We acknowledge Maya Hendija and Dr. Jonathan Michel for assistance with data analysis, and Dr. Janet Sheung, Dr. Moumita Das, and Dr. Michael Rust for helpful discussions and guidance. This research was supported by a William M. Keck Foundation Research Grant and NSF DMREF Award (DMR 2119663) awarded to RMRA and JLR and National Institutes of Health R15 Grants (R15GM123420, 2R15GM123420-02) awarded to RMR-A and RJM.

1. Prepare silanized coverslips and microscope slides to prevent adsorption of proteins to chamber surfaces
NOTE: This is a 2-day process. Silanized slides may be prepared up to 1 month in advance of use.
<ol>
	<li>Place no. 1 coverslips (24 mm x 24 mm) and microscope slides (1 in x 3 in) in a designated rack that will fit in the plasma cleaner. Place rack in plasma cleaner and run for 20 min.</li>
	<li>Transfer coverslips and slides to a new rack designated only for use with silane and place rack in glass container to clean the glasses as described below.
	<ol>
		<li>Immerse coverslips and slides in 100% acetone for 1 h. Immerse coverslips and slides in 100% ethanol for 10 min.</li>
		<li>Immerse coverslips and slides in deionized water (DI) for 5 min. Repeat the cleaning steps two more times.</li>
		<li>Immerse coverslips and slides in freshly prepared 0.1 M KOH for 15 min. Immerse coverslips and slides in fresh DI for 5 min. Repeat this step two more times.</li>
	</ol>
	</li>
	<li>Air dry coverslips and slides for 10 min. Treat cleaned coverslips and slides with silane to produce hydrophobic surfaces as described below. 
	NOTE: Complete the following steps in a fume hood.
	<ol>
		<li>Immerse&#160;dried coverslips and slides in 2% silane (dissolved in toluene) for 5 min. Use a funnel to pour silane back into its designated bottle to reuse up to five times.</li>
		<li>Immerse coverslips and slides in 100% ethanol for 5 min. Replace ethanol with fresh ethanol. Immerse coverslips and slides for 5 min.</li>
		<li>Immerse coverslips and slides in fresh DI for 5 min. Repeat the ethanol and DI wash step two more times using fresh ethanol and DI each time. Air dry coverslips and slides for 10 min.</li>
	</ol>
	</li>
</ol>
2. Preparing active actin-microtubule composite driven by myosin mini-filaments
<ol>
	<li>Remove inactive myosin via actin filament binding and perform pull-down via ultracentrifugation as described below.
	<ol>
		<li>Polymerize actin into filaments. Using a precision micropipette and sterile pipet tips, combine in a microcentrifuge tube: 1.87 &#181;L of DI, 1.3 &#181;L of 10x G-buffer, 1.3 &#181;L of 10x F-buffer, 1.63 &#181;L of 4 M KCl, 4.53 &#181;L of actin (47.6 &#181;M), and 1.08 &#181;L of 100 &#181;M phalloidin. 
		NOTE: To ensure sufficient polymerization, the actin concentration and actin:phalloidin molar ratio should be 18.4 &#181;M and 2:1, respectively.</li>
		<li>Gently pipet the solution up and down to mix and then set on ice in the dark for &#8805;1 h. Cool ultracentrifuge to 4 &#176;C. Remove myosin aliquot from -80 &#176;C and put on ice. 
		NOTE: Complete step 2.2 at this point while actin is polymerizing.</li>
		<li>After &#8805;1 h of actin polymerization, add 1.3 &#181;L of 10 mM ATP and 2 &#181;L of 19 &#181;M myosin to the polymerized actin. 
		NOTE: The actin:myosin molar ratio should be &#62;5 to ensure sufficient removal of inactive myosin motors (i.e., dead heads).</li>
		<li>Gently pipet the solution up and down to mix. Transfer into an ultracentrifuge grade tube. 
		&#8203;Centrifuge at 4 &#176;C and 121,968 x g for 30 min.</li>
	</ol>
	</li>
	<li>Prepare co-entangled composite network of actin filaments and microtubules as described below. 
	NOTE: Begin 30 min before myosin spin-down (step 2.1.4).
	<ol>
		<li>Set a heat block to 37 &#176;C. Use a precision micropipette and sterile pipet tips to add the following to a microcentrifuge tube: 13.9 &#181;L of PEM, 3 &#181;L of 1% Tween20, 1.55 &#181;L of 47.6 &#181;M actin, 0.36 &#181;L of 34.8 &#181;M R-actin, 0.3 &#181;L of 250 mM ATP, 0.87 &#181;L of 100 &#181;M phalloidin, 1.91 &#181;L of 5-488-tubulin, 0.3 &#181;L of 100 mM GTP, and 0.75 &#181;L of 200 &#181;M Taxol, to a total volume of 23 &#181;L. 
		NOTE: The concentrations of actin and tubulin listed are for a composite with 2.9 &#181;M actin and 2.9 &#181;M tubulin. Total protein concentration is c = cA + cT&#160;= 5.8 &#956;M&#160;and molar actin fraction is cA/(cA + cT) = &#934;A = 0.5. See step 2.5 to adjust these values.</li>
		<li>Gently pipet the solution up and down to mix and place on a 37 &#176;C heat block protected from light for 1 h.</li>
	</ol>
	</li>
	<li>Prepare sample chambers for confocal imaging experiments as described below. 
	NOTE: Complete steps 2.1.4 and 2.2.2 during waiting periods.
	<ol>
		<li>Place two silanized slides side-by-side on a hot plate (turned off), lay two strips of thermoplastic sealing film across the slides ~3 mm apart, and place two silanized coverslips over the thermoplastic sealing film to form a sample chamber.</li>
		<li>Turn the hot plate on low setting until coverslips firmly bind to slides with melted thermoplastic sealing film (~1-2 min). Press down with even pressure to ensure bonding while maintaining ~100 &#181;m spacing between the two surfaces.</li>
		<li>Remove chambers and turn off hot plate. Label chambers with (+) and (-). The (+) chamber will be for the active sample (with myosin) and the (-) chamber will be the control (no myosin). Ensure each chamber can accommodate &#8804;10 &#181;L of fluid.</li>
	</ol>
	</li>
	<li>Prepare samples to image as described below. 
	NOTE: It is important to complete this step immediately after steps 2.1 and 2.2 are completed.
	<ol>
		<li>Carefully remove the myosin-actin sample from the ultracentrifuge (step 2.1.4) and immediately pipet up the top 7.5 &#181;L of the supernatant and transfer to a new microcentrifuge tube.</li>
		<li>Remove the actin-microtubule sample from the heat block and gently mix in 1.5 &#181;L of 10x D-Glucose, 1.5 &#181;L of 10x GOC, and 1.5 &#181;L of 1 mM blebbistatin. Divide the solution into two 13.7 &#181;L aliquots and label as (+) and (-).</li>
		<li>Mix in 1.28 &#181;L of the supernatant from step 2.4.1 to (+) aliquot. Mix in 1.28 &#181;L of DI to the (-) aliquot. Slowly flow each solution into the corresponding chamber (step 2.3) via capillary action. Be careful not to introduce air bubbles into the channel.</li>
		<li>Seal the two open ends of each channel with fast-drying epoxy or UV glue. Ensure the adhesive is completely dry before placing on the microscope. Image immediately as described in step 3. 
		NOTE: UV glue is advantageous&#160;because it cures nearly instantly upon UV exposure. However, because blebbistatin is UV sensitive, it is important to only locally illuminate the glue (at the edges of the sample chamber) using a small UV wand to avoid deactivating the blebbistatin.</li>
	</ol>
	</li>
	<li>Optional: vary the protein concentrations to tune the dynamics and structure of the composites. 
	NOTE: The following steps are suggested alterations to the steps above to vary the concentrations of actin, microtubules, and myosin if desired.
	<ol>
		<li>Follow the steps described above except for the following modifications in steps 2.2.1 and 2.4.3.</li>
		<li>To vary the concentrations of actin and microtubules, thereby adjusting c and &#934;A, increase or decrease the volume of actin, R-actin, and 5-488-tubulin used in step 2.2.1, as desired26. When varying the actin concentration, adjust R-actin and phalloidin molar concentrations proportionally to maintain the same molar ratios with actin. Adjust the volume of PEM such that the final volume of the mixture remains 23 &#181;L. All other component volumes and concentrations remain the same.</li>
		<li>To vary the myosin concentration, adjust the volume of myosin added to the (+) aliquot in step 2.4.3 as desired. Adjust the DI volume added to the (-) aliquot accordingly. Adjust the PEM volume in Step 2.2.1 to account for the increase or decrease in myosin (+) and DI (-) volume, ensuring the final volume of each sample ((+) and (-)) is 14.98&#160;&#956;L.</li>
	</ol>
	</li>
</ol>
3. Imaging and characterization of active composites using confocal microscopy
<ol>
	<li>To image actomyosin-microtubule composites prepared in step 2, use a laser scanning confocal microscope (LSCM), or similar microscope, with a 60x 1.4 NA oil-immersion objective. To simultaneously visualize actin filaments and microtubules in separate fluorescence channels, use a 561 nm laser with 565/591 nm excitation/emission filters and a 488 nm laser with 488/525 nm excitation/emission filters.</li>
	<li>Place the sample chamber on the microscope such that the control channel is positioned directly over the objective. Make sure there is an oil interface between the objective and the coverslip.</li>
	<li>Use the stage controls to bring the control composite into focus, then find both surfaces of the sample chamber. Move the z position to the center of the sample chamber. Check for the presence of clear filamentous networks as shown in Figure 2.</li>
	<li>Still visualizing the control chamber, adjust the intensity of each laser to allow for simultaneous visualization of actin filaments and microtubules. Maintain the lowest laser intensity possible to prevent photobleaching (more prevalent in the actin channel) and bleed through (typically from the microtubules into the actin channel).</li>
	<li>To characterize the inactive control sample, collect three time-series (videos) of 256 x 256 square-pixel (213 &#181;m x 213 &#181;m) images at 2.65 fps for a total of &#8805;1000 frames. Collect each time-series in a different region of the sample chamber separated by &#8805;500 &#181;m. Ensure that there is minimal detectable motion and no flow or restructuring.</li>
	<li>Shutter off the 488 nm laser and use the stage controls to move to the (+) chamber.</li>
	<li>Using the 568 nm laser, visualize the microtubules in the (+) channel to ensure proper network formation (Figure 2) and identify the axial center of the sample chamber (which may be different to the center z-position of the control chamber).</li>
	<li>Turn on the 488 nm laser and repeat step 3.5 above with the following modifications. Collect time-series for up to 45 min, stopping acquisition when the sample either moves out of the field of view, ruptures, or photobleaches. Record 5-10 time-series and keep track of the time at which each time-series begins relative to the start of the first time-series.</li>
	<li>Analyze data using DDM, SIA, and PIV as described in Figure 3, Figure 4, Figure 5, and previously17,48,50,51. 
	&#8203;NOTE: The 488 nm laser locally activates myosin ATPase activity by de-activating blebbistatin, so it should only be turned on at the start of&#160;data acquisition such that t = 0 is at the start of the time-series. These acquisition parameters are optimized for differential dynamic microscopy (DDM) analysis as done previously26.</li>
</ol>
4. Preparation of active actin-microtubule composites driven by kinesin motors
NOTE: The following steps create actin-microtubule composites that are driven out-of-equilibrium by kinesin motors or a combination of kinesin and myosin50.
<ol>
	<li>Prepare kinesin and myosin motors as described below.
	<ol>
		<li>If incorporating myosin, follow step 2.1.</li>
		<li>To form kinesin motor clusters that bind and exert forces between pairs of microtubules, use a micropipette and sterile pipet tips to add the following to a sterile 1.5 mL microcentrifuge tube:1.16 &#181;L PEM, 2.74 &#181;L 8.87 &#181;M kinesin dimers, 7.29 &#181;L 83.3 &#181;M NeutrAvidin, 0.81 &#181;L 2mM DTT . Mix gently by pipetting the solution up and down and incubate protected from light (use a black microcentrifuge tube or wrap in foil) for 30 min at 4 &#176;C. 
		NOTE: The molar ratio of kinesin dimers to NA is 1:25.</li>
	</ol>
	</li>
	<li>Follow step 2.3 to prepare sample chambers and make three chambers instead of two. Carry out this step during kinesin incubation (step 4.1.2) and myosin ultracentrifugation (step 4.1.1).</li>
	<li>Prepare co-entangled composite network of actin filaments and microtubules.
	<ol>
		<li>Set heat block to 37 &#176;C. Use a micropipette and sterile pipet tips to add the following to a sterile 1.5 mL microcentrifuge tube: 3.21 &#181;L PEM, 4.5 &#181;L of 1% Tween20, 2.18 &#181;L of 47.6 &#181;M actin, 3.46 &#181;L of 5-R-tubulin, 4.5&#160;&#181;L of 100 mM ATP, 4.5 &#181;L of 10 mM GTP, 1.13 &#181;L of 200 &#181;M Taxol, and 1.57 &#181;L of 20 &#181;M 488-phalloidin. Ensure the total volume is 25 &#181;L.</li>
		<li>Gently pipet the solution up and down to mix and place on the 37 &#176;C heat block protected from light for 1 h. Remove the tube from the heat block and use a micropipette to gently mix in 0.84 &#181;L of 100&#160;&#956;M phalloidin. Incubate for 5-10 min at room temperature, protected from light. 
		NOTE: Adding phalloidin at this step, rather than in step 4.3.1, improves the fluorescence labeling of actin filaments, as 488-phalloidin does not have to compete with unlabeled phalloidin for actin binding sites.</li>
	</ol>
	</li>
	<li>Prepare active composites for confocal imaging.
	<ol>
		<li>Add 1.13 &#181;L of 200 &#956;M blebbistatin, 1.35 &#181;L of 10x Glu, and 1.35 &#181;L of 10x GOC to the solution from step 4.3.2 and mix gently by pipetting up and down. Divide the solution into three 10 &#181;L aliquots and label as (K), (K+M), and (-).</li>
		<li>Mix in 2.54 &#181;L of myosin from step 2.1.4 to the (K+M) aliquot. Mix in 2.54 &#181;L PEM to (K) and (-) aliquots.</li>
		<li>Use a micropipette and sterile pipet tips to add 2.5 &#181;L of kinesin clusters from step 4.1.2 to (K) and (K+M) aliquots. Pipet up and down to mix. Mix in 2.5 &#181;L PEM to (-) using the same technique. 
		NOTE: The concentrations of actin and tubulin listed are for a composite with 2.32 &#181;M actin and 3.48 &#181;M tubulin. Total protein concentration is c = cA&#160;+ cT&#160;= 5.8&#160;&#956;M&#160;and molar actin fraction is cA/(cA&#160;+ cT) =&#160;&#934;A&#160;= 0.4. Kinesin and myosin concentrations are 0.35 &#181;M and 0.47 &#181;M, respectively. See step 2.5 for general guidelines to adjust cA, cT, c, and &#934;A.</li>
		<li>Using a micropipette, slowly flow each solution into the corresponding channel of the prepared sample chambers (step 4.2) via capillary action. Push down very slowly and gently on the pipet so as to not introduce air bubbles into the channel.</li>
		<li>Seal the two open ends of each channel with fast-drying epoxy or UV-curable glue. Ensure the adhesive is completely dry before placing on the microscope. 
		NOTE: It is important that this step is done quickly to minimize the time that the kinesin is acting without being monitored. For this reason, epoxy that cures in 1 min (rather than 5 or 10 min) is recommended. UV-curable glue is advantageous in this regard because it cures nearly instantly upon UV exposure.&#160;</li>
	</ol>
	</li>
	<li>Image prepared samples immediately, following step 3, except for the following important modifications. Because kinesin is not controlled by light-activation, it starts to work immediately after step 4.4.3, so mark this time as t = 0. To image the composite as close to the initial inactive state (t = 0) as possible, image the (K) and (K+M) channels first and note the time elapsed between step 4.4.3 and the beginning of data acquisition (step 3.8). In practice, this elapsed time is ~5 min.</li>
</ol>
5. Incorporating passive crosslinkers into active composites
NOTE: These steps describe how to use biotinylated actin and tubulin subunits and NeutrAvidin (NA) to passively crosslink actin to actin (A-A) or microtubules to microtubules (M-M) in the active composites described in step 4.
<ol>
	<li>Prepare A-A or M-M crosslinker complexes with biotinylated proteins (biotin-actin or biotin-tubulin), NA, and biotin at a ratio of 2:2:1 biotin-actin/tubulin:biotin:NA. Start this process before Step 4 .
	<ol>
		<li>For A-A crosslinkers, use a micropipette and sterile pipet tips to add 2 &#181;L of 11.6 &#181;M biotin-actin, 1.39 &#181;L of 8.33 &#181;M NA, 2.27 &#181;L of 1.02 &#181;M biotin, and 4.34 &#181;L of PEM to a microcentrifuge tube. Mix gently by pipetting up and down.</li>
		<li>For M-M crosslinkers, use a micropipette and sterile pipet tips to add 1.86 &#181;L of 4.55 &#181;M biotin-tubulin, 1.11 &#181;L of 8.33 &#181;M NA, 1.82 &#181;L of 1.02 &#181;M biotin, and 5.21 &#181;L of PEM to a microcentrifuge tube. Mix gently by pipetting up and down.</li>
		<li>Wrap the tube(s) from step 5.1.1 and/or 5.1.2 in thermoplastic sealing film to create a water-tight seal. Place in a flotation raft in a temperature-controlled sonicator bath set to 4 &#176;C.</li>
		<li>Sonicate for 90 min at 4 &#176;C. In practice it is best to put the sonicator in a cold room and add ice packs to the sonication bath to maintain low temperature.</li>
	</ol>
	</li>
	<li>To incorporate crosslinker complexes into samples for imaging, follow step 4.3, modifying step 4.3.1 as described below for A-A crosslinking (step 5.2.1) or M-M crosslinking (step 5.2.2).
	<ol>
		<li>For A-A crosslinking, combine the following in a microcentrifuge tube: 1.94&#160;&#181;L of PEM, 4.50 &#181;L of 1% Tween20, 2.18 &#181;L of 47.6 &#181;M actin, 3.46 &#181;L of 45.5 &#181;M 5-R-tubulin, 1.13 &#181;L of A-A crosslinkers (step 5.1.1), 4.50 &#181;L of 100&#160;mM ATP, 4.50 &#181;L of 10 mM GTP, 1.13 &#181;L of 200 &#181;M Taxol, and 1.57 &#181;L of 20 &#181;M 488-phalloidin. Ensure the total volume is 25 &#181;L.</li>
		<li>For M-M crosslinking, combine the following in a microcentrifuge tube: 1.97&#160;&#181;L of PEM, 4.50 &#181;L of 1% Tween20, 2.18 &#181;L of 47.6 &#181;M actin, 3.76 &#181;L of 45.5 &#181;M 5-R-tubulin, 1.13 &#181;L of 1:4 dilution of M-M crosslinkers (step 5.1.2), 4.50 &#181;L of 100 mM ATP, 4.50 &#181;L of 10 mM GTP, 1.13 &#181;L of 200 &#181;M Taxol, and 1.57 &#181;L of 20 &#181;M 488-phalloidin. Ensure the total volume is 25 &#181;L.</li>
	</ol>
	</li>
	<li>Follow steps 4.3.2-4.5 with the specific concentrations for a crosslinker:actin molar ratio of RA = 0.02 and crosslinker:tubulin molar ratio of RT =&#160;0.005. These RA and RT values result in similar lengths between crosslinkers along actin filaments and microtubules (dA <img alt="Equation 1" src="/files/ftp_upload/64228/64228eq1.png" />&#160;60 nm and dMT&#160;<img alt="Equation 1" src="/files/ftp_upload/64228/64228eq1.png" /> 67 nm), estimated using dA = Imonomer/2RA, where Imonomer is the length of an actin monomer, and dMT = Iring/26RT, where Iring is the length of a ring of 13 tubulins15,17.</li>
</ol>

