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><li>Wioland, H., Jégou, A., Romet-Lemonne, G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Celebrating+20+years+of+live+single-actin-filament+studies+with+five+golden+rules%5BTitle%5D)+AND+%22Proceedings+of+the+National+Academy+of+Sciences+of+the+United+States+of+America%22%5BJournal%5D)">Celebrating 20 years of live single-actin-filament studies with five golden rules.</a> Proceedings of the National Academy of Sciences of the United States of America. 119 (3), 2109506119(2022).</li>
<li>Kuhn, J. R., Pollard, T. D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Real-time+measurements+of+actin+filament+polymerization+by+total+internal+reflection+fluorescence+microscopy%5BTitle%5D)+AND+%22Biophysical+Journal%22%5BJournal%5D)">Real-time measurements of actin filament polymerization by total internal reflection fluorescence microscopy.</a> Biophysical Journal. 88 (2), 1387-1402 (2005).</li>
<li>Brewer, L. R., Bianco, P. R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Laminar+flow+cells+for+single-molecule+studies+of+DNA-protein+interactions%5BTitle%5D)+AND+%22Nature+Methods%22%5BJournal%5D)">Laminar flow cells for single-molecule studies of DNA-protein interactions.</a> Nature Methods. 5 (6), 517-525 (2008).</li>
<li>Jégou, A., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Individual+actin+filaments+in+a+microfluidic+flow+reveal+the+mechanism+of+ATP+hydrolysis+and+give+insight+into+the+properties+of+profilin%5BTitle%5D)+AND+%22PLoS+Biology%22%5BJournal%5D)">Individual actin filaments in a microfluidic flow reveal the mechanism of ATP hydrolysis and give insight into the properties of profilin.</a> PLoS Biology. 9 (9), 1001161(2011).</li>
<li>Gicquel, Y., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Microfluidic+chips+for+in+situ+crystal+x-ray+diffraction+and+in+situ+dynamic+light+scattering+for+serial+crystallography%5BTitle%5D)+AND+%22Journal+of+Visualized+Experiments%3A+JoVE%22%5BJournal%5D)">Microfluidic chips for in situ crystal x-ray diffraction and in situ dynamic light scattering for serial crystallography.</a> Journal of Visualized Experiments: JoVE. (134), e57133(2018).</li>
<li>Chandradoss, S. D., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Surface+passivation+for+single-molecule+protein+studies%5BTitle%5D)+AND+%22Journal+of+Visualized+Experiments%3A+JoVE%22%5BJournal%5D)">Surface passivation for single-molecule protein studies.</a> Journal of Visualized Experiments: JoVE. (86), e50549(2014).</li>
<li>Schaedel, L., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Microtubules+self-repair+in+response+to+mechanical+stress%5BTitle%5D)+AND+%22Nature+Materials%22%5BJournal%5D)">Microtubules self-repair in response to mechanical stress.</a> Nature Materials. 14 (11), 1156-1163 (2015).</li>
<li>Zimmermann, D., Morganthaler, A. N., Kovar, D. R., Suarez, C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((In+vitro+biochemical+characterization+of+cytokinesis+actin-binding+proteins%5BTitle%5D)+AND+%22Methods+in+Molecular+Biology%22%5BJournal%5D)">In vitro biochemical characterization of cytokinesis actin-binding proteins.</a> Methods in Molecular Biology. 1369, 151-179 (2016).</li>
<li>Funk, J., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Profilin+and+formin+constitute+a+pacemaker+system+for+robust+actin+filament+growth%5BTitle%5D)+AND+%22eLife%22%5BJournal%5D)">Profilin and formin constitute a pacemaker system for robust actin filament growth.</a> eLife. 8, 50963(2019).</li>
<li>Pandit, N. G., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Force+and+phosphate+release+from+Arp2%2F3+complex+promote+dissociation+of+actin+filament+branches%5BTitle%5D)+AND+%22Proceedings+of+the+National+Academy+of+Sciences+of+the+United+States+of+America%22%5BJournal%5D)">Force and phosphate release from Arp2/3 complex promote dissociation of actin filament branches.</a> Proceedings of the National Academy of Sciences of the United States of America. 117 (24), 13519-13528 (2020).</li>
<li>Wioland, H., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((ADF%2FCofilin+accelerates+actin+dynamics+by+severing+filaments+and+promoting+their+depolymerization+at+both+ends%5BTitle%5D)+AND+%22Current+Biology%3A+CB%22%5BJournal%5D)">ADF/Cofilin accelerates actin dynamics by severing filaments and promoting their depolymerization at both ends.</a> Current Biology: CB. 27 (13), 1956-1967 (2017).</li>
