslinking protein. This method can provide reliable functional and mechanistic insights into how individual regulatory proteins coordinate actin-microtubule dynamics at a resolution of single filaments or higher-order complexes.

Visualizing Actin and Microtubule Coupling Dynamics In Vitro by Total Internal Reflection Fluorescence TIRF Microscopy

This protocol is a guide for visualizing dynamic actin and microtubules using an in vitro total internal fluorescence (TIRF) microscopy assay.

Traditionally, the actin and microtubule cytoskeletons have been studied as separate entities, restricted to specific cellular regions or processes, and ...

Single-Molecule Diffusion and Assembly on Polymer-Crowded Lipid Membranes

Cellular membranes are highly crowded environments for biomolecular reactions and signaling. Yet, most in vitro experiments probing protein interaction with lipids employ naked bilayer membranes. Such systems lack the complexities of crowding by membrane-embedded proteins and glycans and exclude the associated volume effects encountered on cellular membrane surfaces. Also, the negatively charged glass surface onto which the lipid bilayers are formed prevents the free diffusion of transmembrane biomolecules. Here, we present a well-characterized polymer-lipid membrane as a mimic for crowded lipid membranes. This protocol utilizes polyethylene glycol (PEG)-conjugated lipids as a generalized approach for incorporating crowders into the supported lipid bilayer&#160;(SLB). First, a cleaning procedure of the microscopic slides and coverslips for performing single-molecule experiments is presented. Next, methods for characterizing the PEG-SLBs and performing single-molecule experiments of the binding, diffusion, and assembly of biomolecules using single-molecule tracking and photobleaching are discussed. Finally, this protocol demonstrates how to monitor the nanopore assembly of bacterial pore-forming toxin Cytolysin A (ClyA) on crowded lipid membranes with single-molecule photobleaching analysis. MATLAB codes with example datasets are also included to perform some of the common analyses such as particle tracking, extracting diffusive behavior, and subunit counting.

Here, a protocol to perform and analyze the binding, mobility, and assembly of single molecules on artificial crowded lipid membranes using single-molecule total internal reflection fluorescence (smTIRF) microscopy is presented.

Cellular membranes are highly crowded environments for biomolecular reactions and signaling. Yet, most in vitro experiments probing protein interaction ...

Probing Myosin Ensemble Mechanics in Actin Filament Bundles Using Optical Tweezers

Myosins are motor proteins that hydrolyze ATP to step along actin filament (AF) tracks and are essential in cellular processes such as motility and muscle contraction. To understand their force-generating mechanisms, myosin II has been investigated both at the single-molecule (SM) level and as teams of motors in vitro using biophysical methods such as optical trapping.
These studies showed that myosin force-generating behavior can differ greatly when moving from the single-molecule level in a three-bead arrangement to groups of motors working together on a rigid bead or coverslip surface in a gliding arrangement. However, these assay constructions do not permit evaluating the group dynamics of myosin within viscoelastic structural hierarchy as they would within a cell. We have developed a method using optical tweezers to investigate the mechanics of force generation by myosin ensembles interacting with multiple actin filaments.
These actomyosin bundles facilitate investigation in a hierarchical and compliant environment that captures motor communication and ensemble force output. The customizable nature of the assay allows for altering experimental conditions to understand how modifications to the myosin ensemble, actin filament bundle, or the surrounding environment result in differing force outputs.

Formation of actomyosin bundles in vitro and measuring myosin ensemble force generation using optical tweezers is presented and discussed.

Myosins are motor proteins that hydrolyze ATP to step along actin filament (AF) tracks and are essential in cellular processes such as motility and muscle ...

Bioengineering

The Mechanics of (Poro-)Elastic Contractile Actomyosin Networks As a Model System of the Cell Cytoskeleton