<ol>
	<li>Fletcher, D. A., Mullins, R. D. Cell mechanics and the cytoskeleton. Nature. 463 (7280), 485-492 (2010).</li>
	<li>Koenderink, G. H., Paluch, E. K. Architecture shapes contractility in actomyosin networks. Current Opinion in Cell Biology. 50, 79-85 (2018).</li>
	<li>Dogterom, M., Koenderink, G. H. Actin-microtubule crosstalk in cell biology. Nature Reviews Molecular Cell Biology. 20 (1), 38-54 (2019).</li>
	<li>Burla, F., Mulla, Y., Vos, B. E., Aufderhorst-Roberts, A., Koenderink, G. H. From mechanical resilience to active material properties in biopolymer networks. Nature Reviews Physics. 1 (4), 249-263 (2019).</li>
	<li>Wen, Q., Janmey, P. A. Polymer physics of the cytoskeleton. Current Opinion in Solid State and Materials Science. 15 (5), 177-182 (2011).</li>
	<li>Xiao, Q., Hu, X., Wei, Z., Tam, K. Y. Cytoskeleton molecular motors: structures and their functions in neuron. International Journal of Biological Sciences. 12 (9), 1083-1092 (2016).</li>
	<li>Ajeti, V. et al. Wound healing coordinates actin architectures to regulate mechanical work. Nature Physics. 15 (7), 696-705 (2019).</li>
	<li>Jung, W. et al. Dynamic motions of molecular motors in the actin cytoskeleton. Cytoskeleton. 76 (11-12), 517-531 (2019).</li>
	<li>Pollard, T. D., O&#39;Shaughnessy, B. Molecular mechanism of cytokinesis. Annual Review of Biochemistry. 88 (1), 661-689 (2019).</li>
	<li>Huber, F., Boire, A., L&#243;pez, M. P., Koenderink, G. H. Cytoskeletal crosstalk: when three different personalities team up. Current Opinion in Cell Biology. 32, 39-47 (2015).</li>
	<li>Rivero, F. et al. The role of the cortical cytoskeleton: F-actin crosslinking proteins protect against osmotic stress, ensure cell size, cell shape and motility, and contribute to phagocytosis and development. Journal of Cell Science. 109 (11), 2679-2691 (1996).</li>
	<li>Duclos, G. et al. Topological structure and dynamics of three-dimensional active nematics. Science. 367 (6482), 1120-1124 (2020).</li>
	<li>Baclayon, M. et al. Optical tweezers-based measurements of forces and dynamics at microtubule ends. Optical Tweezers. 1486, 411-435 (2017).</li>
	<li>Gurmessa, B., Fitzpatrick, R., Falzone, T. T., Robertson-Anderson, R. M. Entanglement density tunes microscale nonlinear response of entangled actin. Macromolecules. 49 (10), 3948-3955 (2016).</li>
	<li>Francis, M. L. et al. Non-monotonic dependence of stiffness on actin crosslinking in cytoskeleton composites. Soft Matter. 15 (44), 9056-9065 (2019).</li>
	<li>Ricketts, S. N. et al. Varying crosslinking motifs drive the mesoscale mechanics of actin-microtubule composites. Scientific Reports. 9 (1), 12831 (2019).</li>
	<li>Lee, G. et al. Active cytoskeletal composites display emergent tunable contractility and restructuring. Soft Matter. 17 (47), 10765-10776 (2021).</li>
	<li>Murrell, M. P., Gardel, M. L. F-actin buckling coordinates contractility and severing in a biomimetic actomyosin cortex. Proceedings of the National Academy of Sciences. 109 (51), 20820-20825 (2012).</li>
	<li>Soares e Silva, M. et al. Active multistage coarsening of actin networks driven by myosin motors. Proceedings of the National Academy of Sciences. 108 (23), 9408-9413 (2011).</li>
	<li>Sonn-Segev, A., Bernheim-Groswasser, A., Roichman, Y. Dynamics in steady state in vitro acto-myosin networks. Journal of Physics: Condensed Matter. 29 (16), 163002 (2017).</li>
	<li>Ideses, Y., Sonn-Segev, A., Roichman, Y., Bernheim-Groswasser, A. Myosin II does it all: assembly, remodeling, and disassembly of actin networks are governed by myosin II activity. Soft Matter. 9 (29), 7127 (2013).</li>
	<li>F&#252;rthauer, S. et al. Self-straining of actively crosslinked microtubule networks. Nature Physics. 15 (12), 1295-1300 (2019).</li>
	<li>Lemma, L. M. et al. Multiscale microtubule dynamics in active nematics. Physical Review Letters. 127 (14), 148001 (2021).</li>
	<li>Fan, Y., Wu, K.-T., Aghvami, S. A., Fraden, S., Breuer, K. S. Effects of confinement on the dynamics and correlation scales in kinesin-microtubule active fluids. Physical Review E. 104 (3), 034601 (2021).</li>
	<li>Triclin, S. et al. Self-repair protects microtubules from destruction by molecular motors. Nature Materials. 20 (6), 883-891 (2021).</li>
	<li>Lee, G. et al. Myosin-driven actin-microtubule networks exhibit self-organized contractile dynamics. Science Advances. 7 (6), eabe4334 (2021).</li>
	<li>Ricketts, S. N., Ross, J. L., Robertson-Anderson, R. M. Co-entangled actin-microtubule composites exhibit tunable stiffness and power-law stress relaxation. Biophysical Journal. 115 (6), 1055-1067 (2018).</li>
	<li>Bendix, P. M. et al. A quantitative analysis of contractility in active cytoskeletal protein networks. Biophysical Journal. 94 (8), 3126-3136 (2008).</li>
	<li>Linsmeier, I. et al. Disordered actomyosin networks are sufficient to produce cooperative and telescopic contractility. Nature Communications. 7 (1), 12615 (2016).</li>
	<li>Stam, S. et al. Filament rigidity and connectivity tune the deformation modes of active biopolymer networks. Proceedings of the National Academy of Sciences. 114 (47), E10037-E10045 (2017).</li>
	<li>Yadav, V. et al. Filament nucleation tunes mechanical memory in active polymer networks. Advanced Functional Materials. 29 (49), 1905243 (2019).</li>
	<li>Ennomani, H. et al. Architecture and connectivity govern actin network contractility. Current Biology. 26 (5), 616-626 (2016).</li>
	<li>Alvarado, J., Sheinman, M., Sharma, A., MacKintosh, F. C., Koenderink, G. H. Molecular motors robustly drive active gels to a critically connected state. Nature Physics. 9 (9), 591-597 (2013).</li>
	<li>Alvarado, J., Cipelletti, L., Koenderink, G. H. Uncovering the dynamic precursors to motor-driven contraction of active gels. Soft Matter. 15 (42), 8552-8565 (2019).</li>
	<li>Jung, W., Murrell, M. P., Kim, T. F-actin cross-linking enhances the stability of force generation in disordered actomyosin networks. Computational Particle Mechanics. 2 (4), 317-327 (2015).</li>
	<li>Lenz, M., Thoresen, T., Gardel, M. L., Dinner, A. R. Contractile units in disordered actomyosin bundles arise from f-actin buckling. Physical Review Letters. 108 (23), 238107 (2012).</li>
	<li>Memarian, F.L. et al. Active nematic order and dynamic lane formation of microtubules driven by membrane-bound diffusing motors. Proceedings of the National Academy of Sciences. 118 (52), e2117107118 (2021).</li>
	<li>Needleman, D., Dogic, Z. Active matter at the interface between materials science and cell biology. Nature Reviews Materials. 2 (9), 17048 (2017).</li>
	<li>Foster, P. J., F&#252;rthauer, S., Shelley, M. J., Needleman, D. J. Active contraction of microtubule networks. eLife. 4, e10837 (2015).</li>
	<li>Thijssen, K. et al. Submersed micropatterned structures control active nematic flow, topology, and concentration. Proceedings of the National Academy of Sciences. 118 (38), e2106038118 (2021).</li>
	<li>Sanchez, T., Chen, D. T. N., DeCamp, S. J., Heymann, M., Dogic, Z. Spontaneous motion in hierarchically assembled active matter. Nature. 491 (7424), 431-434 (2012).</li>
	<li>Colen, J. et al. Machine learning active-nematic hydrodynamics. Proceedings of the National Academy of Sciences. 118 (10), e2016708118 (2021).</li>
	<li>Mitchell, K. A., Tan, A. J., Arteaga, J., Hirst, L. S. Fractal generation in a two-dimensional active-nematic fluid. Chaos: An Interdisciplinary Journal of Nonlinear Science. 31 (7), 073125 (2021).</li>
	<li>Pandolfi, R. J., Edwards, L., Johnston, D., Becich, P., Hirst, L. S. Designing highly tunable semiflexible filament networks. Physical Review E. 89 (6), 062602 (2014).</li>
	<li>Tan, A. J. et al. Topological chaos in active nematics. Nature Physics. 15 (10), 1033-1039 (2019).</li>
	<li>Roostalu, J., Rickman, J., Thomas, C., N&#233;d&#233;lec, F., Surrey, T. Determinants of polar versus nematic organization in networks of dynamic microtubules and mitotic motors. Cell. 175 (3), 796-808.e14 (2018).</li>
	<li>Ndlec, F. J., Surrey, T., Maggs, A. C., Leibler, S. Self-organization of microtubules and motors. Nature. 389 (6648), 305-308 (1997).</li>
	<li>Sheung, J. Y. et al. Motor-driven restructuring of cytoskeleton composites leads to tunable time-varying elasticity. ACS Macro Letters. 10 (9), 1151-1158 (2021).</li>
	<li>McGorty, R. PyDDM v0.2.0. Zenodo. (2022).</li>
	<li>Achiriloaie, D. H. et al. Kinesin and myosin motors compete to drive rich multi-phase dynamics in programmable cytoskeletal composites. arXiv. (2021).</li>
	<li>Wulstein, D. M., Regan, K. E., Garamella, J., McGorty, R. J., Robertson-Anderson, R. M. Topology-dependent anomalous dynamics of ring and linear DNA are sensitive to cytoskeleton crosslinking. Science Advances. 5 (12), eaay5912 (2019).</li>
	<li>McGorty, R. Image-Correlation. at &#60;https://github.com/rmcgorty/Image-Correlation&#62;. (2020).</li>
	<li>Robertson, C. Theory and practical recommendations for autocorrelation-based image correlation spectroscopy. Journal of Biomedical Optics. 17 (8), 080801 (2012).</li>
	<li>McGorty, R. Differential Dynamic Microscopy - Python. at &#60;https://github.com/rmcgorty/Differential-Dynamic-Microscopy---Python&#62;. (2021).</li>
	<li>Cerbino, R., Trappe, V. Differential dynamic microscopy: probing wave vector dependent dynamics with a microscope. Physical Review Letters. 100 (18), 188102 (2008).</li>
	<li>Robertson-Anderson, R. M. Optical tweezers microrheology: from the basics to advanced techniques and applications. ACS Macro Letters. 7 (8), 968-975 (2018).</li>
	<li>Pollard, T. D. Polymerization of ADP-actin. Journal of Cell Biology. 99 (3), 769-777 (1984).</li>
	<li>Cou&#233;, M., Brenner, S. L., Spector, I., Korn, E. D. Inhibition of actin polymerization by latrunculin A. FEBS Letters. 213 (2), 316-318 (1987).</li>
	<li>Pollard, T. D. Actin and actin-binding proteins. Cold Spring Harbor Perspectives in Biology. 8 (8), a018226 (2016).</li>
	<li>Kumar, N. Taxol-induced polymerization of purified tubulin. Mechanism of action. Journal of Biological Chemistry. 256 (20), 10435-10441 (1981).</li>
	<li>K&#228;s, J. et al. F-actin, a model polymer for semiflexible chains in dilute, semidilute, and liquid crystalline solutions. Biophysical Journal. 70 (2), 609-625 (1996).</li>
	<li>Viamontes, J., Narayanan, S., Sandy, A. R., Tang, J. X. Orientational order parameter of the nematic liquid crystalline phase of F -actin. Physical Review E. 73 (6), 061901 (2006).</li>
	<li>Hitt, A. L., Cross, A. R., Williams, R. C. Microtubule solutions display nematic liquid crystalline structure. Journal of Biological Chemistry. 265 (3), 1639-1647 (1990).</li>
	<li>Andexer, J. N., Richter, M. Emerging enzymes for ATP regeneration in biocatalytic processes. ChemBioChem. 16 (3), 380-386 (2015).</li>
	<li>Farhadi, L. et al. Actin and microtubule crosslinkers tune mobility and control co-localization in a composite cytoskeletal network. Soft Matter. 16 (31), 7191-7201 (2020).</li>
	<li>Falzone, T. T., Lenz, M., Kovar, D. R., Gardel, M. L. Assembly kinetics determine the architecture of &#945;-actinin crosslinked F-actin networks. Nature Communications. 3 (1), 861 (2012).</li>
	<li>Thoresen, T., Lenz, M., Gardel, M. L. Reconstitution of contractile actomyosin bundles. Biophysical Journal. 100 (11), 2698-2705 (2011).</li>
	<li>Sanghvi-Shah, R., Weber, G. F. Intermediate filaments at the junction of mechanotransduction, migration, and development. Frontiers in Cell and Developmental Biology. 5, 81 (2017).</li>
	<li>Shen, Y. et al. Effects of vimentin intermediate filaments on the structure and dynamics of in vitro multicomponent interpenetrating cytoskeletal networks. Physical Review Letters. 127 (10), 108101 (2021).</li>
	<li>Laan, L., Roth, S., Dogterom, M. End-on microtubule-dynein interactions and pulling-based positioning of microtubule organizing centers. Cell Cycle. 11 (20), 3750-3757 (2012).</li>
</ol>

