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 ce...

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.

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			<td>(3-Mercaptopropyl)trimethoxysilane</td>
			<td>Sigma-Aldrich Company</td>
			<td>175617</td>
			<td>Stored under Argon atmosphere at 4 &#176;C&#160;</td>
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			<td>Acetic acid</td>
			<td>Bio-Lab ltd</td>
			<td>1070521</td>
			<td></td>
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			<td>Alexa-Fluor 488</td>
			<td>Invitrogene</td>
			<td>A10254</td>
			<td>Diluted with DMSO, stored under Argon atmosphere at -20 &#176;C&#160;</td>
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			<td>Alexa-Fluor 647</td>
			<td>Invitrogene</td>
			<td>A20347</td>
			<td>Diluted with DMSO, stored under Argon atmosphere at -20 &#176;C&#160;</td>
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			<td>BSA</td>
			<td>Sigma -Aldrich Company</td>
			<td>A3059</td>
			<td>Stored at 4 &#176;C&#160;</td>
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			<td>Catalase</td>
			<td>Sigma -Aldrich Company</td>
			<td>C9322</td>
			<td>The stock bottle is kept under dry atmosphere (silica gel) at -20 &#176;C</td>
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			<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>
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			<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.&#160; The stock bottle is kept under dry atmosphere (silica gel) at 4 &#176;C</td>
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			<td>Creatine phosphate</td>
			<td>Roche Life Science Products</td>
			<td>10621714001</td>
			<td>When dissolved should be kept at -20 &#176;C and used within 3 months. The stock bottle is kept under Argon atmosphere and stored at 4 &#176;C</td>
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			<td>DTT</td>
			<td>Roche Life Science Products</td>
			<td>10708984001</td>
			<td>When dissolved should be kept at -20 &#176;C and used within 3 months</td>
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			<td>Dual view Simultaneous Imaging System&#160;</td>
			<td>Photometrics</td>
			<td>DV2-CUBE</td>
			<td></td>
		</tr>
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			<td>EGTA</td>
			<td>MP Biomedicals</td>
			<td>195174</td>
			<td></td>
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			<td>EM-CCD Camera</td>
			<td>Andor Technology Ltd</td>
			<td>DV 887</td>
			<td></td>
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			<td>EM-CCD Camera</td>
			<td>Photometrics</td>
			<td>Evolve Delta</td>
			<td></td>
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			<td>Ethanol</td>
			<td>Bio-Lab ltd</td>
			<td>525050300</td>
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			<td>Flourescence Lamp</td>
			<td>Rapp Optoelectronic</td>
			<td></td>
			<td></td>
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			<td>Fluoresbrite YG Microspheres</td>
			<td>Polysciences</td>
			<td>17151-10</td>
			<td>200 nm diameter</td>
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		<tr>
			<td>Glucose</td>
			<td>ICN Biomedicals Inc</td>
			<td>194024</td>
			<td>When dissolved should be kept at -20 &#176;C and used within 3 months.</td>
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		<tr>
			<td>Glucose oxidase</td>
			<td>Sigma-Aldrich Company</td>
			<td>G7141</td>
			<td>Kept in -20 &#176;C and used within 3 months. The stock powder is kept under Argon atmosphere and kept at -20 &#176;C</td>
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			<td>Glycerol</td>
			<td>ICN Biomedicals Inc</td>
			<td>800687</td>
			<td></td>
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			<td>Glycine</td>
			<td>MP Biomedicals</td>
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			<td>Sigma-Aldrich Company</td>
			<td>216763</td>
			<td>Stored at 4 &#176;C&#160;</td>
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			<td>EMD Millipore Corp.</td>
			<td>529552</td>
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			<td>Bio-Lab ltd</td>
			<td>1368052100</td>
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			<td>MgCl2</td>
			<td>EMD Millipore Corp.</td>
			<td>442615</td>
			<td></td>
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			<td>Microscope</td>
			<td>Leica Microsystems</td>
			<td>DMI3000</td>
			<td></td>
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			<td>mPEG-mal</td>
			<td>Nanocs</td>
			<td>PG1-ML-5k&#160;</td>
			<td>Mw = 5000 Da. Divided to small batches by weight. Stored under Argon atmosphere at -20 &#176;C</td>
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			<td>Nile red microspheres</td>
			<td>Spherotech</td>
			<td>FP-2056-2&#160;</td>
			<td>2300 nm diameter</td>
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			<td>Objective (10x)</td>
			<td>Leica Germany</td>
			<td>HC PL AP0</td>
			<td>UPlanFL Numerical Aperture = 0.3</td>
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			<td>Objective (2.5x)</td>
			<td>Leica Germany</td>
			<td>506304</td>
			<td>&#160;Plan-NEOFLUAR Numerical Aperture = 0.075</td>
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			<td>Oven</td>
			<td>WTC Binder</td>
			<td></td>
			<td></td>
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			<td>Parafilm</td>
			<td>Amcor</td>
			<td>PM-996</td>
			<td></td>
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			<td>PBS Buffer</td>
			<td>Sigma-Aldrich Company</td>
			<td>P4417</td>
			<td></td>
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			<td>Vincet Associates</td>
			<td>VMM D1</td>
			<td></td>
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			<td>Silica gel</td>
			<td>Merck</td>
			<td>1.01907.5000</td>
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			<td>Sonicator</td>
			<td>Elma</td>
			<td>Elmasonic P</td>
			<td></td>
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			<td>Sulfuric acid</td>
			<td>Carlo Erba reagents</td>
			<td>410301</td>
			<td></td>
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			<td>DV2 Dual-Channel Simultaneous-Imaging System</td>
			<td>Photometrics</td>
			<td></td>
			<td></td>
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			<td>TRIS</td>
			<td>MP Biomedicals</td>
			<td>819620</td>
			<td></td>
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			<td>UV-VIS Spectrophotometer</td>
			<td>Pharmacia</td>
			<td>Ultraspec 2100 pro</td>
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			<td>MICROMAN E</td>
			<td>Gilson</td>
			<td>FD10001</td>
			<td>1&#8211;10 uL</td>
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			<td>MATLAB R2017b</td>
			<td>MathWorks</td>
			<td></td>
			<td>Data quantification&#160;</td>
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			<td>MetaMorph&#160;</td>
			<td>Molecular devices</td>
			<td></td>
			<td>Control software of the optical imaging system; data quantification (particle tracking analysis, network mesh size)</td>
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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.

