We would like to thank Dina Aranovich for protein purification and labelling. G.L. is grateful to the Israel Ministry of Science, Technology and Space for the Jabotinsky PhD Scholarship. A.B.G. is grateful to the Israel Science Foundation (grant 2101/20) and to the Ministry of Science and Technology - State of Israel (grant 3-17491) for financial support.

1. Glass surface treatment and passivation:
NOTE: This section includes three major steps (see Figure 1): (i) cleaning and hydrophilization, (ii) silanization, and (iii) surface passivation.
<ol>
	<li>Cleaning and hydrophilization
	<ol>
		<li>Use Piranha solution for glass surface cleaning. 
		NOTE: Piranha solution is a mixture of 30% H2O2 and 70% H2SO4 (Table of Materials). Its prime goal is to remove organic contaminations from coverslip surfaces and expose the OH groups on the glass surface. In this manner, the glass surface becomes hydrophilic.</li>
		<li>Place 10-12 #1.5 (22 mm x 22 mm) glass coverslips (Table of Materials) in a homemade polytetrafluoroethylene holder (base area: 5.5 cm&#160;x 5.5 cm). Transfer the polytetrafluoroethylene holder into a 400 mL beaker (Table of Materials), and incubate it with 300 mL of Piranha solution for 2 h. Do this step on ice and in a chemical hood. 
		NOTE: The polytetrafluoroethylene pole screwed to the holder base is 12 cm long, longer than the beaker (11 cm), which allows for the safe transfer/pull-out of the holder into/from the Piranha solution. Since the Piranha solution can still be highly reactive and very hot, it is imperative to stop the reaction. The easiest way to stop the reaction is to pour the Piranha solution into (tap) water to achieve a 50x dilution (at least). The solution can then be safely discarded. Never keep the used Piranha solution in a closed glass bottle &#8211; this could end in a bottle explosion.</li>
		<li>Transfer the polytetrafluoroethylene holder into a clean beaker filled with fresh double-distilled water (DDW) to wash excess Piranha from the coverslips.</li>
		<li>Transfer the beaker into a sonication bath (Table of Materials), and sonicate at 80 Hz at full power for 10 min at 25 &#176;C. Repeat this step two more times with fresh DDW. Use fresh DDW at each time.</li>
	</ol>
	</li>
	<li>Silanization
	<ol>
		<li>Transfer the holder into a clean beaker (400 mL) filled with 300 mL of pure methanol (Table of Materials) and then into a sonication bath (Table of Materials). Sonicate at 80 Hz at full power for 30 min at 25 &#176;C.</li>
		<li>After sonication, transfer the holder into a silane solution prepared using 15 mL of DDW, 3.1 mL of acetic acid (Table of Materials), 340 mL of methanol, and 7.6 mL of silane ((3-Mercaptopropyl) trimethoxysilane) (Table of Materials). Seal the beaker with parafilm, and keep it in the fridge at 4 &#176;C overnight. 
		NOTE: The preparation and handling of the silane solution must be performed in a chemical hood. After the silanization step, put the used silane solution (waste) in a dedicated glass bottle within the chemical hood.</li>
		<li>On the next day, transfer the holder into a clean beaker filled with 300 mL of pure methanol for 15 s. Then, transfer the holder into a clean beaker, not before drying the bottom of the polytetrafluoroethylene holder to remove excess methanol. Transfer the beaker to the oven (Table of Materials) to dry the glass coverslips for 5 min at 120 &#176;C.</li>
	</ol>
	</li>
	<li>Surface passivation with an inert polymer
	<ol>
		<li>Achieve surface passivation by incubating the glass coverslips with an inert polymer (Table of Materials). For each experiment, take two coverslips and place them in a Petri dish coated with a parafilm layer initially cleaned with ethanol (EtOH) (DDW/EtOH: 30/70 vol/vol) (Table of Materials).</li>
		<li>Incubate each coverslip with 1 mL of a 4 mg&#183;mL&#8722;1 5 kDa molecular weight methoxy polyethylene glycol maleimide (mPEG-mal, Mw = 5 kDa) (Table of Materials) in 1x phosphate buffered saline (PBS) (Table of Materials) for 1 h at 22 &#176;C. 
		NOTE: Placing the coverslips on a hydrophobic parafilm layer confines the hydrophilic PEG polymer solution to the glass coverslip surface. The incubation time is critical for achieving optimal passivation. Use incubation times ranging between 45 min and 2 h. Above this incubation time, the glass coverslips surface can deteriorate, leading to protein adhesion and a loss of transparency.</li>