The authors have nothing to disclose

A key advance of the reconstituted system described above is its modularity and tunability, so users are encouraged to modify the concentrations of proteins, motors, crosslinkers, etc. to suit their desired outcomes, whether it be to emulate a particular cellular process or engineer a material with specific functionality or mechanical properties. Limitations on the concentration range of actin and tubulin are set at the lower limit by the critical concentration needed to polymerize actin (~0.2 &#956;M)57,58,59 and tubulin (~3 - 4&#160;&#956;M)60, and at the upper limit by the transition to nematic alignment of actin filaments (~90 &#956;M)61,62 or microtubules (~35&#160;&#956;M)63. Actin monomers and tubulin dimers should be polymerized into filaments together, rather than mixed together after polymerization, to ensure that they form homogenously interpenetrating percolated networks that synergistically support each other. The novel dynamics that the composites exhibit rely on this interaction. While it is generally important to follow all steps as outlined in the protocol to successfully reproduce the results shown, some steps are more exacting, while others have room to modify and adjust to fit specific needs and available resources.
For example, one important step in ensuring reproducible results is properly preparing and storing the reagents following the guidelines provided in the Table of Materials. Cytoskeletal proteins (actin, tubulin, myosin, kinesin) are labile and should be aliquoted, flash-frozen with liquid nitrogen, and stored at -80 &#176;C in single-use aliquots. Once removed from -80 &#176;C, aliquots should be kept on ice. Cytoskeletal proteins do not reliably retain function after additional freeze-thaw cycles.
Microtubules are more sensitive to depolymerization and denaturing than actin. Once removed from -80 &#176;C, tubulin should be kept on ice before polymerization, and used within 12 h. Once polymerized, microtubules should be kept at room temperature. It is also critical to stabilize microtubules with taxol to prevent depolymerization. Phalloidin-stabilization of actin filaments is likewise important to suppress the ATP-consuming actin treadmilling that competes with myosin and kinesin activity.
Ultracentrifugation of myosin motors is another critical step, as it removes inactive myosin dead heads. Not removing the enzymatically inactive monomers results in passive crosslinking of the actin network and loss of activity. To prolong ATPase activity of motors, an ATP regeneration system such as creatine phosphate and creatine phosphokinase64 can be incorporated.
Finally, maintaining composite activity requires inhibiting adsorption of filaments and motors to the walls of the sample chamber, which can be achieved by passivation of the microscope coverslips and slides. Motor proteins are particularly prone to adsorption, which results in the composite being pulled to the surface of the sample chamber, moving out of the field-of-view, collapsing to 2D, and no longer undergoing activity. Silanizing the coverslips and slides is an effective way to passivate the surfaces and prevent adsorption (see step 1). An alternative passivation method used effectively in in vitro cytoskeleton experiments is coating the surface with a lipid bilayer, similar to the cell membrane18. This method is advantageous if one wishes to tether proteins to the surface or introduce other specific protein-surface interactions, because the bilayer can be functionalized. For optical tweezers experiments, passivation of the microspheres is also critical, and can be achieved by coating carboxylated microspheres with BSA or PEG via carbodiimide crosslinker chemistry48.
There are a few aspects of the presented protocols that researchers may consider altering to suit their needs. Firstly, researchers may choose to replace non-native biotin-NA crosslinkers with biological crosslinkers, such as alpha-actinin or MAP65 that crosslink actin and microtubules, respectively28,65,66. The use of non-native crosslinkers in the composites described here is motivated by their enhanced reproducibility, stability, and tunability compared to native crosslinkers. Because of the strong biotin-NA bond, crosslinkers can be assumed to be permanent, rather than most native crosslinkers that transiently bind with wide-ranging turnover rates. The dynamics of transient crosslinking complicates parsing the contributions from crosslinkers and motors to the dynamics. Moreover, biotin-NA linkers can be versatilely used to crosslink both actin and microtubules, as well as crosslink actin to microtubules. In this way, an unambiguous comparison between crosslinking motifs can be made, keeping all other variables (e.g., crosslinker size, binding affinity, stoichiometry, etc.) fixed. Finally, the reagents needed to incorporate biotin-NA linkers are widely commercially available, well-characterized, and commonly used in many biophysics labs. However, one of the key strengths of the in vitro platform described here is its modularity, so researchers should be able to seamlessly replace biotin-NA linkers with native linkers should they choose.
Secondly, in the current protocol, actin monomers and tubulin dimers are polymerized into filaments together in a centrifuge tube prior to adding to the sample chamber. Flowing the solution of entangled filamentous proteins into the sample chamber may cause flow alignment, especially of the microtubules, which breaks the desired isotropy and homogeneity of the composites. Indeed, a major advance in prior work on steady-state actin-microtubule composites was the ability to co-polymerize actin and microtubules in situ (in the sample chamber) to ensure formation of isotropic interpenetrating networks of actin and microtubules15,16,27. However, extending this approach to active composites would require adding the motors to the sample prior to actin and tubulin polymerization and having the entire sample incubate together at 37 &#176;C prior to experiments. Tests of this variation to the protocol have resulted in reduced actin polymerization and no discernible motor activity, likely due to competing ATPase activity and the prolonged 37&#160;&#176;C incubation of the motors. Fortunately, there is no discernible flow alignment of composites when following the current protocols, as can be seen in Figure 2, Figure 3, and Figure 6. Nevertheless, researchers are encouraged to design protocols that allow for in situ formation of active composites.
Another point of consideration is the fluorescence labeling scheme, which entails sparsely labeling all of the actin filaments and microtubules in the network. This labeling approach was optimized to directly visualize the structure of the network rather than inferring structure and dynamics via tracer filaments or microspheres. However, the tradeoff is that individual filaments are not brightly labeled and resolvable. One approach that researchers could take to both resolve single filaments as well as visualize network structure is to dope in pre-formed filaments labeled with another fluorophore, so both the surrounding network and individual filaments could be imaged simultaneously. However, when using more than two fluorophores and excitation/emission channels, bleed-through between channels is often hard to eliminate, so care must be taken in choosing the fluorophores, filters, and laser intensities.
A related limitation is the inability to visualize the myosin or kinesin motors in the composites. The fluorescent-labeled actin monomers and tubulin dimers used are commercially available, whereas visualization of myosin or kinesin in composites requires in-house labeling. Researchers are encouraged to take the next step to label motors, as done previously18,67, to be able to unequivocally link motor activity and binding to the dynamics and structures that our composites exhibit.
Finally, it is important to note that, in the current protocol, the onset and duration of kinesin activity is not controlled. Because the myosin activity is controlled using photo-deactivation of blebbistatin, as described above, to build in similar light-activation of kinesin, one can incorporate light-activated ATP.
To build up the complexity of the designs described here, to better mimic cellular conditions and broaden the dynamic-structure-function parameter space, future work will focus on incorporating intermediate filaments, such as vimentin68,69, as well as other motors such as dynein13,70. Gelsolin will also be incorporated at different concentrations to control actin length14, as well as tau protein to control microtubule stiffness.
In summary, the presented protocols describe how to design, create, and characterize the dynamics, structure, and mechanics of cytoskeleton-inspired active matter systems, that contain two separate active force-generating components that act on different substrates in a single system. This tunable and modular platform brings reconstitution efforts one important step closer to mimicking the cellular cytoskeleton and offers the unique ability to program its properties across a wide phase space by independently incorporating, removing, and tuning the different components. Moreover, all components of this versatile system are commercially available (see Table of Materials), except for the kinesin dimers which are purified in the Ross Lab, as described previously50, and available upon request. Finally, all analysis code is freely available through GitHub49 and is based on free programming languages and software (Python and Fiji). The transparent dissemination of protocols to design these systems will hopefully make this platform more accessible to a diverse group of users with different expertise, backgrounds, institutional affiliations, and research goals.