<li>Pollard, T. D., Mooseker, M. S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Direct+measurement+of+actin+polymerization+rate+constants+by+electron+microscopy+of+actin+filaments+nucleated+by+isolated+microvillus+cores%5BTitle%5D)+AND+%22The+Journal+of+Cell+Biology%22%5BJournal%5D)">Direct measurement of actin polymerization rate constants by electron microscopy of actin filaments nucleated by isolated microvillus cores.</a> The Journal of Cell Biology. 88 (3), 654-659 (1981).</li>
<li>Kovar, D. R., Harris, E. S., Mahaffy, R., Higgs, H. N., Pollard, T. D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Control+of+the+assembly+of+ATP-+and+ADP-actin+by+formins+and+profilin%5BTitle%5D)+AND+%22Cell%22%5BJournal%5D)">Control of the assembly of ATP- and ADP-actin by formins and profilin.</a> Cell. 124 (2), 423-435 (2006).</li>
<li>Jégou, A., Carlier, M. -F., Romet-Lemonne, G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Formin+mDia1+senses+and+generates+mechanical+forces+on+actin+filaments%5BTitle%5D)+AND+%22Nature+Communications%22%5BJournal%5D)">Formin mDia1 senses and generates mechanical forces on actin filaments.</a> Nature Communications. 4, 1883(2013).</li>
<li>Breitsprecher, D., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Rocket+launcher+mechanism+of+collaborative+actin+assembly+defined+by+single-molecule+imaging%5BTitle%5D)+AND+%22Science%22%5BJournal%5D)">Rocket launcher mechanism of collaborative actin assembly defined by single-molecule imaging.</a> Science. 336 (6085), 1164-1168 (2012).</li>
<li>Courtemanche, N., Lee, J. Y., Pollard, T. D., Greene, E. C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Tension+modulates+actin+filament+polymerization+mediated+by+formin+and+profilin%5BTitle%5D)+AND+%22Proceedings+of+the+National+Academy+of+Sciences+of+the+United+States+of+America%22%5BJournal%5D)">Tension modulates actin filament polymerization mediated by formin and profilin.</a> Proceedings of the National Academy of Sciences of the United States of America. 110 (24), 9752-9757 (2013).</li>
<li>Niedermayer, T., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Intermittent+depolymerization+of+actin+filaments+is+caused+by+photo-induced+dimerization+of+actin+protomers%5BTitle%5D)+AND+%22Proceedings+of+the+National+Academy+of+Sciences%22%5BJournal%5D)">Intermittent depolymerization of actin filaments is caused by photo-induced dimerization of actin protomers.</a> Proceedings of the National Academy of Sciences. 109 (27), 10769-10774 (2012).</li>
<li>Gateva, G., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Tropomyosin+isoforms+specify+functionally+distinct+actin+filament+populations+in+vitro%5BTitle%5D)+AND+%22Current+Biology%3A+CB%22%5BJournal%5D)">Tropomyosin isoforms specify functionally distinct actin filament populations in vitro.</a> Current Biology: CB. 27 (5), 705-713 (2017).</li>
<li>Aratyn, Y. S., Schaus, T. E., Taylor, E. W., Borisy, G. G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Intrinsic+dynamic+behavior+of+fascin+in+filopodia%5BTitle%5D)+AND+%22Molecular+Biology+of+the+Cell%22%5BJournal%5D)">Intrinsic dynamic behavior of fascin in filopodia.</a> Molecular Biology of the Cell. 18 (10), 3928-3940 (2007).</li>
<li>Pollard, T. D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Rate+constants+for+the+reactions+of+ATP-+and+ADP-actin+with+the+ends+of+actin+filaments%5BTitle%5D)+AND+%22The+Journal+of+Cell+Biology%22%5BJournal%5D)">Rate constants for the reactions of ATP- and ADP-actin with the ends of actin filaments.</a> The Journal of Cell Biology. 103, Pt 2 2747-2754 (1986).</li>
<li>Wioland, H., Jegou, A., Romet-Lemonne, G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Torsional+stress+generated+by+ADF%2Fcofilin+on+cross-linked+actin+filaments+boosts+their+severing%5BTitle%5D)+AND+%22Proceedings+of+the+National+Academy+of+Sciences+of+the+United+States+of+America%22%5BJournal%5D)">Torsional stress generated by ADF/cofilin on cross-linked actin filaments boosts their severing.</a> Proceedings of the National Academy of Sciences of the United States of America. 116 (7), 2595-2602 (2019).</li>
<li>Colombo, J., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((A+functional+family+of+fluorescent+nucleotide+analogues+to+investigate+actin+dynamics+and+energetics%5BTitle%5D)+AND+%22Nature+Communications%22%5BJournal%5D)">A functional family of fluorescent nucleotide analogues to investigate actin dynamics and energetics.</a> Nature Communications. 12 (1), 548(2021).</li>