Cells can actively change their shapes and become motile, a property that depends on their ability to actively reorganize their internal structure. This feature is attributed to the mechanical and dynamic properties of the cell cytoskeleton, notably, the actomyosin cytoskeleton, which is an active gel of polar actin filaments, myosin motors, and accessory proteins that exhibit intrinsic contraction properties. The usually accepted view is that the cytoskeleton behaves as a viscoelastic material. However, this model cannot always explain the experimental results, which are more consistent with a picture describing the cytoskeleton as a poroelastic active material-an elastic network embedded with cytosol. Contractility gradients generated by the myosin motors drive the flow of the cytosol across the gel pores, which infers that the mechanics of the cytoskeleton and the cytosol are tightly coupled. One main feature of poroelasticity is the diffusive relaxation of stresses in the network, characterized by an effective diffusion constant that depends on the gel elastic modulus, porosity, and cytosol (solvent) viscosity. As cells have many ways to regulate their structure and material properties, our current understanding of how cytoskeleton mechanics and cytosol flow dynamics are coupled remains poorly understood. Here, an in vitro reconstitution approach is employed to characterize the material properties of poroelastic actomyosin gels as a model system for the cell cytoskeleton. Gel contraction is driven by myosin motor contractility, which leads to the emergence of a flow of the penetrating solvent. The paper describes how to prepare these gels and run experiments. We also discuss how to measure and analyze the solvent flow and gel contraction both at the local and global scales. The various scaling relations used for data quantification are given. Finally, the experimental challenges and common pitfalls are discussed, including their relevance to cell cytoskeleton mechanics.

The Mechanics of Poro-Elastic Contractile Actomyosin Networks As a Model System of the Cell Cytoskeleton

In this work, an in vitro reconstitution approach is employed to study the poroelasticity of actomyosin gels under controlled conditions. The dynamics of the actomyosin gel and the embedded solvent are quantified, through which the network poroelasticity is demonstrated. We also discuss the experimental challenges, common pitfalls, and relevance to cell cytoskeleton mechanics.

Cells can actively change their shapes and become motile, a property that depends on their ability to actively reorganize their internal structure. This ...

Analyses of Actin Dynamics, Clutch Coupling and Traction Force for Growth Cone Advance

To establish functional networks, neurons must migrate to their appropriate destinations and then extend axons toward their target cells. These processes depend on the advances of growth cones that located at the tips of neurites. Axonal growth cones generate driving forces by sensing their local microenvironment and modulating cytoskeletal dynamics and actin-adhesion coupling (clutch coupling). Decades of research have led to the identification of guidance molecules, their receptors, and downstream signaling cascades for regulating neuronal migration and axonal guidance; however, the molecular machineries required for generating forces to drive growth cone advance and navigation are just beginning to be elucidated. At the leading edge of neuronal growth cones, actin filaments undergo retrograde flow, which is powered by actin polymerization and actomyosin contraction. A clutch coupling between F-actin retrograde flow and adhesive substrate generates traction forces for growth cone advance. The present study describes a detailed protocol for monitoring F-actin retrograde flow by single speckle imaging. Importantly, when combined with an F-actin marker Lifeact, this technique can quantify 1) the F-actin polymerization rate and 2) the clutch coupling efficiency between F-actin retrograde flow and the adhesive substrate. Both are critical variables for generating forces for growth cone advance and navigation. In addition, the present study describes a detailed protocol of traction force microscopy, which can quantify 3) traction force generated by growth cones. Thus, by coupling the analyses of&#160;single speckle imaging and traction force microscopy, investigators can monitor the molecular mechanics underlying growth cone advance and navigation.

To advance, growth cones must exert traction forces against the external environment. The generation of traction forces is dependent on actin dynamics and clutch coupling. The present study describes methods for analyzing actin dynamics, clutch coupling and traction forces for growth cone advance.

To establish functional networks, neurons must migrate to their appropriate destinations and then extend axons toward their target cells. These processes ...

Neuroscience

Micromanipulation Techniques Allowing Analysis of Morphogenetic Dynamics and Turnover of Cytoskeletal Regulators

Examining the spatiotemporal dynamics of proteins can reveal their functional importance in various contexts. In this article, it is discussed how fluorescent recovery after photobleaching (FRAP) and photoactivation techniques can be used to study the spatiotemporal dynamics of proteins in subcellular locations. We also show how these techniques enable straightforward determination of various parameters linked to actin cytoskeletal regulation and cell motility. Moreover, the microinjection of cells is additionally described as an alternative treatment (potentially preceding or complementing the aforementioned photomanipulation techniques) to trigger instantaneous effects of translocated proteins on cell morphology and function. Micromanipulation such as protein injection or local application of plasma membrane-permeable drugs or cytoskeletal inhibitors can serve as powerful tool to record immediate consequences of a given treatment on cell behavior at the single cell and subcellular level. This is exemplified here by immediate induction of lamellipodial cell edge protrusion by the injection of recombinant Rac1 protein, as established a quarter-century ago. In addition, we provide a protocol for determining the turnover of enhanced green fluorescent protein (EGFP)-VASP, an actin filament polymerase prominently accumulating at lamellipodial tips of B16-F1 cells, employing FRAP and including associated data analysis and curve fitting. We also present guidelines for estimating the rates of lamellipodial actin network polymerization, as exemplified by cells expressing EGFP-tagged &#946;-actin. Finally, instructions are given for how to investigate the rates of actin monomer mobility within the cell cytoplasm, followed by actin incorporation at sites of rapid filament assembly, such as the tips of protruding lamellipodia, using photoactivation approaches. None of these protocols is&#160;restricted to components or regulators of the actin cytoskeleton, but can easily be extended to explore in analogous fashion the spatiotemporal dynamics and function of proteins in various different subcellular structures or functional contexts.