An erratum was issued for: <a href="https://www.jove.com/t/64228">Reconstituting and Characterizing Actin-Microtubule Composites with Tunable Motor-Driven Dynamics and Mechanics</a>. The Authors section was updated.
Mehrzad Sasanpour1 
Daisy H. Achiriloaie1,2 
Gloria Lee1 
Gregor Leech1 
Christopher Currie1 
K. Alice Lindsay3 
Jennifer L. Ross3 
Ryan J. McGorty1 
Rae M. Robertson-Anderson1 
1Department of Physics and Biophysics, University of San Diego 
2W. M. Keck Science Department, Scripps College, Pitzer College, and Claremont McKenna College 
3Department of Physics, Syracuse University
to:
Mehrzad Sasanpour1 
Daisy H. Achiriloaie1,2 
Gloria Lee1 
Gregor Leech1 
Maya Hendija1 
K. Alice Lindsay3 
Jennifer L. Ross3 
Ryan J. McGorty1 
Rae M. Robertson-Anderson1 
1Department of Physics and Biophysics, University of San Diego 
2W. M. Keck Science Department, Scripps College, Pitzer College, and Claremont McKenna College 
3Department of Physics, Syracuse University

An erratum was issued for:&#160;<a href="https://www.jove.com/t/64228">Reconstituting and Characterizing Actin-Microtubule Composites with Tunable Motor-Driven Dynamics and Mechanics</a>. The Authors section was updated.