Two glass coverslips are used per experiment. The glass coverslips are cleaned and passivated with PEG polymers. Passivation is essential for preventing the solubilized proteins from adhering to the glass surfaces at the early experimental stages and for minimizing the interaction of the contracting network with the glass walls. Failure to achieve good passivation can lead to inefficient contraction and, in extreme cases, can even inhibit actin network formation.
Figure 1<...

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 ...

the-mechanics-poro-elastic-contractile-actomyosin-networks-as-model

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Bioengineering

701 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

Rotating Cell Culture Systems for Human Cell Culture: Human Trophoblast Cells as a Model

The field of human trophoblast research aids in understanding the complex environment established during placentation. Due to the nature of these studies, human in vivo experimentation is impossible. A combination of primary cultures, explant cultures and trophoblast cell lines1 support our understanding of invasion of the uterine wall2 and remodeling of uterine spiral arteries3,4 by extravillous trophoblast cells (EVTs), which is required for successful establishment of pregnancy. Despite the wealth of knowledge gleaned from such models, it is accepted that in vitro cell culture models using EVT-like cell lines display altered cellular properties when compared to their in vivo counterparts5,6. Cells cultured in the rotating cell culture system (RCCS) display morphological, phenotypic, and functional properties of EVT-like cell lines that more closely mimic differentiating in utero EVTs, with increased expression of genes mediating invasion (e.g. matrix metalloproteinases (MMPs)) and trophoblast differentiation7,8,9. The Saint Georges Hospital Placental cell Line-4 (SGHPL-4) (kindly donated by Dr. Guy Whitley and Dr. Judith Cartwright) is an EVT-like cell line that was used for testing in the RCCS.
The design of the RCCS culture vessel is based on the principle that organs and tissues function in a three-dimensional (3-D) environment. Due to the dynamic culture conditions in the vessel, including conditions of physiologically relevant shear, cells grown in three dimensions form aggregates based on natural cellular affinities and differentiate into organotypic tissue-like assemblies10,11,12 . The maintenance of a fluid orbit provides a low-shear, low-turbulence environment similar to conditions found in vivo. Sedimentation of the cultured cells is countered by adjusting the rotation speed of the RCCS to ensure a constant free-fall of cells. Gas exchange occurs through a permeable hydrophobic membrane located on the back of the bioreactor. Like their parental tissue in vivo, RCCS-grown cells are able to respond to chemical and molecular gradients in three dimensions (i.e. at their apical, basal, and lateral surfaces) because they are cultured on the surface of porous microcarrier beads. When grown as two-dimensional monolayers on impermeable surfaces like plastic, cells are deprived of this important communication at their basal surface. Consequently, the spatial constraints imposed by the environment profoundly affect how cells sense and decode signals from the surrounding microenvironment, thus implying an important role for the 3-D milieu13.
We have used the RCCS to engineer biologically meaningful 3-D models of various human epithelial tissues7,14,15,16. Indeed, many previous reports have demonstrated that cells cultured in the RCCS can assume physiologically relevant phenotypes that have not been possible with other models10,17-21. In summary, culture in the RCCS represents an easy, reproducible, high-throughput platform that provides large numbers of differentiated cells that are amenable to a variety of experimental manipulations. 
In the following protocol, using EVTs as an example, we clearly describe the steps required to three-dimensionally culture adherent cells in the RCCS.