		<li>At the end of the incubation process, rinse each coverslip with 5 mL of DDW, and dry with a flow of N2 (gas). Do not dry the coverslips under a vacuum. Since the coverslips must be kept wet after passivation, immediately put 1 mL of 10 mM Tris on the pegylated surfaces. Use the coverslips within 2 h. 
		&#8203;NOTE: The various steps must be performed in a laminar flow chamber to prevent glass surface contamination.</li>
	</ol>
	</li>
</ol>
2. Protein purification
<ol>
	<li>Purify G-actin from rabbit skeletal muscle acetone powder using the gel filtration method24.
	<ol>
		<li>Actin purification takes 1 week. Keep the purified G-actin in G-buffer (5 mM Tris pH 7.8, 0.01% NaN3, 0.1 mM CaCl2, 0.2 mM ATP, and 1 mM DTT), and store it on ice. Use the solution within 2 weeks. 
		NOTE: Replace the ice every couple of days.</li>
		<li>Label the actin with a maleimide-modified fluorescent dye16 (Table of Materials). Purify the GST-fascin based on the method of Ono et al.25. Flash-freeze both proteins in liquid N2, and store at &#8722;80 &#176;C.</li>
	</ol>
	</li>
	<li>Use a standard protocol to purify myosin II from rabbit skeletal muscle26. The purified myosin II (dimers) stock buffer includes 10 mM Tris pH 7.4, 0.5 M KCl, and 35% w/v sucrose. Flash-freeze the myosin in liquid N2, and store at &#8722;80 &#176;C. 
	NOTE: The high concentration of KCl keeps the myosin motors in a dimeric form, while the high percentage of sucrose assures that the activity of the motors is not hampered upon freezing. Use a dedicated pipette for viscous fluid handling when working with myosin II stock solutions (Table of Materials).
	<ol>
		<li>Label the myosin II dimers with a fluorescent dye (Table of Materials) at pairs of engineered cysteine residues19,27. Use a chicken gizzard RLC mutant A for that purpose26. Flash-freeze the labeled myosin II in liquid N2, and store at &#8722;80 &#176;C. 
		NOTE: This labeling procedure assures that the labeled myosin is active as the native myosin II protein.</li>
	</ol>
	</li>
	<li>Use a UV-Vis spectrophotometer (Table of Materials) to measure the absorbance of the stock protein solutions, and deduce the protein concentrations with the Beer-Lambert law using the following extinction coefficients: G-actin (&#949;290 = 26,460 M&#8722;1&#183;cm&#8722;1), GST-fascin (&#949;280 = 99,330 M&#8722;1&#183;cm&#8722;1), myosin II dimer (&#949;280 = 268,800 M&#8722;1&#183;cm&#8722;1), and RLC mutant A (&#949;280 = 3,960 M&#8722;1&#183;cm&#8722;1)19.</li>
</ol>
3. Sample preparation
NOTE: Polymerize the actin monomers in the presence of large aggregates of myosin II motors and the strong passive crosslinker fascin to produce macroscopically contractile elastic actomyosin networks19,23. Adding fluorescent beads to the solution allows for tracking the solvent flow during gel contraction.
<ol>
	<li>Protein (&#34;actomyosin&#34;) solution
	<ol>
		<li>Except for G-actin, keep the proteins at &#8722;80&#176;C in small aliquots, and thaw them at 37 &#176;C before use. The actomyosin solution&#39;s total volume is 40 &#181;L.</li>
		<li>Prepare the solution by mixing 10 mM Tris pH 7.4, 2 mM MgCl2, 25 mM KCl, 1 mM ATP, an ATP regenerating system (0.5 mg&#183;mL&#8722;1 creatine kinase and 5 mM creatine phosphate), 200 &#181;M EGTA (chelates Ca2+ ions), an anti-bleaching solution (0.1 mg&#183;mL&#8722;1 glucose oxidase, 0.018 mg&#183;mL&#8722;1 catalase, and 5 mg&#183;mL&#8722;1 glucose), 5 &#181;M G-actin, 280 nM GST-fascin, and 1.67 nM myosin II. The percentage of labeled G-actin is 5% (molar percentage). 
		NOTE: Use a low percentage of labeled G-actin to avoid the saturation of the fluorescent signal at the advanced stages of gel contraction.</li>
	</ol>
	</li>
	<li>Preparing myosin II motor aggregates
	<ol>
		<li>Add the myosin motors to the protein solution in the form of aggregates. To form large myosin aggregates (150 myosin dimers/aggregate), dilute the myosin stock solution with 10 mM Tris pH 7.4 to reach a final concentration of 25 mM KCl.</li>
		<li>Keep the diluted myosin solution for 10 min at room temperature (RT), and then transfer it to ice until use. The motors are fully active for up to 2 h.</li>
	</ol>
	</li>
	<li>Measuring the solvent flow
	<ol>