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

The cytoskeleton is a dynamic composite network of interacting biopolymers that provides structural and mechanical support to cells. Associated molecular motors and binding proteins restructure and adapt the cytoskeleton to allow cells to grow, change shape, stiffen, move, and even self-heal, enabling myriad cellular processes ranging from migration and division to mechanosensing1,2. Beyond its significance in cellular biophysics, the cytoskeleton is also a quintessential example of active matter with potential materials applications ranging from wound healing and drug delivery to filtration and soft robotics1,3,4,5,6,7,8,9.
The two key characteristics that endow the cytoskeleton with its unique structural and mechanical diversity and multifunctionality are: 1) its composite nature, comprising multiple interacting protein filaments, such as semiflexible actin filaments and rigid microtubules, as well as their associated binding and crosslinking proteins3,5,10; and 2) its ability to continuously restructure, move, coarsen, and perform work via energy-consuming motors, such as myosins and kinesins, pushing and pulling on the filamentous proteins1,7,11,12,13. While this elegant complexity enables the cytoskeleton to mediate processes as diverse as cell motility, cytokinesis, and wound-healing3,6,7,11, it hampers the ability of researchers to reproduce the signature in vivo characteristics of the cytoskeleton in reconstituted in vitro systems.
Current frontier reconstitution efforts focus on composites of entangled and crosslinked actin filaments and microtubules3,10,14,15,16,17, force-generating actomyosin networks2,8,18,19,20,21, and active nematics driven by kinesin-microtubule interactions22,23,24,25,26. Steady-state actin-microtubule composites have been shown to display emergent mechanical properties15,16,27, such as enhanced filament mobility and increased stiffness compared to single-component systems27. Studies on in vitro actomyosin systems have reported a wide range of structural and dynamical properties that depend on the concentrations of actin, myosin, and crosslinkers28,29,30,31. For example, with sufficient crosslinking, actomyosin networks undergo large-scale contraction and coarsening2,28,30,32,33,34,35,36, whereas without crosslinkers, networks display rapid, destabilizing flow and rupturing19,29. Reconstituted microtubule-based active nematics that use clusters of kinesin motors to crosslink and pull on microtubule bundles have been reported to exhibit long lasting turbulent flows, extension, buckling, fracturing, and healing12,22,23,24,25,37,38,39,40,41,42,43,44,45,46,47.
More recently, actin-microtubule composites driven by myosin II mini-filaments have been shown to lead to more ordered contraction and network integrity compared to the disordered flow and network rupturing that actomyosin networks without crosslinkers exhibit17,26,48. Moreover, the combination of composite robustness and force-generation is optimized when actin and microtubules are present at comparable concentrations. Key emergent features in this region of formulation space include enhanced mechanical strength26, coordinated motion of actin and microtubules26, steady sustained contraction, and mesoscale restructuring17.
Here, protocols are described to engineer and tune co-entangled and crosslinked composites of microtubules and actin filaments that are pushed out of equilibrium by myosin II mini-filaments and kinesin clusters acting on actin filaments and microtubules, respectively (Figure 1). The dynamics, structure, and mechanics of this class of composites can be tuned by the relative concentrations of the filaments, motors, and crosslinkers to exhibit a rich phase space of advective and turbulent flow, isotropic contraction, acceleration, deceleration, de-mixing, stiffening, relaxation, and rupturing. The focus of this work is on preparing and tuning this class of active cytoskeletal composites. However, to aid researchers in benchmarking and characterizing the described active composites, effective imaging methods using multi-spectral confocal microscopy are also detailed. Finally, results of key computational analysis methods that can be used to measure the dynamics, structure, and mechanics of the composites are presented. Researchers are encouraged to adopt these methods-which include differential dynamic microscopy (DDM), spatial image autocorrelation (SIA), and particle image velocimetry (PIV)-as they have been optimized to characterize the complex dynamics and structural diversity of the composites17,26,49.
The steps described below focus on preparing the composites and imaging them using confocal microscopy. Protocols describing post-acquisition data analysis and optical tweezers measurements can be found in previous works&#160;17,26,48,50,&#160;and provided upon request. All materials are listed in the Table of Materials provided.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>(-)-Blebbistatin 
			&#160;Abbreviation used in paper: blebbistatin</td>
			<td>Sigma Aldrich</td>
			<td>B0560</td>
			<td>Stock Concentration: 200 &#956;M in DMSO 
			Storage: dessicated, in DMSO, -20&#186;C 
			Stock and Experiment Recipes: dissolve 1 mg of powder to 200 &#956;M in DMSO 
			Storage, Handling, Troubleshooting Notes: limited shelf-life, typically stops functioning reliably after 3-4 months. purchase and prepare new solution every 3 months.&#160;</td>
		</tr>
		<tr>
			<td>1:20 488-tubulin:tubulin mixture 
			&#160;Abbreviation used in paper: 5-488-tubulin</td>
			<td>NA</td>
			<td>NA</td>
			<td>Stock Concentration: 5 mg/ml in PEM 
			Storage: single use aliquots, -80&#186;C, avoid freeze-thaw cycles 
			Stock and Experiment Recipes: mix tubulin and 488-tubulin at a 20:1 ratio, flash freeze with LN2 
			Storage, Handling, Troubleshooting Notes: each aliquot can be used for up to 12 hrs stored on ice at 4&#186;C, protect from light</td>
		</tr>
		<tr>
			<td>1:20 R-tubulin:tubulin mixture 
			&#160;Abbreviation used in paper: 5-R-tubulin</td>
			<td>NA</td>
			<td>NA</td>
			<td>Stock Concentration: 5 mg/ml in PEM 
			Storage: single use aliquots, -80&#186;C, avoid freeze-thaw cycles 
			Stock and Experiment Recipes: mix tubulin and rhodamine tubulin at a 20:1 ratio, flash freeze with LN2 
			Storage, Handling, Troubleshooting Notes: each aliquot can be used for up to 12 hrs stored on ice at 4&#186;C, protect from light</td>
		</tr>
		<tr>
			<td>actin (biotin): skeletal muscle 
			&#160;Abbreviation used in paper: biotin-actin</td>
			<td>Cytoskeleton</td>
			<td>AB07</td>
			<td>Stock Concentration: 1 mg/ml in G-buffer 
			Storage: single use aliquots, -80&#186;C, avoid freeze-thaw cycles 
			Stock and Experiment Recipes: reconstitute to 1 mg/ml in G-buffer, flash freeze with LN2 
			Storage, Handling, Troubleshooting Notes: (1) immediately prior to use dilute to 0.5 mg/ml in PEM, (2) once removed from -80&#186;C, store&#160; aliquot on ice at 4&#186;C for up to 1 week</td>
		</tr>
		<tr>
			<td>actin (rhodamine): rabbit skeletal muscle 
			&#160;Abbreviation used in paper: R-actin</td>
			<td>Cytoskeleton</td>
			<td>AR05</td>
			<td>Stock Concentration: 1.5 mg/ml in G-buffer 
			Storage: single use aliquots, -80&#186;C, avoid freeze-thaw cycles 
			Stock and Experiment Recipes: reconstitute to 1.5 mg/ml in G-buffer, flash freeze with LN2 
			Storage, Handling, Troubleshooting Notes: once removed from -80&#186;C, store&#160; aliquot on ice at 4&#186;C, can be used for up to 1 week</td>
		</tr>
		<tr>
			<td>adenosine triphosphate 
			&#160;Abbreviation used in paper: ATP</td>
			<td>Thermo Fisher Scientific</td>
			<td>A1048</td>
			<td>Stock Concentration: 100 mM 
			Storage: in solution (pH 7), -20&#186;C 
			Stock and Experiment Recipes: reconsitute in DI H20, bring pH to 7 with NaOH 
			Storage, Handling, Troubleshooting Notes: routinely check pH and adjust as needed, hydrolyzes over time, replace every ~6-12 months</td>
		</tr>
		<tr>
			<td>AlexaFluor488 Phalloidin 
			&#160;Abbreviation used in paper: 488-phalloidin</td>
			<td>Thermo Fisher Scientific</td>
			<td>A12379</td>
			<td>Stock Concentration: 100 &#956;M DMSO 
			Storage: protected from light, dessicated, -20&#186;C 
			Stock and Experiment Recipes: reconstitute to 100 &#956;M with DMSO 
			Storage, Handling, Troubleshooting Notes: immediately prior to use dilute to 20 &#956;M in PEM (1 &#956;L in 4 &#956;L PEM)</td>
		</tr>
		<tr>
			<td>AlexaFluor488&#8211;labeled actin 
			&#160;Abbreviation used in paper: 488-actin</td>
			<td>Thermo Fisher Scientific</td>
			<td>A12373</td>
			<td>Stock Concentration: 1.5 mg/ml in G-buffer 
			Storage: single use aliquots, -80&#186;C, avoid freeze-thaw cycles 
			Stock and Experiment Recipes: reconstitute to 1.5 mg/ml in G-buffer, flash freeze with LN2 
			Storage, Handling, Troubleshooting Notes: this item has been discontinued</td>
		</tr>
		<tr>
			<td>Basic Plasma Cleaner 
			&#160;Abbreviation used in paper: plasma cleaner</td>
			<td>Harrick Plasma</td>
			<td>PDC-32G</td>
			<td></td>
		</tr>
		<tr>
			<td>Bemis&#160;Parafilm M Laboratory Wrapping Film 
			&#160;Abbreviation used in paper: transparent film</td>
			<td>Thermo Fisher Scientific</td>
			<td>13-374-5</td>
			<td></td>
		</tr>
		<tr>
			<td>D-(+)-Glucose 
			&#160;Abbreviation used in paper:&#160;</td>
			<td>Thermo Fisher Scientific</td>
			<td>A1682836</td>
			<td>Stock Concentration: 100x 