Traditional, two dimensional cell culture techniques often result in altered characteristics with respect to differentiation markers, cytokines and growth factors. Three-dimensional cell culture in the rotating cell culture system (RCCS) reestablishes expression of many of these factors as shown here with an extravillous trophoblast cell line.

The field of human trophoblast research aids in  understanding the complex environment established during placentation. Due to the  nature of these ...

Alginate Microcapsule as a 3D Platform for Propagation and Differentiation of Human Embryonic Stem Cells (hESC) to Different Lineages

Human embryonic stem cells (hESC) are emerging as an attractive alternative source for cell replacement therapy since they can be expanded in culture indefinitely and differentiated to any cell types in the body. Various types of biomaterials have also been used in stem cell cultures to provide a microenvironment mimicking the stem cell niche1-3. The latter is important for promoting cell-to-cell interaction, cell proliferation, and differentiation into specific lineages as well as tissue organization by providing a three-dimensional (3D) environment4 such as encapsulation. The principle of cell encapsulation involves entrapment of living cells within the confines of semi-permeable membranes in 3D cultures2. These membranes allow for the exchange of nutrients, oxygen and stimuli across the membranes, whereas antibodies and immune cells from the host that are larger than the capsule pore size are excluded5. Here, we present an approach to culture and differentiate hESC DA neurons in a 3D microenvironment using alginate microcapsules. We have modified the culture conditions2 to enhance the viability of encapsulated hESC. We have previously shown that the addition of p160-Rho-associated coiled-coil kinase (ROCK) inhibitor, Y-27632 and human fetal fibroblast-conditioned serum replacement medium (hFF-CM) to the 3D platform significantly enhanced the viability of encapsulated hESC in which the cells expressed definitive endoderm marker genes1. We have now used this 3D platform for the propagation of hESC and efficient differentiation to DA neurons. Protein and gene expression analyses after the final stage of DA neuronal differentiation showed an increased expression of tyrosine hydroxylase (TH), a marker for DA neurons, &gt;100 folds after 2 weeks. We hypothesized that our 3D platform using alginate microcapsules may be useful to study the proliferation and directed differentiation of hESC to various lineages. This 3D system also allows the separation of feeder cells from hESC during the process of differentiation and also has potential for immune-isolation during transplantation in the future.

Alginate Microcapsule as a 3D Platform for Propagation and Differentiation of Human Embryonic Stem Cells hESC to Different Lineages

We have optimized a microencapsulation technique as an effective 3D platform for propagation and differentiation of embryonic stem cells to endoderm and dopaminergic (DA) neurons. It also provides an opportunity for immune-isolation of cells from the host during transplantation. This platform can be adapted for other cell types.

Human embryonic stem cells (hESC) are emerging as an attractive alternative source for cell replacement therapy since they can be expanded in culture ...