		<li>To track the solvent flow across the gel pores, add the fluorescent beads to the actomyosin solution. Reduce the interaction between the beads and the actomyosin network by passivating the beads. Practically, incubate 1 &#181;L of beads (Table of Materials) with 5 &#181;M G-actin for 20 min at RT, and then remove excess G-actin by centrifugation (Table of Materials) (conditions: 6,000 x g for 5 min at 4 &#176;C).</li>
		<li>Repeat this step with 10 mg&#183;mL&#8722;1 bovine serum albumin (BSA) (Table of Materials) to block (possibly) the remaining uncoated surface. Use the beads at a final dilution of 1:10,000 vol/vol in the experiments. 
		NOTE: It is important to adapt the beads&#39; diameter to the gel pore size such that the bead/pore size ratio is &#60;1 at all stages.</li>
	</ol>
	</li>
	<li>Effect of solution viscosity
	<ol>
		<li>Control the viscosity of the solution by adding glycerol (Table of Materials) to the actomyosin solution. Adding glycerol at a weight percent ranging from 0% to 34% induces a corresponding increase in the solution viscosity from &#951;&#969; to 2.76 &#951;&#969;, where &#951;&#969;&#160;is the viscosity of water at 20 &#176;C28. 
		&#8203;NOTE: It is essential to use a dedicated pipette for viscous fluid handling when working with glycerol (Table of Materials).</li>
	</ol>
	</li>
</ol>
4. Running an experiment
<ol>
	<li>Homemade sample holder handling
	<ol>
		<li>Take one of the PEG-passivated coverslips, place a greased parafilm spacer on top of it, and place it in the sample holder. 
		NOTE: One can replace the parafilm spacer with any spacer.</li>
	</ol>
	</li>
	<li>Preparing the actomyosin solution
	<ol>
		<li>Prepare the actomyosin solution on ice in a microcentrifuge tube by incorporating the various microscopic constituents, adding the myosin II motor aggregates, and adding EGTA last. Mix the solution well. The addition of EGTA initiates actomyosin network polymerization, which sets the starting time of the experiment.</li>
	</ol>
	</li>
	<li>Put 1.1 &#181;L of that solution on the holder-mounted coverslip, and place the second coverslip on top. Screw the holder to seal the sample. This process slightly squeezes the drop, which adopts a disc-like shape. The drop diameters range between 2,800-3,000 &#181;m.</li>
	<li>Place the sample holder on the microscope, and start the acquisition. Prepare the microscope in advance to reduce the initial acquisition time. It typically takes 1-2 min after mixing to start the sample imaging.</li>
</ol>
5. Microscopy techniques
<ol>
	<li>Image the samples using an inverted fluorescent microscope (Table of Materials) controlled by dedicated software (Table of Materials).
	<ol>
		<li>Use low magnifications of 2.5x/0.075 Plan-NEOFLUAR objective (Table of Materials) to visualize the whole gel area and follow its macroscopic contraction with time.</li>
		<li>Use a 10x/0.3 Ph1 UPlanFL objective (Table of Materials) to characterize the network porosity and structure and to follow the network self-organization and contraction with time. 
		NOTE: This objective is also useful for localizing the myosin II motor aggregates within the network and following the movement of fluorescent beads across the gel pores. Higher magnifications can be used at the expense of a reduced field of view, which is even more significant in dual-color imaging mode (Table of Materials).</li>
	</ol>
	</li>
	<li>Excite the samples at 488 nm (actin/fluorescent beads) and 561 nm (myosin motor aggregates/fluorescent beads). Acquire the images at 100 ms per frame with dedicated software (Table of Materials) in streaming mode with an electron multiplying charge-coupled device (EMCCD) camera (Table of Materials). 
	NOTE: The fluorescent lamp (Table of Materials) intensity should be kept at the lowest possible value to avoid signal saturation during network contraction.</li>
	<li>Simultaneous two-color imaging
	<ol>
		<li>Use this mode for two purposes: (i) detecting the localization of the myosin motor aggregates within the actin network, and (i) characterizing the outward solvent flow inside and outside during network contraction.</li>
		<li>Excite the actin and myosin II motors at 488 nm and 561 nm, respectively, and record the images simultaneously on an EMCCD camera (Table of Materials) using a dual-emission apparatus (Table of Materials). Use the same imaging system to simultaneously image the actin gel (488 nm) and the fluorescent beads (561 nm).</li>
	</ol>
	</li>
</ol>