			Storage: store at stock concentration (100x) or 10x concentration, dessicated, at -20&#186;C 
			Stock and Experiment Recipes: reconstitute powder to 4.5 mg/ml in DI H20 
			Storage, Handling, Troubleshooting Notes: final concentration in solution should 45 &#956;g/mL&#160;</td>
		</tr>
		<tr>
			<td>D-Biotin 
			&#160;Abbreviation used in paper: biotin</td>
			<td>Fisher Scientific</td>
			<td>BP232-1</td>
			<td>Stock Concentration: 1.02 mM in PEM 
			Storage: dessicated, 4&#186;C</td>
		</tr>
		<tr>
			<td>deionized nanopure water 
			&#160;Abbreviation used in paper: DI</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Dimethyldichlorosilane 
			&#160;Abbreviation used in paper: silane</td>
			<td>Thermo Fisher Scientific</td>
			<td>D/3820/PB05</td>
			<td>Stock Concentration: 2% dissolved in Toulene</td>
		</tr>
		<tr>
			<td>Dithiothreitol 
			&#160;Abbreviation used in paper: DTT</td>
			<td>Thermo Fisher Scientific</td>
			<td>R0861</td>
			<td>Stock Concentration: 1 M in DMSO 
			Storage: dessicated, -20&#186;C 
			Stock and Experiment Recipes: dilute to 2 mM in PEM immediately before each experiment</td>
		</tr>
		<tr>
			<td>DMSO Anhydrous 
			&#160;Abbreviation used in paper: DMSO</td>
			<td>Thermo Fisher Scientific</td>
			<td>D12345</td>
			<td></td>
		</tr>
		<tr>
			<td>F-Buffer 
			&#160;Abbreviation used in paper: F-buffer</td>
			<td>NA</td>
			<td>NA</td>
			<td>Stock Concentration: 10x 
			Storage: dessicated, -20&#186;C 
			Stock and Experiment Recipes: 10 mM Imidazole (pH 7.0), 50 mM KCl, 1 mM MgCl2, 1 mM EGTA, 0.2 mM ATP</td>
		</tr>
		<tr>
			<td>G-Buffer 
			&#160;Abbreviation used in paper: G-buffer</td>
			<td>NA</td>
			<td>NA</td>
			<td>Stock Concentration: 10x 
			Storage: dessicated, -20&#186;C 
			Stock and Experiment Recipes: 2.0 mM Tris (pH 8), 0.2 mM ATP, 0.5 mM DTT, 0.1 mM CaCl2. Store at -20&#176;C.</td>
		</tr>
		<tr>
			<td>glass microscope slide 
			&#160;Abbreviation used in paper: slide</td>
			<td>Thermo Fisher Scientific</td>
			<td>22-310397</td>
			<td></td>
		</tr>
		<tr>
			<td>Glucose oxidase + catalase + &#946;-mercaptoethanol 
			&#160;Abbreviation used in paper: GOC</td>
			<td>Sigma Aldrich</td>
			<td>G2133-250KU, C1345, 63689&#160;</td>
			<td>Stock Concentration: 100x 
			Storage: store at stock concentration (100x) or 10x concentration, dessicated, at -20&#186;C 
			Stock and Experiment Recipes: For 100x: 4.3 mg/ml glucose oxidase, 0.7 mg/ml catalase, 0.5% v/v&#160; &#946;-mercaptoethanol in DI H20 
			Storage, Handling, Troubleshooting Notes: final concentration in solution should be: 0.005% &#946;-mercaptoethanol, 43 &#956;g/mL glucose oxidase, 7 &#956;g/mL catalase</td>
		</tr>
		<tr>
			<td>glu-GOC oxygen scavenging system 
			&#160;Abbreviation used in paper: glu-GOC</td>
			<td>NA</td>
			<td>NA</td>
			<td>Stock Concentration: 100x 
			Storage: prepare fresh each time 
			Stock and Experiment Recipes: mix equal parts Glu and GOC and add at 1/100 final sample volume immediately before imaging 
			Storage, Handling, Troubleshooting Notes: prepare from Glu and GOC immediately before imaging</td>
		</tr>
		<tr>
			<td>Guanosine triphosphate 
			&#160;Abbreviation used in paper: GTP</td>
			<td>Thermo Fisher Scientific</td>
			<td>R0461</td>
			<td>Stock Concentration: 100 mM 
			Storage: 100 &#956;L aliquots at -20&#186;C</td>
		</tr>
		<tr>
			<td>Instant Mix 1-minute epoxy 
			&#160;Abbreviation used in paper: epoxy</td>
			<td>Loctite</td>
			<td>1366072&#160;</td>
			<td></td>
		</tr>
		<tr>
			<td>Kinesin-1 401 BIO 6x HIS 
			&#160;Abbreviation used in paper: kinesin</td>
			<td>Prepared in JL Ross Lab at Syracuse University</td>
			<td>NA</td>
			<td>Stock Concentration: 8.87 &#956;M in PEM 
			Storage: single use aliquots, -80&#186;C, avoid freeze-thaw cycles 
			Storage, Handling, Troubleshooting Notes: biotinylated dimers form kinesin clusters, each aliquot can be used for up to 12 hrs stored on ice at 4&#186;C</td>
		</tr>
		<tr>
			<td>NeutrAvidin 
			&#160;Abbreviation used in paper: NA</td>
			<td>Thermo Fisher Scientific</td>
			<td>31000</td>
			<td>Stock Concentration: 5 mg/ml in PEM 
			Storage: dessicated, -20&#186;C 
			Stock and Experiment Recipes: reconstitute powder to 5 mg/ml in PEM</td>
		</tr>
		<tr>
			<td>No 1. glass coverslips (24 mm x 24 mm) 
			&#160;Abbreviation used in paper: coverslip</td>
			<td>Thermo Fisher Scientific</td>
			<td>12-548-CP</td>
			<td></td>
		</tr>
		<tr>
			<td>Paclitaxel&#160; 
			&#160;Abbreviation used in paper: Taxol</td>
			<td>Thermo Fisher Scientific</td>
			<td>P3456</td>
			<td>Stock Concentration: 2 mM in DMSO 
			Storage: protected from light, dessicated, -20&#186;C 
			Stock and Experiment Recipes: reconstitute to 2 mM with DMSO 
			Storage, Handling, Troubleshooting Notes: immediately prior to use dilute to 200 &#956;M in DMSO (0.4 &#956;L in 3.6 &#956;L DMSO)</td>
		</tr>
		<tr>
			<td>PEM-100 
			&#160;Abbreviation used in paper: PEM</td>
			<td>NA</td>
			<td>NA</td>
			<td>Stock Concentration: 1x 
			Storage: room temperature (RT) 
			Stock and Experiment Recipes: 100 mM K-PIPES (pH 6.8), 2 mM EGTA, 2 mM MgCl2 
			Storage, Handling, Troubleshooting Notes: use KOH to adjust pH to 6.8, recheck pH often and adjust accordingly</td>
		</tr>
		<tr>
			<td>phalloidin 
			Abbreviation used in paper: phalloidin</td>
			<td>Thermo Fisher Scientific</td>
			<td>P3457</td>
			<td>Stock Concentration: 100 &#956;M in DMSO 
			Storage: protected from light, dessicated, -20&#186;C, adhere closely to storage/handling conditions 
			Stock and Experiment Recipes: reconstitute to 100 &#956;M with DMSO 
			Storage, Handling, Troubleshooting Notes: susceptible to impurities in its preparation and denaturing, identifiable as large amorphous aggregates of actin in samples</td>
		</tr>
		<tr>
			<td>porcine brain tubulin 
			&#160;Abbreviation used in paper: tubulin</td>
			<td>Cytoskeleton</td>
			<td>T240</td>
			<td>Stock Concentration: 5 mg/ml in PEM 
			Storage: single use aliquots, -80&#186;C, avoid freeze-thaw cycles 
			Stock and Experiment Recipes: reconstitute powder to 5 mg/ml in PEM, flash freeze with LN2 
			Storage, Handling, Troubleshooting Notes: each aliquot can be used for up to 12 hrs stored on ice at 4&#186;C</td>
		</tr>
		<tr>
			<td>Potassium Chloride 
			&#160;Abbreviation used in paper: KCl</td>
			<td>Thermo Fisher Scientific</td>
			<td>AM9640G</td>
			<td>Stock Concentration: 4 M 
			Storage: RT</td>
		</tr>
		<tr>
			<td>Rabbit skeletal actin 
			&#160;Abbreviation used in paper: actin</td>
			<td>Cytoskeleton</td>
			<td>AKL99</td>
			<td>Stock Concentration: 2 mg/ml in G-buffer 
			Storage: single use aliquots, -80&#186;C, avoid freeze-thaw cycles 
			Stock and Experiment Recipes: reconstitute to 2 mg/ml in G-buffer, flash freeze with LN2 
			Storage, Handling, Troubleshooting Notes: once removed from -80&#186;C, store&#160; aliquot on ice at 4&#186;C, can be used for up to 1 week</td>
		</tr>
		<tr>
			<td>Rabbit skeletal myosin II 
			&#160;Abbreviation used in paper: myosin</td>
			<td>Cytoskeleton</td>
			<td>MY02</td>
			<td>Stock Concentration: 10 mg/ml in PEM 
			Storage: single use aliquots, -80&#186;C, avoid freeze-thaw cycles 
			Stock and Experiment Recipes: reconstitute powder to 10 mg/ml in PEM, flash freeze with LN2 
			Storage, Handling, Troubleshooting Notes: monomers form minifilaments at low KCl, each aliquot can be used for up to 12 hrs stored on ice at 4&#186;C</td>
		</tr>
		<tr>
			<td>Tubulin (biotin): porcine brain 
			&#160;Abbreviation used in paper: biotin-tubulin</td>
			<td>Cytoskeleton</td>
			<td>T333P</td>
			<td>Stock Concentration: 5 mg/ml in PEM 
			Storage: single use aliquots, -80&#186;C, avoid freeze-thaw cycles 
			Stock and Experiment Recipes: reconstitute powder to 5 mg/ml in PEM, flash freeze with LN2 
			Storage, Handling, Troubleshooting Notes: immediately prior to use dilute to 0.5 mg/ml in PEM</td>
		</tr>
		<tr>
			<td>Tubulin (fluorescent HiLyte 488): porcine brain 
			&#160;Abbreviation used in paper: 488-tubulin</td>
			<td>Cytoskeleton</td>
			<td>TL488M</td>
			<td>Stock Concentration: 5 mg/ml in PEM 
			Storage: single use aliquots, -80&#186;C, avoid freeze-thaw cycles 
			Stock and Experiment Recipes: reconstitute powder to 5 mg/ml in PEM, flash freeze with LN2 
			Storage, Handling, Troubleshooting Notes: each aliquot can be used for up to 12 hrs stored on ice at 4&#186;C, protect from light</td>
		</tr>
		<tr>
			<td>tubulin (rhodamine): porcine brain 
			&#160;Abbreviation used in paper: R-tubulin</td>
			<td>Cytoskeleton</td>
			<td>TL590M</td>
			<td>Stock Concentration: 5 mg/ml in PEM 
			Storage: single use aliquots, -80&#186;C, avoid freeze-thaw cycles 
			Stock and Experiment Recipes: reconstitute powder to 5 mg/ml in PEM, flash freeze with LN2 
			Storage, Handling, Troubleshooting Notes: each aliquot can be used for up to 12 hrs stored on ice at 4&#186;C, protect from light</td>
		</tr>
		<tr>
			<td>Tween 20 
			&#160;Abbreviation used in paper: Tween20</td>
			<td>Thermo Fisher Scientific</td>
			<td>J20605.AP</td>
			<td>Stock Concentration: 1% v/v in DI H20 
			Storage: RT</td>
		</tr>
		<tr>
			<td>ultracentrifuge grade microtubes 
			&#160;Abbreviation used in paper: Beckman-Coulter Optima Max XP&#160;</td>
			<td>Beckman Coultier</td>
			<td>343776</td>
			<td>Storage, Handling, Troubleshooting Notes: 8x34 mm PC</td>
		</tr>
		<tr>
			<td>UV light curing glue 
			&#160;Abbreviation used in paper: UV glue</td>
			<td>Pharda</td>
			<td>SKG-2869</td>
			<td></td>
		</tr>
	</tbody>
</table>