Magnetic Resonance Elastography Methodology for the Evaluation of Tissue Engineered Construct Growth

Traditional mechanical testing often results in the destruction of the sample, and in the case of long term tissue engineered construct studies, the use of destructive assessment is not acceptable. A proposed alternative is the use of an imaging process called magnetic resonance elastography. Elastography is a nondestructive method for determining the engineered outcome by measuring local mechanical property values (i.e., complex shear modulus), which are essential markers for identifying the structure and functionality of a tissue. As a noninvasive means for evaluation, the monitoring of engineered constructs with imaging modalities such as magnetic resonance imaging (MRI) has seen increasing interest in the past decade1. For example, the magnetic resonance (MR) techniques of diffusion and relaxometry have been able to characterize the changes in chemical and physical properties during engineered tissue development2. The method proposed in the following protocol uses microscopic magnetic resonance elastography (&mu;MRE) as a noninvasive MR based technique for measuring the mechanical properties of small soft tissues3. MRE is achieved by coupling a sonic mechanical actuator with the tissue of interest and recording the shear wave propagation with an MR scanner4. Recently, &mu;MRE has been applied in tissue engineering to acquire essential growth information that is traditionally measured using destructive mechanical macroscopic techniques5. In the following procedure, elastography is achieved through the imaging of engineered constructs with a modified Hahn spin-echo sequence coupled with a mechanical actuator. As shown in Figure 1, the modified sequence synchronizes image acquisition with the transmission of external shear waves; subsequently, the motion is sensitized through the use of oscillating bipolar pairs. Following collection of images with positive and negative motion sensitization, complex division of the data produce a shear wave image. Then, the image is assessed using an inversion algorithm to generate a shear stiffness map6. The resulting measurements at each voxel have been shown to strongly correlate (R2&gt;0.9914) with data collected using dynamic mechanical analysis7. In this study, elastography is integrated into the tissue development process for monitoring human mesenchymal stem cell (hMSC) differentiation into adipogenic and osteogenic constructs as shown in Figure 2.

The procedure demonstrates the methodology of magnetic resonance elastography for monitoring the engineered outcome of adipose and osteogenic tissue engineered constructs through noninvasive local assessment of the mechanical properties using microscopic magnetic resonance elastography (&mu;MRE).

Traditional mechanical testing often results in the destruction of the sample, and in the case of long term tissue engineered construct studies, the use ...

Monitoring the Wall Mechanics During Stent Deployment in a Vessel

Clinical trials have reported different restenosis rates for various stent designs1. It is speculated that stent-induced strain concentrations on the arterial wall lead to tissue injury, which initiates restenosis2-7. This hypothesis needs further investigations including better quantifications of non-uniform strain distribution on the artery following stent implantation. A non-contact surface strain measurement method for the stented artery is presented in this work. ARAMIS stereo optical surface strain measurement system uses two optical high speed cameras to capture the motion of each reference point, and resolve three dimensional strains over the deforming surface8,9. As a mesh stent is deployed into a latex vessel with a random contrasting pattern sprayed or drawn on its outer surface, the surface strain is recorded at every instant of the deformation. The calculated strain distributions can then be used to understand the local lesion response, validate the computational models, and formulate hypotheses for further in vivo study.

Stent-induced arterial strain distributions are characterized using an optical surface strain measurement system. This visualization technique is used to gain insights into the impact of stent implantation on the host vessel.

Clinical trials have reported different restenosis rates for various stent designs1. It is speculated that stent-induced strain concentrations on the ...

Robotic Production of Cancer Cell Spheroids with an Aqueous Two-phase System for Drug Testing

Cancer cell spheroids present a relevant in vitro model of avascular tumors for anti-cancer drug testing applications. A detailed protocol for producing both mono-culture and co-culture spheroids in a high throughput 96-well plate format is described in this work. This approach utilizes an aqueous two-phase system to confine cells into a drop of the denser aqueous phase immersed within the second aqueous phase. The drop rests on the well surface and keeps cells in close proximity to form a single spheroid. This technology has been adapted to a robotic liquid handler to produce size-controlled spheroids and expedite the process of spheroid production for compound screening applications. Spheroids treated with a clinically-used drug show reduced cell viability with increase in the drug dose. The use of a standard micro-well plate for spheroid generation makes it straightforward to analyze viability of cancer cells of drug-treated spheroids with a micro-plate reader. This technology is straightforward to implement both robotically and with other liquid handling tools such as manual pipettes.