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J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Nonequilibrium+mechanics+and+dynamics+of+motor-activated+gels.">Nonequilibrium mechanics and dynamics of motor-activated gels.</a> Physical Review Letters. 100 (1), 018104 (2008).</li><li>Melero, C., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Light-induced+molecular+adsorption+of+proteins+using+the+PRIMO+system+for+micro-patterning+to+study+cell+responses+to+extracellular+matrix+proteins.">Light-induced molecular adsorption of proteins using the PRIMO system for micro-patterning to study cell responses to extracellular matrix proteins.</a> Journal of Visualized Experiments. (152), e60092 (2019).</li><li>Bendix, P. M., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+quantitative+analysis+of+contractility+in+active+cytoskeletal+protein+networks.">A quantitative analysis of contractility in active cytoskeletal protein networks.</a> Biophysical Journal. 94 (8), 3126-3136 (2008).</li></ol>

The authors have nothing to disclose.

Here, an in vitro approach is employed to characterize the mechanics of poroelastic actomyosin gels as a model system of the cell cytoskeleton and, more generally, of the cell cytoplasm, which has been shown to behave as a poroelastic material3,5. The rheology of the cell cytoskeleton (cytoplasm) has been characterized by a poroelastic diffusion constant, which dictates how long it takes for the intracellular cytosolic fluid to redistribute within the cell following applied mechanical stress. Cells have been proposed to use this mechanism to regulate their shape5.
Motivated by these findings, we have developed a framework to study poroelasticity in vitro. Using the proposed model system, we provide insights into how myosin contractility contributes to the emergence of a fluid flow and how network velocity and solvent velocity directionality and speed correlate in time and space. Through this, we demonstrate that the network and embedded solvent are interconnected and cannot be considered separate objects. We describe the experimental setup and provide a detailed protocol for preparing the gels and running an experiment. A detailed framework for data quantification is also presented. The proposed framework can be easily adapted to study poroelasticity in a variety of experimental setups and systems, regardless of the specificity of the experimental system investigated. 
We validate the poroelasticity of the actomyosin networks by analyzing the changes in the time scale of stress relaxation. We choose to demonstrate this by varying the viscosity of the medium by adding various amounts of glycerol to the actomyosin solution. The advantage of using glycerol is that it does not affect the porosity and structure of the networks, and, thus, the sole effect on stress relaxation is the increase in friction between the moving solution and the gel pores23. Changes in network porosity would have engendered an additional complication, as both the elastic modulus and network pore size influence the relaxation time in a non-linear manner23,31,32. 
Running successful experiments requires the passivation of the glass coverslips with an inert polymer. This step is crucial. The procedure employed here for glass passivation can be replaced with other procedures if the quality of the passivation is preserved33,34. Defective surface passivation can give rise to massive protein adhesion, which can inhibit network assembly. Glass passivation is also important to prevent the interaction of the contracting gel with the glass surfaces. This would lead to an additional friction term, in addition to the fluid-gel friction considered here, which would be much more difficult to estimate or quantify. Aside from surface passivation, successful experiments require the use of fresh actin. Moreover, the activity of the motors is also crucial, and the preparation of fresh batches of myosin should be repeated every 3-4 months. 
Overall, this experimental approach has been shown to be successful for studying poroelasticity in vitro. A limitation of the currently employed experimental procedure is how to control the initial gel dimensions more robustly. One option would be to use preformed chambers of controllable dimensions. The use of preformed chambers would also allow not only the researcher to better control the system size but also to easily change the geometry of the system.

Living cells have unique mechanical properties. Besides the ability to passively react to applied forces, they are also capable of actively generating forces in response to external stimuli1. These characteristics, which are essential for a variety of cellular processes, notably during cell motility, are primarily attributed to the mechanical and dynamic properties of the cell cytoskeleton, especially the actomyosin cytoskeleton, which is an active gel of polar actin filaments, myosin molecular motors, and accessory proteins. These actomyosin networks exhibit intrinsic self-organization and contraction properties driven by the myosin motor proteins, which crosslink the actin filaments and actively generate mechanical stresses in the network fueled by ATP hydrolysis2.
Numerous experimental and theoretical studies have been conducted to study the material properties of the cytoskeleton3. The commonly accepted view is that the cytoskeleton behaves as a viscoelastic material4. This means that on short timescales, the cytoskeleton behaves as an elastic material, and on long timescales, it behaves as a viscous fluid due to the crosslinking proteins and myosin motor detachment (and reattachment), which allows the network to dynamically turnover. In many situations, however, the viscoelastic model cannot describe the experimental results, which are more consistent with a picture describing the cytoskeleton and, more generally, the cell cytoplasm being described as a poroelastic active material5,6. Two main features characterize these types of materials. (i) The first main feature is the generation of a flow of the penetrating cytosol (the &#34;solvent&#34;) across the gel pores by contractility gradients driven by the myosin motors, which underlies processes such as cell blebbing7, motility8, and cell shape oscillations9. The emergence of such cytosolic flows can be local, for blebbing, or global, like in cell motility. In the latter case, the contractile-applied stresses at the cell rear drive the flow of the cytosolic fluid toward the cell front, which replenishes the protein pool needed for lamellipodia assembly8. (ii) The second main feature is that the relaxation of stresses is diffusive and is characterized by an effective diffusion constant, , which depends on the gel elastic modulus, gel porosity, and solvent viscosity5. The poroelastic diffusion constant determines how fast the system responds to an applied stress. Higher diffusion constants correspond to faster stress redistribution. This, in turn, determines how long it takes for the intracellular cytosolic fluid to be redistributed within the cell following applied mechanical stress, be it external or internal, such as the active contractile stresses generated by myosin motors. These examples, thus, demonstrate that the mechanics of the cytoskeleton and the cytosol are tightly coupled and cannot be treated separately3.
As cells can regulate their mechanical properties in a variety of ways, the interplay between network mechanics and fluid flow dynamics remains poorly understood. A powerful alternative approach is to use in vitro reconstituted systems that allow for full control of the various microscopic constituents and the system parameters, which renders these model systems optimal for physical analysis10,11. This approach has been successfully employed to study the impact of protein composition and system geometry on actin-based motility12,13,14,15,16,17,18, the 2D patterning of actomyosin networks19,20,21,22, and the interplay between network contractility and fluid flow dynamics of poroelastic actomyosin gels, which is the focus of this paper23.
In this manuscript, the preparation of contractile elastic actomyosin networks of controllable dimensions and material properties is discussed based on the work of Ideses et al.23. The dynamics of the contracting gel and the drained solvent are analyzed and quantified, through which it is demonstrated that these actomyosin gels can be described as a poroelastic active material. Studying the effect of solvent viscosity on stress diffusivity further confirms the poroelastic nature of these networks. The various scaling relations used for data quantification are provided. Finally, the experimental challenges, the common pitfalls, and the relevance of the experimental results to the cell cytoskeleton are also discussed.