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.

Watch this Scientific Journal Video about Reconstituting and Characterizing Actin-Microtubule Composites with Tunable Motor-Driven Dynamics and Mechanics at JoVE.com

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

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 ...

reconstituting-characterizing-actin-microtubule-composites-with

Research

JoVE Journal

Biology

2.9K Views.  University of San Diego. 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. 

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

AC Electrokinetic Phenomena Generated by Microelectrode Structures

The field of AC electrokinetics is rapidly growing due to its ability to perform dynamic fluid and particle manipulation on the micro- and nano-scale, which is essential for Lab-on-a-Chip applications. AC electrokinetic phenomena use electric fields to generate forces that act on fluids or suspended particles (including those made of dielectric or biological material) and cause them to move in astonishing ways1, 2. Within a single channel, AC electrokinetics can accomplish many essential on-chip operations such as active micro-mixing, particle separation, particle positioning and micro-pattering. A single device may accomplish several of those operations by simply adjusting operating parameters such as frequency or amplitude of the applied voltage. Suitable electric fields can be readily created by micro-electrodes integrated into microchannels. It is clear from the tremendous growth in this field that AC electrokinetics will likely have a profound effect on healthcare diagnostics3-5, environmental monitoring6 and homeland security7.

In general, there are three AC Electrokinetic phenomena (AC electroosmosis, dielectrophoresis and AC electrothermal effect) each with unique dependencies on the operating parameters. A change in these operating parameters can cause one phenomena to become dominant over another, thus changing the particle or fluid behavior. 

It is difficult to predict the behavior of particles and fluids due to the complicated physics that underlie AC electrokinetics. It is the goal of this publication to explain the physics and elucidate particle and fluid behavior. Our analysis also covers how to fabricate the electrode structures that generate them, and how to interpret a wide number of experimental observations using several popular device designs. This video article will help scientists and engineers understand these phenomena and may encourage them to start using AC Electrokinetics in their research.

Manipulating fluids and suspended particles in the micro- and nano-scale is becoming more of a reality as enabling technologies, like AC electrokinetics, continue to develop. Here, we discuss the physics behind AC electrokinetics, how to fabricate these devices and how to interpret the experimental observations.

The field of AC electrokinetics is rapidly growing due to its ability to perform dynamic fluid and particle manipulation on the micro- and nano-scale, ...

Assembly, Loading, and Alignment of an Analytical Ultracentrifuge Sample Cell

The analytical ultracentrifuge (AUC) is a powerful biophysical tool that allows us to record macromolecular sedimentation profiles during high speed centrifugation. When properly planned and executed, an AUC sedimentation velocity or sedimentation equilibrium experiment can reveal a great deal about a protein in regards to size and shape, sample purity, sedimentation coefficient, oligomerization states and protein-protein interactions.
This technique, however, requires a rigorous level of technical attention. Sample cells hold a sectored center piece sandwiched between two window assemblies. They are sealed with a torque pressure of around 120-140 in/lbs. Reference buffer and sample are loaded into the centerpiece sectors and then after sealing, the cells are precisely aligned into a titanium rotor so that the optical detection systems scan both sample and reference buffer in the same radial path midline through each centerpiece sector while rotating at speeds of up to 60, 000 rpm and under very high vacuum
Not only is proper sample cell assembly critical, sample cell components are very expensive and must be properly cared for to ensure they are in optimum working condition in order to avoid leaks and breakage during experiments. Handle windows carefully, for even the slightest crack or scratch can lead to breakage in the centrifuge. The contact between centerpiece and windows must be as tight as possible; i.e. no Newton s rings should be visible after torque pressure is applied. Dust, lint, scratches and oils on either the windows or the centerpiece all compromise this contact and can very easily lead to leaking of solutions from one sector to another or leaking out of the centerpiece all together. Not only are precious samples lost, leaking of solutions during an experiment will cause an imbalance of pressure in the cell that often leads to broken windows and centerpieces. In addition, plug gaskets and housing plugs must be securely in place to avoid solutions being pulled out of the centerpiece sector through the loading holes by the high vacuum in the centrifuge chamber. Window liners and gaskets must be free of breaks and cracks that could cause movement resulting in broken windows.
This video will demonstrate our procedures of sample cell assembly, torque, loading and rotor alignment to help minimize component damage, solution leaking and breakage during the perfect AUC experiment.

The analytical ultracentrifuge (AUC) sample cell holds sample and reference buffer and during experiments and is exposed to high vacuum and rotor speeds up to 60,000 rpm. This video will demonstrate the rigorous attention to detail necessary for assembly, loading and alignment of this very important component of an AUC experiment.