A protocol for robotic printing of cancer cell spheroids in a high throughput 96-well plate format using an aqueous two-phase system is presented.

Cancer cell spheroids present a relevant in vitro model of avascular tumors for anti-cancer drug testing applications. A detailed protocol for producing ...

Fabrication of Extracellular Matrix-derived Foams and Microcarriers as Tissue-specific Cell Culture and Delivery Platforms

Cell function is mediated by interactions with the extracellular matrix (ECM), which has complex tissue-specific composition and architecture. The focus of this article is on the methods for fabricating ECM-derived porous foams and microcarriers for use as biologically-relevant substrates in advanced 3D in vitro cell culture models or as pro-regenerative scaffolds and cell delivery systems for tissue engineering and regenerative medicine. Using decellularized tissues or purified insoluble collagen as a starting material, the techniques can be applied to synthesize a broad array of tissue-specific bioscaffolds with customizable geometries. The approach involves mechanical processing and mild enzymatic digestion to yield an ECM suspension that is used to fabricate the three-dimensional foams or microcarriers through controlled freezing and lyophilization procedures. These pure ECM-derived scaffolds are highly porous, yet stable without the need for chemical crosslinking agents or other additives that may negatively impact cell function. The scaffold properties can be tuned to some extent by varying factors such as the ECM suspension concentration, mechanical processing methods, or synthesis conditions. In general, the scaffolds are robust and easy to handle, and can be processed as tissues for most standard biological assays, providing a versatile and user-friendly 3D cell culture platform that mimics the native ECM composition. Overall, these straightforward methods for fabricating customized ECM-derived foams and microcarriers may be of interest to both biologists and biomedical engineers as tissue-specific cell-instructive platforms for in vitro and in vivo applications.

The tissue-specific extracellular matrix (ECM) is a key mediator of cell function. This article describes methods for synthesizing pure ECM-derived foams and microcarriers that are stable in culture without the need for chemical crosslinking for applications in advanced 3D in vitro cell culture models or as pro-regenerative bioscaffolds.

Cell function is mediated by interactions with the extracellular matrix (ECM), which has complex tissue-specific composition and architecture. The focus ...

Simulating the Mechanics of Lens Accommodation via a Manual Lens Stretcher

The goal of this protocol is to mimic the biomechanics of physiological accommodation in a cost-efficient, practical manner. Accommodation is achieved through the contraction of the ciliary body and relaxation of zonule fibers, which results in the thickening of the lens necessary for near vision. Here, we present a novel, simple method in which accommodation is replicated by tensing the zonules connected to the lens capsule via a manual lens stretcher (MLS). This method monitors the radial stretching achieved by a lens when subjected to a consistent force and allows for a comparison of accommodating lenses, which can be stretched, to non-accommodating lenses, which cannot be stretched. Importantly, the stretcher couples to the zonules directly, and not to the sclera of the eye, thus only requiring the lens, zonules, and ciliary body rather than the entire globe sample. This difference can significantly decrease the cost of acquiring donor cadaver lenses by about 62% compared to acquiring an entire globe.

We present an efficient method of studying lens accommodation by using a manual lens stretcher. The protocol mimics physiological accommodation by pulling the zonules connected around the lens capsule, thereby, stretching the lens.

The goal of this protocol is to mimic the biomechanics of physiological accommodation in a cost-efficient, practical manner. Accommodation is achieved ...

Recombinant Collagen I Peptide Microcarriers for Cell Expansion and Their Potential Use As Cell Delivery System in a Bioreactor Model

Tissue engineering is a promising field, focused on developing solutions for the increasing demand on tissues and organs regarding transplantation purposes. The process to generate such tissues is complex, and includes an appropriate combination of specific cell types, scaffolds, and physical or biochemical stimuli to guide cell growth and differentiation. Microcarriers represent an appealing tool to expand cells in a three-dimensional (3D) microenvironment, since they provide higher surface-to volume ratios and mimic more closely the in vivo situation compared to traditional two-dimensional methods. The vascular system, supplying oxygen and nutrients to the cells and ensuring waste removal, constitutes an important building block when generating engineered tissues. In fact, most constructs fail after being implanted due to lacking vascular support. In this study, we present a protocol for endothelial cell expansion on recombinant collagen-based microcarriers under dynamic conditions in spinner flask and bioreactors, and we explain how to determine in this setting cell viability and functionality. In addition, we propose a method for cell delivery for vascularization purposes without additional detachment steps necessary. Furthermore, we provide a strategy to evaluate the cell vascularization potential in a perfusion bioreactor on a decellularized&#160;biological matrix. We believe that the use of the presented methods could lead to the development of new cell-based therapies for a large range of tissue engineering applications in the clinical practice.