<table><tbody><tr><td>(3-Mercaptopropyl)trimethoxysilane</td><td>Sigma-Aldrich Company</td><td>175617</td><td>Stored under Argon atmosphere at 4 &amp;deg;C&amp;nbsp;</td></tr><tr><td>Acetic acid</td><td>Bio-Lab ltd</td><td>1070521</td><td></td></tr><tr><td>Alexa-Fluor 488</td><td>Invitrogene</td><td>A10254</td><td>Diluted with DMSO, stored under Argon atmosphere at -20 &amp;deg;C&amp;nbsp;</td></tr><tr><td>Alexa-Fluor 647</td><td>Invitrogene</td><td>A20347</td><td>Diluted with DMSO, stored under Argon atmosphere at -20 &amp;deg;C&amp;nbsp;</td></tr><tr><td>BSA</td><td>Sigma -Aldrich Company</td><td>A3059</td><td>Stored at 4 &amp;deg;C&amp;nbsp;</td></tr><tr><td>Catalase</td><td>Sigma -Aldrich Company</td><td>C9322</td><td>The stock bottle is kept under dry atmosphere (silica gel) at -20 &amp;deg;C</td></tr><tr><td>Coverslips</td><td>Mezel-glaser</td><td>CG2222-1.5</td><td>Kept in milliQ-water after the Piranha treatment and used within 3 weeks</td></tr><tr><td>Creatine kinase</td><td>Roche Life Science Products</td><td>10736988001</td><td>Prepared fresh in glycine buffer, kep on ice, and used within 3 days.&amp;nbsp; The stock bottle is kept under dry atmosphere (silica gel) at 4 &amp;deg;C</td></tr><tr><td>Creatine phosphate</td><td>Roche Life Science Products</td><td>10621714001</td><td>When dissolved should be kept at -20 &amp;deg;C and used within 3 months. The stock bottle is kept under Argon atmosphere and stored at 4 &amp;deg;C</td></tr><tr><td>DTT</td><td>Roche Life Science Products</td><td>10708984001</td><td>When dissolved should be kept at -20 &amp;deg;C and used within 3 months</td></tr><tr><td>Dual view Simultaneous Imaging System&amp;nbsp;</td><td>Photometrics</td><td>DV2-CUBE</td><td></td></tr><tr><td>EGTA</td><td>MP Biomedicals</td><td>195174</td><td></td></tr><tr><td>EM-CCD Camera</td><td>Andor Technology Ltd</td><td>DV 887</td><td></td></tr><tr><td>EM-CCD Camera</td><td>Photometrics</td><td>Evolve Delta</td><td></td></tr><tr><td>Ethanol</td><td>Bio-Lab ltd</td><td>525050300</td><td></td></tr><tr><td>Flourescence Lamp</td><td>Rapp Optoelectronic</td><td></td><td></td></tr><tr><td>Fluoresbrite YG Microspheres</td><td>Polysciences</td><td>17151-10</td><td>200 nm diameter</td></tr><tr><td>Glucose</td><td>ICN Biomedicals Inc</td><td>194024</td><td>When dissolved should be kept at -20 &amp;deg;C and used within 3 months.</td></tr><tr><td>Glucose oxidase</td><td>Sigma-Aldrich Company</td><td>G7141</td><td>Kept in -20 &amp;deg;C and used within 3 months. The stock powder is kept under Argon atmosphere and kept at -20 &amp;deg;C</td></tr><tr><td>Glycerol</td><td>ICN Biomedicals Inc</td><td>800687</td><td></td></tr><tr><td>Glycine</td><td>MP Biomedicals</td><td>808822</td><td></td></tr><tr><td>Hydrogen Peroxide</td><td>Sigma-Aldrich Company</td><td>216763</td><td>Stored at 4 &amp;deg;C&amp;nbsp;</td></tr><tr><td>KCl</td><td>EMD Millipore Corp.</td><td>529552</td><td></td></tr><tr><td>Methanol</td><td>Bio-Lab ltd</td><td>1368052100</td><td></td></tr><tr><td>MgCl&lt;sub&gt;2&lt;/sub&gt;</td><td>EMD Millipore Corp.</td><td>442615</td><td></td></tr><tr><td>Microscope</td><td>Leica Microsystems</td><td>DMI3000</td><td></td></tr><tr><td>mPEG-mal</td><td>Nanocs</td><td>PG1-ML-5k&amp;nbsp;</td><td>Mw = 5000 Da. Divided to small batches by weight. Stored under Argon atmosphere at -20 &amp;deg;C</td></tr><tr><td>Nile red microspheres</td><td>Spherotech</td><td>FP-2056-2&amp;nbsp;</td><td>2300 nm diameter</td></tr><tr><td>Objective (10x)</td><td>Leica Germany</td><td>HC PL AP0</td><td>UPlanFL Numerical Aperture = 0.3</td></tr><tr><td>Objective (2.5x)</td><td>Leica Germany</td><td>506304</td><td>&amp;nbsp;Plan-NEOFLUAR Numerical Aperture = 0.075</td></tr><tr><td>Oven</td><td>WTC Binder</td><td></td><td></td></tr><tr><td>Parafilm</td><td>Amcor</td><td>PM-996</td><td></td></tr><tr><td>PBS Buffer</td><td>Sigma-Aldrich Company</td><td>P4417</td><td></td></tr><tr><td>Shutter Driver</td><td>Vincet Associates</td><td>VMM D1</td><td></td></tr><tr><td>Silica gel</td><td>Merck</td><td>1.01907.5000</td><td></td></tr><tr><td>Sonicator</td><td>Elma</td><td>Elmasonic P</td><td></td></tr><tr><td>Sulfuric acid</td><td>Carlo Erba reagents</td><td>410301</td><td></td></tr><tr><td>DV2 Dual-Channel Simultaneous-Imaging System</td><td>Photometrics</td><td></td><td></td></tr><tr><td>TRIS</td><td>MP Biomedicals</td><td>819620</td><td></td></tr><tr><td>UV-VIS Spectrophotometer</td><td>Pharmacia</td><td>Ultraspec 2100 pro</td><td></td></tr><tr><td>MICROMAN E</td><td>Gilson</td><td>FD10001</td><td>1&amp;ndash;10 uL</td></tr><tr><td>MATLAB R2017b</td><td>MathWorks</td><td></td><td>Data quantification&amp;nbsp;</td></tr><tr><td>MetaMorph&amp;nbsp;</td><td>Molecular devices</td><td></td><td>Control software of the optical imaging system; data quantification (particle tracking analysis, network mesh size)</td></tr></tbody></table>