<li>Spudich, J. A., Watt, S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((The+regulation+of+rabbit+skeletal+muscle+contraction.+I.+Biochemical+studies+of+the+interaction+of+the+tropomyosin-troponin+complex+with+actin+and+the+proteolytic+fragments+of+myosin%5BTitle%5D)+AND+%22The+Journal+of+Biological+Chemistry%22%5BJournal%5D)">The regulation of rabbit skeletal muscle contraction. I. Biochemical studies of the interaction of the tropomyosin-troponin complex with actin and the proteolytic fragments of myosin.</a> The Journal of Biological Chemistry. 246 (15), 4866-4871 (1971).</li>
<li>Romet-Lemonne, G., Guichard, B., Jégou, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Using+microfluidics+single+filament+assay+to+study+formin+control+of+actin+assembly%5BTitle%5D)+AND+%22Methods+in+Molecular+Biology%22%5BJournal%5D)">Using microfluidics single filament assay to study formin control of actin assembly.</a> Methods in Molecular Biology. 1805, 75-92 (2018).</li>
<li>Gieselmann, R., Kwiatkowski, D. J., Janmey, P. A., Witke, W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Distinct+biochemical+characteristics+of+the+two+human+profilin+isoforms%5BTitle%5D)+AND+%22European+Journal+of+Biochemistry%22%5BJournal%5D)">Distinct biochemical characteristics of the two human profilin isoforms.</a> European Journal of Biochemistry. 229 (3), 621-628 (1995).</li>
<li>Lin, D. C., Lin, S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Actin+polymerization+induced+by+a+motility-related+high-affinity+cytochalasin+binding+complex+from+human+erythrocyte+membrane%5BTitle%5D)+AND+%22Proceedings+of+the+National+Academy+of+Sciences+of+the+United+States+of+America%22%5BJournal%5D)">Actin polymerization induced by a motility-related high-affinity cytochalasin binding complex from human erythrocyte membrane.</a> Proceedings of the National Academy of Sciences of the United States of America. 76 (5), 2345-2349 (1979).</li>
<li>Casella, J. F., Maack, D. J., Lin, S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Purification+and+initial+characterization+of+a+protein+from+skeletal+muscle+that+caps+the+barbed+ends+of+actin+filaments%5BTitle%5D)+AND+%22The+Journal+of+Biological+Chemistry%22%5BJournal%5D)">Purification and initial characterization of a protein from skeletal muscle that caps the barbed ends of actin filaments.</a> The Journal of Biological Chemistry. 261 (23), 10915-10921 (1986).</li>
<li>Kremneva, E., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Cofilin-2+controls+actin+filament+length+in+muscle+sarcomeres%5BTitle%5D)+AND+%22Developmental+Cell%22%5BJournal%5D)">Cofilin-2 controls actin filament length in muscle sarcomeres.</a> Developmental Cell. 31 (2), 215-226 (2014).</li>
<li>Le Clainche, C., Carlier, M. -F. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Actin-based+motility+assay%5BTitle%5D)+AND+%22Current+Protocols+in+Cell+Biology%22%5BJournal%5D)">Actin-based motility assay.</a> Current Protocols in Cell Biology. , Chapter 12 1-20 (2004).</li>
<li>Vignjevic, D., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Formation+of+filopodia-like+bundles+in+vitro+from+a+dendritic+network%5BTitle%5D)+AND+%22The+Journal+of+Cell+Biology%22%5BJournal%5D)">Formation of filopodia-like bundles in vitro from a dendritic network.</a> The Journal of Cell Biology. 160 (6), 951-962 (2003).</li>
<li>Duellberg, C., Cade, N. I., Holmes, D., Surrey, T. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((The+size+of+the+EB+cap+determines+instantaneous+microtubule+stability%5BTitle%5D)+AND+%22eLife%22%5BJournal%5D)">The size of the EB cap determines instantaneous microtubule stability.</a> eLife. 5, 13470(2016).</li>
<li>Duellberg, C., Cade, N. I., Surrey, T. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Microtubule+aging+probed+by+microfluidics-assisted+tubulin+washout%5BTitle%5D)+AND+%22Molecular+Biology+of+the+Cell%22%5BJournal%5D)">Microtubule aging probed by microfluidics-assisted tubulin washout.</a> Molecular Biology of the Cell. 27 (22), 3563-3573 (2016).</li>
<li>Suzuki, E. L., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Geometrical+constraints+greatly+hinder+formin+mDia1+activity%5BTitle%5D)+AND+%22Nano+Letters%22%5BJournal%5D)">Geometrical constraints greatly hinder formin mDia1 activity.</a> Nano Letters. 20 (1), 22-32 (2020).</li>
<li>Wioland, H., Suzuki, E., Cao, L., Romet-Lemonne, G., Jegou, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((The+advantages+of+microfluidics+to+study+actin+biochemistry+and+biomechanics%5BTitle%5D)+AND+%22Journal+of+Muscle+Research+and+Cell+Motility%22%5BJournal%5D)">The advantages of microfluidics to study actin biochemistry and biomechanics.</a> Journal of Muscle Research and Cell Motility. 41 (1), 175-188 (2020).</li>
</ol>