We propose a cell expansion protocol on macroporous microcarriers and their use as delivery system in a perfusion bioreactor to seed a decellularized tissue matrix. We also include different techniques to determine cell proliferation and viability of cells cultured on microcarriers. Furthermore, we demonstrate functionality of cells after bioreactor cultures.

Tissue engineering is a promising field, focused on developing solutions for the increasing demand on tissues and organs regarding transplantation ...

Spontaneous Formation and Rearrangement of Artificial Lipid Nanotube Networks as a Bottom-Up Model for Endoplasmic Reticulum

We present a convenient method to form a bottom-up structural organelle model for the endoplasmic reticulum (ER). The model consists of highly dense lipidic nanotubes that are, in terms of morphology and dynamics, reminiscent of ER. The networks are derived from phospholipid double bilayer membrane patches adhering to a transparent Al2O3 substrate. The adhesion is mediated by Ca2+ in the ambient buffer. Subsequent depletion of Ca2+ by means of BAPTA/EDTA causes retraction of the membrane, resulting in spontaneous lipid nanotube network formation. The method only comprises phospholipids and microfabricated surfaces for simple formation of an ER model and does not require the addition of proteins or chemical energy (e.g., GTP or ATP). In contrast to the 3D morphology of the cellular endoplasmic reticulum, the model is two-dimensional (albeit the nanotube dimensions, geometry, structure, and dynamics are maintained). This unique in vitro ER model consists of only a few components, is easy to construct, and can be observed under a light microscope. The resulting structure can be further decorated for additional functionality, such as the addition of ER-associated proteins or particles to study transport phenomena among the tubes. The artificial networks described here are suitable structural models for the cellular ER, whose unique characteristic morphology has been shown to be related to its biological function, whereas details regarding formation of the tubular domain and rearrangements within are still not completely understood. We note that this method uses Al2O3 thin-film-coated microscopy coverslips, which are commercially available but require special orders. Therefore, it is advisable to have access to a microfabrication facility for preparation.

Solid-supported, protein-free, double phospholipid bilayer membranes (DLBM) can be transformed into complex and dynamic lipid nanotube networks and can serve as 2D bottom-up models of the endoplasmic reticulum.

We present a convenient method to form a bottom-up structural organelle model for the endoplasmic reticulum (ER). The model consists of highly dense ...

Patterning the Geometry of Human Embryonic Stem Cell Colonies on Compliant Substrates to Control Tissue-Level Mechanics

Human embryonic stem cells demonstrate a unique ability to respond to morphogens in vitro by self-organizing patterns of cell fate specification that correspond to primary germ layer formation during embryogenesis. Thus, these cells represent a powerful tool with which to examine the mechanisms that drive early human development. We have developed a method to culture human embryonic stem cells in confined colonies on compliant substrates that provides control over both the geometry of the colonies and their mechanical environment in order to recapitulate the physical parameters that underlie embryogenesis. The key feature of this method is the ability to generate polyacrylamide hydrogels with defined patterns of extracellular matrix ligand at the surface to promote cell attachment. This is achieved by fabricating stencils with the desired geometric patterns, using these stencils to create patterns of extracellular matrix ligand on glass coverslips, and transferring these patterns to polyacrylamide hydrogels during polymerization. This method is also compatible with traction force microscopy, allowing the user to measure and map the distribution of cell-generated forces within the confined colonies. In combination with standard biochemical assays, these measurements can be used to examine the role mechanical cues play in fate specification and morphogenesis during early human development.

Extracellular matrix ligands can be patterned onto polyacrylamide hydrogels to enable the culture of human embryonic stem cells in confined colonies on compliant substrates. This method can be combined with traction force microscopy and biochemical assays to examine the interplay between tissue geometry, cell-generated forces, and fate specification.

Human embryonic stem cells demonstrate a unique ability to respond to morphogens in vitro by self-organizing patterns of cell fate specification that ...