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.

Watch this Scientific Journal Video about The Mechanics of Poro-Elastic Contractile Actomyosin Networks As a Model System of the Cell Cytoskeleton at JoVE.com

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.

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 ...

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Nanyang Technological University

Research

JoVE Journal

Bioengineering

713 Views.  Ben Gurion University of the Negev. 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.

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

Micropipette Aspiration of Substrate-attached Cells to Estimate Cell Stiffness

Growing number of studies show that biomechanical properties of individual cells play major roles in multiple cellular functions, including cell proliferation, differentiation, migration and cell-cell interactions. The two key parameters of cellular biomechanics are cellular deformability or stiffness and the ability of the cells to contract and generate force. Here we describe a quick and simple method to estimate cell stiffness by measuring the degree of membrane deformation in response to negative pressure applied by a glass micropipette to the cell surface, a technique that is called Micropipette Aspiration or Microaspiration.
Microaspiration is performed by pulling a glass capillary to create a micropipette with a very small tip (2-50 &mu;m diameter depending on the size of a cell or a tissue sample), which is then connected to a pneumatic pressure transducer and brought to a close vicinity of a cell under a microscope. When the tip of the pipette touches a cell, a step of negative pressure is applied to the pipette by the pneumatic pressure transducer generating well-defined pressure on the cell membrane. In response to pressure, the membrane is aspirated into the pipette and progressive membrane deformation or "membrane projection" into the pipette is measured as a function of time. The basic principle of this experimental approach is that the degree of membrane deformation in response to a defined mechanical force is a function of membrane stiffness. The stiffer the membrane is, the slower the rate of membrane deformation and the shorter the steady-state aspiration length.The technique can be performed on isolated cells, both in suspension and substrate-attached, large organelles, and liposomes.
Analysis is performed by comparing maximal membrane deformations achieved under a given pressure for different cell populations or experimental conditions. A "stiffness coefficient" is estimated by plotting the aspirated length of membrane deformation as a function of the applied pressure. Furthermore, the data can be further analyzed to estimate the Young's modulus of the cells (E), the most common parameter to characterize stiffness of materials. It is important to note that plasma membranes of eukaryotic cells can be viewed as a bi-component system where membrane lipid bilayer is underlied by the sub-membrane cytoskeleton and that it is the cytoskeleton that constitutes the mechanical scaffold of the membrane and dominates the deformability of the cellular envelope. This approach, therefore, allows probing the biomechanical properties of the sub-membrane cytoskeleton.

Here we describe a quick and simple method to measure cell stiffness. The general principle of this approach is to measure membrane deformation in response to well-defined negative pressure applied through a micropipette to the cell surface. This method provides a powerful tool to study biomechanical properties of substrate-attached cells.

Growing number of studies show that biomechanical properties of individual cells play major roles in multiple cellular functions, including cell ...

Live Cell Response to Mechanical Stimulation Studied by Integrated Optical and Atomic Force Microscopy

To understand the mechanism by which living cells sense mechanical forces, and how they respond and adapt to their environment, a new technology able to investigate cells behavior at sub-cellular level with high spatial and temporal resolution was developed. Thus, an atomic force microscope (AFM) was integrated with total internal reflection fluorescence (TIRF) microscopy and fast-spinning disk (FSD) confocal microscopy. The integrated system is broadly applicable across a wide range of molecular dynamic studies in any adherent live cells, allowing direct optical imaging of cell responses to mechanical stimulation in real-time. Significant rearrangement of the actin filaments and focal adhesions was shown due to local mechanical stimulation at the apical cell surface that induced changes into the cellular structure throughout the cell body. These innovative techniques will provide new information for understanding live cell restructuring and dynamics in response to mechanical force. A detailed protocol and a representative data set that show live cell response to mechanical stimulation are presented.