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 lengths. The solution around the filaments is constantly renewed, exposing them to constant protein concentrations. Being able to rapidly (&#60;1 s, Figure 3D,E) switch between different protein solutions to which the filaments are exposed allows one to perform time-controlled sequential experiments, which are often instrumental for kinetic studies. Finally, the viscous drag exerted by the flowing solution on the filaments can be exploited to apply controlled mechanical stress on the filaments (Representative Results section). One should note that moderate fluid flows (Mid Flow pressure settings) bring filaments close enough to the surface (~250 nm) to efficiently image them with TIRF while generating minimal tension (&#60;1 pN)14.
Compared to classical single-filament assays, however, microfluidics will require larger volumes of protein solutions: typically a few 100 &#181;L, when a standard experiment could be done with less than 10 &#181;L. This can be a limitation when using precious proteins. Classical experiments can be used to help establish the relevant experimental conditions (e.g., absolute or relative concentrations of different proteins) before starting a series of microfluidic experiments. Another limitation, as for any other single-filament techniques in vitro, comes from the imperfect passivation of the coverslip surface. Reproducibility in coverslip cleaning and binding of the passivation layer (BSA, PEGylation, etc.) is always hard to control. A one-step passivation technique based on PEG-silane surface treatment has become the technique of choice in many laboratories7,15. As such, the effective density of filament seeds may vary between experiments by roughly twofold, even when repeated as accurately as possible. One should aim for a satisfactory range of filament surface density and be prepared to repeat the experiment if needed. Commonly encountered issues when working with microfluidics and/or single actin filaments are discussed in Supplementary File 1.
For the basic protocol presented here, one should note that spectrin-actin seeds, which can be viewed as short stabilized filaments, are randomly oriented as they stick to the surface. As a consequence, as the filaments grown from these seeds align with the flow, their portion closest to the seed will be sharply bent, each with its own angle. The length over which filaments are bent is usually very small when filaments are exposed to mid or high flows. In fact, this length will generally be smaller than the diffraction limit (~200 nm) and will thus not be easily detected. Importantly, the ABPs that are sensitive to filament curvature will bind and function differently in this highly bent region. To avoid biasing results, the simplest is to exclude this region from the analysis21.
Before we started using microfluidics to manipulate and visualize single actin filaments, it had already been used to study single DNA filaments3, which are far more flexible. This can give rise to notable differences, as the flow can dramatically unwind DNA and change its apparent length dramatically. Microfluidics can also be used, very similarly to the method presented here, to study microtubules; these are much stiffer but can nonetheless be made to align with the flow in order to measure their elongation and depolymerization, taking advantage of the rapid switch of conditions31,32, or be bent by a perpendicular flow to measure microtubule plasticity7.
We have presented here the protocol for the basic experiment, where filaments are anchored by one end only and where the flow direction in the field of view is the same throughout the experiment. These two characteristics can be varied. For example, filaments can be anchored by multiple points in order to generate a different force profile along the filament. Likewise, the flow direction can be varied (in the vicinity of the junction between the entry channels, in the flow chamber) to locally bend filaments, as the unanchored part will be pointing in a different direction than the anchored filament segment21. Filaments elongating from randomly anchored spectrin-actin seeds can also be exposed to cross-linking proteins to form bundles33 (see section 10). By combining microfluidics with other techniques (micropatterning, optical tweezers, etc.) or designing microfluidic chambers with compartments to modify flow lines, multiple configurations can be created to study specific ABP activity onto single filaments or to form small actin networks34. The number of combinations, together with the advantage and versatility of microfluidics, offers many tools to researchers in order to decipher the spatio-temporal regulation of actin networks at the molecular scale.