We describe how micro- and photomanipulation techniques such as FRAP and photoactivation enable the determination of motility parameters and the spatiotemporal dynamics of proteins within migrating cells. Experimental readouts include subcellular dynamics and turnover of motility regulators or of the underlying actin cytoskeleton.

Examining the spatiotemporal dynamics of proteins can reveal their functional importance in various contexts. In this article, it is discussed how ...

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.

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

Reconstituting and Characterizing Actin-Microtubule Composites with Tunable Motor-Driven Dynamics and Mechanics

The composite cytoskeleton, comprising interacting networks of semiflexible actin filaments and rigid microtubules, restructures and generates forces using motor proteins such as myosin II and kinesin to drive key processes such as migration, cytokinesis, adhesion, and mechanosensing. While actin-microtubule interactions are key to the cytoskeleton's versatility and adaptability, an understanding of their interplay with myosin and kinesin activity is still nascent. This work describes how to engineer tunable three-dimensional composite networks of co-entangled actin filaments and microtubules that undergo active restructuring and ballistic motion, driven by myosin II and kinesin motors, and are tuned by the relative concentrations of actin, microtubules, motor proteins, and passive crosslinkers. Protocols for fluorescence labeling of the microtubules and actin filaments to most effectively visualize composite restructuring and motion using multi-spectral confocal imaging are also detailed. Finally, the results of data analysis methods that can be used to quantitatively characterize non-equilibrium structure, dynamics, and mechanics are presented. Recreating and investigating this tunable biomimetic platform provides valuable insight into how coupled motor activity, composite mechanics, and filament dynamics can lead to myriad cellular processes from mitosis to polarization to mechano-sensation.

This paper presents protocols for engineering and characterizing tunable three-dimensional composite networks of co-entangled actin filaments and microtubules. Composites undergo active restructuring and ballistic motion, driven by myosin II and kinesin motors, and are tuned by the relative concentrations of actin, microtubules, motor proteins, and passive crosslinkers.

The composite cytoskeleton, comprising interacting networks of semiflexible actin filaments and rigid microtubules, restructures and generates forces ...

Studying the Cytoskeleton

The cytoskeletal architecture can be studied using different microscopic and biochemical techniques. Electron microscopy was instrumental in discovering the cytoskeletal architecture around the 1960s, which allowed obtaining structural information at a high-resolution level. However, the sample preparation procedure often limits this ability in biological samples. Several protocols have been developed over the years to optimize sample preparation. In one of the protocols known as rotary shadowing or metal replica, the cytoskeletal filaments are fixed using chemicals such as glutaraldehyde or formaldehyde, followed by dehydration of the samples using alcohol to a critical point of drying and then embedded into a resin, followed by thin sectioning. The sample is positively or negatively stained with heavy metal salts such as platinum and observed under the electron microscope.&#160;&#160;
However, electron microscopy techniques only produce static images and cannot help in studying the dynamic structure of the cytoskeletal filaments and their functions. A better alternative to understanding the cytoskeleton&#39;s structure and function is to study it using fluorescence microscopy. This technique has helped researchers analyze the molecular processes of living cells in real-time through an approach known as live-cell imaging. A commonly used protocol for live-cell imaging involves synthesizing fluorescently labeled proteins within a cell as fusion proteins with GFP or green fluorescent protein. Alternatively, protein subunits of cytoskeletal structures such as purified actin or tubulin can be covalent to a small fluorescent dye in vitro and injected into a living cell. These fusion or tagged proteins are then observed under a fluorescent microscope to study their dynamic behavior in the cell.

The cytoskeletal architecture can be studied using different microscopic and biochemical techniques. Electron microscopy was instrumental in discovering ...