This paper aims to instruct the reader in the operation of an integrated atomic force-optical imaging microscope for mechanical stimulation of live cells in culture. A step-by-step protocol is presented. A representative data set that shows live cell response to mechanical stimulation is presented.

To understand the mechanism by which living cells sense mechanical forces, and how they respond and adapt to their environment, a new technology able to ...

Biology

A Typical Workflow to Simulate Cytoskeletal Systems

Many cytoskeletal systems are now sufficiently well known to permit their precise quantitative modeling. Microtubule and actin filaments are well characterized, and the associated proteins are often known, as well as their abundance and the interactions between these elements. Thus, computer simulations can be used to investigate the collective behavior of the system precisely, in a way that is complementary to experiments. Cytosim is an Open Source cytoskeleton simulation suite designed to handle large systems of flexible filaments with associated proteins such as molecular motors. It also offers the possibility to simulate passive crosslinkers, diffusible crosslinkers, nucleators, cutters, and discrete versions of the motors that only step on unoccupied lattice sites on a filament. Other objects complement the filaments by offering spherical or more complicated geometry that can be used to represent chromosomes, the nucleus, or vesicles in the cell.
Cytosim offers simple command-line tools for running a simulation and displaying its results, which are versatile and do not require programming skills. In this workflow, step-by-step instructions are given to i) install the necessary environment on a new computer, ii) configure Cytosim to simulate the contraction of a 2D actomyosin network, and iii) produce a visual representation of the system. Next, the system is probed by systematically varying a key parameter: the number of crosslinkers. Finally, the visual representation of the system is complemented by the numerical quantification of contractility to view, in a graph, how contractility depends on the composition of the system. Overall, these different steps constitute a typical workflow that can be applied with few modifications to tackle many other problems in the cytoskeletal field.

This protocol demonstrates how to use Cytosim, an Open Source cytoskeleton simulation, to investigate the behavior of a network of filaments connected by molecular motors and passive crosslinkers. A generic workflow with step-by-step instructions is followed to vary the number of crosslinkers and plot the resulting network contractility.

Many cytoskeletal systems are now sufficiently well known to permit their precise quantitative modeling. Microtubule and actin filaments are well ...

Developmental Biology

Optogenetic Inhibition of Rho1-Mediated Actomyosin Contractility in Drosophila

Optogenetic Inhibition of Rho1-Mediated Actomyosin Contractility Coupled with Measurement of Epithelial Tension in Drosophila Embryos

Contractile forces generated by actin and non-muscle myosin II ("actomyosin contractility") are critical for morphological changes of cells and tissues at multiple length scales, such as cell division, cell migration, epithelial folding, and branching morphogenesis. An in-depth understanding of the role of actomyosin contractility in morphogenesis requires approaches that allow the rapid inactivation of actomyosin, which is difficult to achieve using conventional genetic or pharmacological approaches. The presented protocol demonstrates the use of a CRY2-CIBN based optogenetic dimerization system, Opto-Rho1DN, to inhibit actomyosin contractility in Drosophila embryos with precise temporal and spatial controls. In this system, CRY2 is fused to the dominant negative form of Rho1 (Rho1DN), whereas CIBN is anchored to the plasma membrane. Blue light-mediated dimerization of CRY2 and CIBN results in rapid translocation of Rho1DN from the cytoplasm to the plasma membrane, where it inactivates actomyosin by inhibiting endogenous Rho1. In addition, this article presents a detailed protocol for coupling Opto-Rho1DN-mediated inactivation of actomyosin with laser ablation to investigate the role of actomyosin in generating epithelial tension during Drosophila ventral furrow formation. This protocol can be applied to many other morphological processes that involve actomyosin contractility in Drosophila embryos with minimal modifications. Overall, this optogenetic tool is a powerful approach to dissect the function of actomyosin contractility in controlling tissue mechanics during dynamic tissue remodeling.

Author Spotlight: Optogenetic Inhibition of Rho1-Mediated Actomyosin Contractility Coupled with Measurement of Epithelial Tension in Drosophila Embryos  

Actomyosin contractility plays an important role in cell and tissue morphogenesis. However, it is challenging to manipulate actomyosin contractility in vivo acutely. This protocol describes an optogenetic system that rapidly inhibits Rho1-mediated actomyosin contractility in Drosophila embryos, revealing the immediate loss of epithelial tension after the inactivation of actomyosin in vivo.

Contractile forces generated by actin and non-muscle myosin II ("actomyosin contractility") are critical for morphological changes of cells and tissues at ...