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 that they are anchored by one of their ends only. Similarly, when using TIRF imaging, their vertical fluctuations should be visible by changes in intensity along their length and time. Depending on the applied flow rates, one may need to adjust the TIRF penetration depth to optimize the image quality of the actin filaments acquired by TIRF.
When exposing filaments to polymerization conditions (see section 8), filament elongation should be regular (i.e., the elongation at the end of the filament is not impeded by surface interaction or permanent sticking). In addition, the measured filament barbed end elongation rate should match the expected value according to the actin concentration in the tube1,20, indicating that the tube solution has been correctly flowed up to the microfluidic chamber (Figure 4A). Similarly, when exposed to a buffer solution, filaments should depolymerize steadily at a rate that reflects their ADP-content4 (Figure 4A). When exposing already grown actin filaments to a solution of fluorescently labeled cofilin, cofilin clusters will be nucleated and grow toward both the pointed and barbed ends (Figure 4B) at a rate that is dependent on the cofilin concentration. When assessing a potential cross-linking activity of an ABP, such as fascin (Figure 4C), close-by actin filaments forming bundles will be easily detected by their higher fluorescence intensity and a change in their lateral fluctuations.
The flow of liquid applies a viscous friction force on actin filaments that are anchored to the surface of the microfluidic chamber. The friction force coefficient on F-actin is &#951; = 6.10-4 pN&#183;s/&#181;m2, expressed per filament micron length14. At intermediate flow rates, as filament height fluctuates around a constant average of 250 nm above the surface, there exists a force gradient from the free-floating end up to the filament anchoring point. One can therefore compute the applied tension at any point along the filament, using F = 6&#951;&#960;Lv, where v is the local flow velocity 250 nm above the surface (Figure 1B) and L is the downstream filament segment length (i.e., from the considered point up to the free end). For higher flow rates, the filament average height is not constant but increases linearly from the anchoring point to the free end, remains below 250 nm on average, and will vary depending on the flow rates, thus leading to a more complex tension force profile along the filament21.
<img alt="Figure 4" class="xfigimg" src="/files/ftp_upload/63891/63891fig04.jpg" /> 
Figure 4: Representative results. Typical experiments in which actin filaments are polymerized from spectrin-actin seeds and exposed to different ABPs. For the sake of clarity, only a fraction of the field of view is shown. (A) Outcome from the basic polymerization-depolymerization experiment (section 8). Filaments are polymerized with a solution of 0.8 &#181;M 10% Alexa-488 labeled G-actin, aged for 15 min to convert all subunits into ADP-actin (not shown), and depolymerized when exposed to F-buffer only. Bottom: kymographs used to quantify the polymerization and depolymerization rates. Acquired at 1 frame/5 s, 200 ms exposure time, 150 mW 488 nm laser at 9% power, TIRF (laser penetration depth 250 nm). (B) Fragmentation of single actin filaments by 500 nM mCherry-cofilin-1. Actin is labeled with ATP-ATTO48822 (yellow) and cofilin-1 is fused to mCherry (blue). Top: fraction of a field of view. Note: protein aggregates on the surface. Bottom: kymograph showing the binding of cofilin-1 to a filament (arrowheads show cofilin-1 domain nucleation events), leading to a fragmentation event (lightning symbol). Acquired at 1 frame/4 s, 200 ms exposure, 150 mW 488 nm laser at 16% and 100 mW 561 nm laser at 12% power, epifluorescence. (C) Bundling of actin filaments by fascin (section 10.5). Filaments were first polymerized with 0.8 &#181;M 5% Alexa-488 labeled G-actin and bundled with 200 nM fascin. Compared with single filaments, filament bundles appear two- to threefold brighter and not perfectly aligned with the flow. Acquired at 1 frame/10 s, 200 ms exposure, 20% 200 W Mercury lamp intensity, epifluorescence. (A-C) The background was subtracted with ImageJ&#39;s ad hoc function. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/63891/63891fig04large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Protein name</td>
			<td>species</td>
			<td>Uniprot ref (sequence)</td>
			<td>original purification protocol ref.</td>
			<td>comments</td>
		</tr>
		<tr>
			<td>actin</td>
			<td>rabbit</td>
			<td>P68135 (full length)</td>
			<td>23</td>
			<td>For fluorescent labeling, see ref 24</td>
		</tr>
		<tr>
			<td>profilin1</td>
			<td>human</td>
			<td>P07737 (full length)</td>
			<td>25</td>
			<td>see also ref 11</td>
		</tr>
		<tr>
			<td>Spectrin-actin seed</td>
			<td>human</td>
			<td>N/A</td>
			<td>26, 27</td>
			<td>see also ref 11</td>
		</tr>
		<tr>
			<td>cofilin1</td>
			<td>mouse</td>
			<td>P18760 (full length)</td>
			<td>28</td>
			<td></td>
		</tr>
		<tr>
			<td>gelsolin</td>
			<td>human</td>
			<td>P06396 (full length)</td>
			<td>29</td>
			<td></td>
		</tr>
		<tr>
			<td>mDia1 formin</td>
			<td>mouse</td>
			<td>O08808 (aa 552&#8211;1255)</td>
			<td>13</td>
			<td>more detailed protocol in ref 24</td>
		</tr>
		<tr>
			<td>fascin1</td>
			<td>human</td>
			<td>Q16658 (full length)</td>
			<td>30</td>
			<td></td>
		</tr>
	</tbody>
</table>
Table 1: Actin and actin-binding proteins23,24,25,26,27,28,29,30
<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Reagent</td>
			<td>concentration</td>
		</tr>
		<tr>
			<td>Tris-HCl pH 7.4</td>
			<td>5 mM</td>
		</tr>
		<tr>
			<td>KCl</td>
			<td>50 mM</td>
		</tr>
		<tr>
			<td>MgCl2</td>
			<td>1 mM</td>
		</tr>
		<tr>
			<td>EGTA</td>
			<td>0.2 mM</td>
		</tr>
		<tr>
			<td>ATP</td>
			<td>0.2 mM</td>
		</tr>
		<tr>
			<td>DTT</td>
			<td>10 mM</td>
		</tr>
		<tr>
			<td>DABCO</td>
			<td>1 mM</td>
		</tr>
	</tbody>
</table>
Table 2: F-buffer composition. DABCO and a relatively high concentration of DTT are used to limit photo-induced damage to filaments due to light exposure during fluorescence microscopy experiments.
<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Setting names</td>
			<td>Pressure (mBar)</td>
			<td>Flow rate (nL/min)</td>
		</tr>
		<tr>
			<td>Max pressure</td>
			<td>300</td>
			<td>~ 30 000 (in dominant channel)</td>
		</tr>
		<tr>
			<td>High pressure</td>
			<td>150</td>
			<td>~ 15 000 (in dominant channel)</td>
		</tr>
		<tr>
			<td>Mid pressure</td>
			<td>12</td>
			<td>~ 1500 (in dominant channel)</td>
		</tr>
		<tr>
			<td>&#8216;Change&#8217; pressure</td>
			<td>12 for all inlets, 
			5 for outlet</td>
			<td>~ 500 (in each inlet)</td>
		</tr>
	</tbody>
</table>
Table 3: Correspondence between applied pressures and measured flow rates. The resulting flow rates highly depend on the experimental setup. Values are given for a microfluidic chamber with a 1 cm long main channel of cross-section 20 &#181;m x 800 &#181;m (height x width), connected to each reservoir with 80 cm long PEEK tubing.
Supplementary File 1: Classical issues, causes, and solutions. They commonly encountered issues when working with microfluidics and/or single actin filaments. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/63891/Supplementary File 1.docx" target="_blank">Please click here to view a larger version of this figure.</a>