Simplified, High-throughput Analysis of Single-cell Contractility using Micropatterned Elastomers

Cellular contractile force generation is a fundamental trait shared by virtually all cells. These contractile forces are crucial to proper development, function at both the cellular and tissue levels,and regulate the mechanical systems in the body. Numerous biological processes are force-dependent, including motility, adhesion, and division of single-cells, as well as contraction and relaxation of organs such as the heart, bladder, lungs, intestines, and uterus. Given its importance in maintaining proper physiological function, cellular contractility can also drive disease processes when exaggerated or disrupted. Asthma, hypertension, preterm labor, fibrotic scarring, and underactive bladder are all examples of mechanically driven disease processes that could potentially be alleviated with proper control of cellular contractile force. Here, we present a comprehensive protocol for utilizing a novel microplate-based contractility assay technology known as fluorescently labeled elastomeric contractible surfaces (FLECS), that provides simplified and intuitive analysis of single-cell contractility in a massively scaled manner. Herein, we provide a step-wise protocol for obtaining two six-point dose-response curves describing the effects of two contractile inhibitors on the contraction of primary human bladder smooth muscle cells in a simple procedure utilizing just a single FLECS assay microplate, to demonstrate proper technique to users of the method. Using FLECS Technology, all researchers with basic biological laboratories and fluorescent microscopy systems gain access to studying this fundamental but difficult-to-quantify functional cell phenotype, effectively lowering the entry barrier into the field of force biology and phenotypic screening of contractile cell force.

This work presents a flexible protocol for utilizing fluorescently labeled elastomeric contractible surfaces (FLECS) Technology in microwell format for simplified, hands-off quantification of single-cell contractile forces based on visualized displacements of fluorescent protein micropatterns.

Cellular contractile force generation is a fundamental trait shared by virtually all cells. These contractile forces are crucial to proper development, ...

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 ...

Quantifying Cytoskeleton Dynamics Using Differential Dynamic Microscopy

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

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

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

In Vitro Reconstitution of the Actin Cytoskeleton Inside Giant Unilamellar Vesicles

The actin cytoskeleton, the principal mechanical machinery in the cell, mediates numerous essential physical cellular activities, including cell deformation, division, migration, and adhesion. However, studying the dynamics and structure of the actin network in vivo is complicated by the biochemical and genetic regulation within live cells. To build a minimal model devoid of intracellular biochemical regulation, actin is encapsulated inside giant unilamellar vesicles (GUVs, also called liposomes). The biomimetic liposomes are cell-sized and facilitate a quantitative insight into the mechanical and dynamical properties of the cytoskeleton network, opening a viable route for bottom-up synthetic biology. To generate liposomes for encapsulation, the inverted emulsion method (also referred to as the emulsion transfer method) is utilized, which is one of the most successful techniques for encapsulating complex solutions into liposomes to prepare various cell-mimicking systems. With this method, a mixture of proteins of interest is added to the inner buffer, which is later emulsified in a phospholipid-containing mineral oil solution to form monolayer lipid droplets. The desired liposomes are generated from monolayer lipid droplets crossing a lipid/oil-water interface. This method enables the encapsulation of concentrated actin polymers into the liposomes with desired lipid components, paving the way for in vitro reconstitution of a biomimicking cytoskeleton network.

In Vitro Reconstitution of the Actin Cytoskeleton Inside Giant Unilamellar Vesicles

In this manuscript, we demonstrate the experimental techniques to encapsulate the F-actin cytoskeleton into giant unilamellar lipid vesicles (also called liposomes), and the method to form a cortex-biomimicking F-actin layer at the inner leaflet of the liposome membrane.

The actin cytoskeleton, the principal mechanical machinery in the cell, mediates numerous essential physical cellular activities, including cell ...

Reconstitution of Membrane-Tethered Minimal Actin Cortices on Supported Lipid Bilayers

The surface of a living cell provides a versatile active platform for numerous cellular processes, which arise from the interplay of the plasma membrane with the underlying actin cortex. In the past decades, reconstituted, minimal systems based on supported lipid bilayers in combination with actin filament networks have proven to be very instrumental in unraveling basic mechanisms and consequences of membrane-tethered actin networks, as well as in studying the functions of individual membrane-associated proteins. Here, we describe how to reconstitute such active composite systems in vitro that consist of fluid supported lipid bilayers coupled via membrane-associated actin-binding proteins to dynamic actin filaments and myosin motors that can be readily observed via total internal reflection fluorescence microscopy. An open-chamber design allows one to assemble the system in a step-by-step manner and to systematically control many parameters such as linker protein concentration, actin concentration, actin filament length, actin/myosin ratio, as well as ATP levels. Finally, we discuss how to control the quality of the system, how to detect and troubleshoot commonly occurring problems, and some limitations of this system in comparison with the living cell surface.

This protocol describes the formation of supported lipid bilayers and the addition of cytoskeletal filaments and motor proteins to study the dynamics of reconstituted, membrane-tethered cytoskeletal networks using fluorescence microscopy.

The surface of a living cell provides a versatile active platform for numerous cellular processes, which arise from the interplay of the plasma membrane ...

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 ...

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

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Live Cell Response to Mechanical Stimulation Studied by Integrated Optical and Atomic Force Microscopy

A Typical Workflow to Simulate Cytoskeletal Systems

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Simplified, High-throughput Analysis of Single-cell Contractility using Micropatterned Elastomers

Probing Myosin Ensemble Mechanics in Actin Filament Bundles Using Optical Tweezers

Quantifying Cytoskeleton Dynamics Using Differential Dynamic Microscopy

In Vitro Reconstitution of the Actin Cytoskeleton Inside Giant Unilamellar Vesicles

Reconstitution of Membrane-Tethered Minimal Actin Cortices on Supported Lipid Bilayers

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