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 ...

using-microfluidics-fluorescence-microscopy-to-study-assembly

Nanyang Technological University

Research

JoVE Journal

Biochemistry

2.6K Views.  Institut Jacques Monod. We combineren fluorescentiemicroscopie met microfluïdica om te bestuderen hoe actinebindende eiwitten de assemblage en de demontage van actinefilamenten reguleren. Met microfluïdica zijn we in staat om reacties te kwantificeren die tegelijkertijd op tientallen individuele actinefilamenten plaatsvinden, terwijl we biochemische en mechanische omstandigheden nauwkeurig regelen. Vergeet niet dat oplossingen eerst parallel worden geïnjecteerd totdat ze de kamer bereiken, en vervolgens worden fi...

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 ...

Study of the Actin Cytoskeleton in Live Endothelial Cells Expressing GFP-Actin

The microvascular endothelium plays an important role as a selectively permeable barrier to fluids and solutes. The adhesive junctions between endothelial cells regulate permeability of the endothelium, and many studies have indicated the important contribution of the actin cytoskeleton to determining junctional integrity1-5. A cortical actin belt is thought to be important for the maintenance of stable junctions1, 2, 4, 5. In contrast, actin stress fibers are thought to generate centripetal tension within endothelial cells that weakens junctions2-5. Much of this theory has been based on studies in which endothelial cells are treated with inflammatory mediators known to increase endothelial permeability, and then fixing the cells and labeling F-actin for microscopic observation. However, these studies provide a very limited understanding of the role of the actin cytoskeleton because images of fixed cells provide only snapshots in time with no information about the dynamics of actin structures5. 
Live-cell imaging allows incorporation of the dynamic nature of the actin cytoskeleton into the studies of the mechanisms determining endothelial barrier integrity. A major advantage of this method is that the impact of various inflammatory stimuli on actin structures in endothelial cells can be assessed in the same set of living cells before and after treatment, removing potential bias that may occur when observing fixed specimens. Human umbilical vein endothelial cells (HUVEC) are transfected with a GFP-&beta;-actin plasmid and grown to confluence on glass coverslips. Time-lapse images of GFP-actin in confluent HUVEC are captured before and after the addition of inflammatory mediators that elicit time-dependent changes in endothelial barrier integrity. These studies enable visual observation of the fluid sequence of changes in the actin cytoskeleton that contribute to endothelial barrier disruption and restoration. 
Our results consistently show local, actin-rich lamellipodia formation and turnover in endothelial cells. The formation and movement of actin stress fibers can also be observed. An analysis of the frequency of formation and turnover of the local lamellipodia, before and after treatment with inflammatory stimuli can be documented by kymograph analyses. These studies provide important information on the dynamic nature of the actin cytoskeleton in endothelial cells that can used to discover previously unidentified molecular mechanisms important for the maintenance of endothelial barrier integrity.

Microscopic imaging of live endothelial cells expressing GFP-actin allows characterization of dynamic changes in cytoskeletal structures. Unlike techniques that use fixed specimens, this method provides a detailed assessment of temporal changes in the actin cytoskeleton in the same cells before, during, and after various physical, pharmacological, or inflammatory stimuli.

The microvascular endothelium plays an important role as a selectively permeable barrier to fluids and solutes. The adhesive junctions between endothelial ...

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 ...

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

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