The analytical ultracentrifuge (AUC) is a powerful biophysical tool that allows us to record macromolecular sedimentation profiles during high speed ...

Examining the Conformational Dynamics of Membrane Proteins in situ with Site-directed Fluorescence Labeling

Two electrode voltage clamp electrophysiology (TEVC) is a powerful tool to investigate the mechanism of ion transport1 for a wide variety of membrane proteins including ion channels2, ion pumps3, and transporters4. Recent developments have combined site-specific fluorophore labeling alongside TEVC to 
concurrently examine the conformational dynamics at specific residues and function of these proteins on the surface of single cells.
We will describe a method to study the conformational dynamics of membrane proteins by simultaneously monitoring fluorescence and current changes using voltage-clamp fluorometry. This approach can be used to examine the molecular motion of membrane proteins site-specifically following cysteine replacement and site-directed fluorophore labeling5,6. Furthermore, this method provides an approach to determine distance constraints between specific residues7,8. 
This is achieved by selectively attaching donor and acceptor fluorophores to two mutated cysteine residues of interest.
In brief, these experiments are performed following functional expression of the desired protein on the surface of Xenopus leavis oocytes. The large surface area of these oocytes enables facile functional measurements and a robust fluorescence signal5. It is also possible to readily change the extracellular conditions such as pH, ligand or cations/anions, which can provide further information on the mechanism of membrane proteins4. Finally, recent developments 
have also enabled the manipulation of select internal ions following co-expression with a second protein9.
Our protocol is described in multiple parts. First, cysteine scanning mutagenesis proceeded by fluorophore labeling is completed at residues located at the interface of the transmembrane and extracellular domains. Subsequent experiments are designed to identify residues which demonstrate large changes in fluorescence intensity (&lt;5%)3 upon a conformational change of the protein. Second, these changes in fluorescence intensity are compared to the kinetic parameters of 
the membrane protein in order to correlate the conformational dynamics to the function of the protein10. This enables a rigorous biophysical analysis of the molecular motion of the target protein. Lastly, two residues of the holoenzyme can be labeled with a donor and acceptor fluorophore in order to determine distance constraints using donor photodestruction methods. It is also possible to monitor the relative movement of protein subunits following labeling with a donor and acceptor fluorophore.

Examining the Conformational Dynamics of Membrane Proteins in situ with Site-directed Fluorescence Labeling

We will describe a method which measures the kinetics of ion transport of membrane proteins alongside site-specific analysis of conformational changes using fluorescence on single cells. This technique is adaptable to ion channels, transporters and ion pumps and can be utilized to determine distance constraints between protein subunits.

Two electrode voltage clamp electrophysiology (TEVC) is a powerful tool to investigate the mechanism of ion transport1 for a wide variety of membrane ...

Selective Capture of 5-hydroxymethylcytosine from Genomic DNA

5-methylcytosine (5-mC) constitutes ~2-8% of the total cytosines in human genomic DNA and impacts a broad range of biological functions, including gene expression, maintenance of genome integrity, parental imprinting, X-chromosome inactivation, regulation of development, aging, and cancer1. Recently, the presence of an oxidized 5-mC, 5-hydroxymethylcytosine (5-hmC), was discovered in mammalian cells, in particular in embryonic stem (ES) cells and neuronal cells2-4. 5-hmC is generated by oxidation of 5-mC catalyzed by TET family iron (II)/&alpha;-ketoglutarate-dependent dioxygenases2, 3. 5-hmC is proposed to be involved in the maintenance of embryonic stem (mES) cell, normal hematopoiesis and malignancies, and zygote development2, 5-10. To better understand the function of 5-hmC, a reliable and straightforward sequencing system is essential. Traditional bisulfite sequencing cannot distinguish 5-hmC from 5-mC11. To unravel the biology of 5-hmC, we have developed a highly efficient and selective chemical approach to label and capture 5-hmC, taking advantage of a bacteriophage enzyme that adds a glucose moiety to 5-hmC specifically12.
Here we describe a straightforward two-step procedure for selective chemical labeling of 5-hmC. In the first labeling step, 5-hmC in genomic DNA is labeled with a 6-azide-glucose catalyzed by &beta;-GT, a glucosyltransferase from T4 bacteriophage, in a way that transfers the 6-azide-glucose to 5-hmC from the modified cofactor, UDP-6-N3-Glc (6-N3UDPG). In the second step, biotinylation, a disulfide biotin linker is attached to the azide group by click chemistry. Both steps are highly specific and efficient, leading to complete labeling regardless of the abundance of 5-hmC in genomic regions and giving extremely low background. Following biotinylation of 5-hmC, the 5-hmC-containing DNA fragments are then selectively captured using streptavidin beads in a density-independent manner. The resulting 5-hmC-enriched DNA fragments could be used for downstream analyses, including next-generation sequencing.
Our selective labeling and capture protocol confers high sensitivity, applicable to any source of genomic DNA with variable/diverse 5-hmC abundances. Although the main purpose of this protocol is its downstream application (i.e., next-generation sequencing to map out the 5-hmC distribution in genome), it is compatible with single-molecule, real-time SMRT (DNA) sequencing, which is capable of delivering single-base resolution sequencing of 5-hmC.

Described is a two-step labeling process using &beta;-glucosyltransferase (&beta;-GT) to transfer an azide-glucose to 5-hmC, followed by click chemistry to transfer a biotin linker for easy and density-independent enrichment. This efficient and specific labeling method enables enrichment of 5-hmC with extremely low background and high-throughput epigenomic mapping via next-generation sequencing.

5-methylcytosine (5-mC) constitutes ~2-8% of the total cytosines in human genomic DNA and impacts a broad range of biological functions, including gene ...

Discovering Protein Interactions and Characterizing Protein Function Using HaloTag Technology

Research in proteomics has exploded in recent years with advances in mass spectrometry capabilities that have led to the characterization of numerous proteomes, including those from viruses, bacteria, and yeast.&#160; In comparison, analysis of the human proteome lags behind, partially due to the sheer number of proteins which must be studied, but also the complexity of networks and interactions these present. To specifically address the challenges of understanding the human proteome, we have developed HaloTag technology for protein isolation, particularly strong for isolation of multiprotein complexes and allowing more efficient capture of weak or transient interactions and/or proteins in low abundance. &#160;HaloTag is a genetically encoded protein fusion tag, designed for covalent, specific, and rapid immobilization or labelling of proteins with various ligands. Leveraging these properties, numerous applications for mammalian cells were developed to characterize protein function and here we present methodologies including: protein pull-downs used for discovery of novel interactions or functional assays, and cellular localization. We find significant advantages in the speed, specificity, and covalent capture of fusion proteins to surfaces for proteomic analysis as compared to other traditional non-covalent approaches. We demonstrate these and the broad utility of the technology using two important epigenetic proteins as examples, the human bromodomain protein BRD4, and histone deacetylase HDAC1.&#160; These examples demonstrate the power of this technology in enabling &#160;the discovery of novel interactions and characterizing cellular localization in eukaryotes, which will together further understanding of human functional proteomics. &#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;

HaloTag technology is a multifunctional technology which has shown significant success in isolation of both small and large protein complexes from mammalian cells.&#160; Here we highlight the advantages of this technology compared to existing alternatives and demonstrate its utility to study numerous aspects of protein function inside eukaryotic cells.

Research in proteomics has exploded in recent years with advances in mass spectrometry capabilities that have led to the characterization of numerous ...

Flash-and-Freeze: A Novel Technique to Capture Membrane Dynamics with Electron Microscopy

Cells constantly change their membrane architecture and protein distribution, but it is extremely difficult to visualize these events at a temporal and spatial resolution on the order of ms and nm, respectively. We have developed a time-resolved electron microscopy technique, &#34;flash-and-freeze,&#34; that induces cellular events with optogenetics and visualizes the resulting membrane dynamics by freezing cells at defined time points after stimulation. To demonstrate this technique, we expressed channelrhodopsin, a light-sensitive cation channel, in mouse hippocampal neurons. A flash of light stimulates neuronal activity and induces neurotransmitter release from synaptic terminals through the fusion of synaptic vesicles. The optogenetic stimulation of neurons is coupled with high-pressure freezing to follow morphological changes during synaptic transmission. Using a commercial instrument, we captured the fusion of synaptic vesicles and the recovery of the synaptic vesicle membrane. To visualize the sequence of events, large datasets were generated and analyzed blindly, since morphological changes were followed in different cells over time. Nevertheless, flash-and-freeze allows the visualization of membrane dynamics in electron micrographs with ms temporal resolution.

We developed a novel technique in electron microscopy, &#34;flash-and-freeze,&#34; that enables the visualization of membrane dynamics with ms temporal resolution. This technique combines the optogenetic stimulation of neurons with high-pressure freezing. Here, we demonstrate the procedures and describe the protocols in detail.

Cells constantly change their membrane architecture and protein distribution, but it is extremely difficult to visualize these events at a temporal and ...

Exploiting Live Imaging to Track Nuclei During Myoblast Differentiation and Fusion

Nuclear positioning within cells is important for multiple cellular processes in development and regeneration. The most intriguing example of nuclear positioning occurs during skeletal muscle differentiation. Muscle fibers (myofibers) are multinucleated cells formed by the fusion of muscle precursor cells (myoblasts) derived from muscle stem cells (satellite cells) that undergo proliferation and differentiation. Correct nuclear positioning within myofibers is required for the proper muscle regeneration and function. The common procedure to assess myoblast differentiation and myofiber formation relies on fixed cells analyzed by immunofluorescence, which impedes the study of nuclear movement and cell behavior over time. Here, we describe a method for the analysis of myoblast differentiation and myofiber formation by live cell imaging. We provide a software for automated nuclear tracking to obtain a high-throughput quantitative characterization of nuclear dynamics and myoblast behavior (i.e., the trajectory) during differentiation and fusion.

Skeletal muscle differentiation is a highly dynamic process, which particularly relies on nuclear positioning. Here, we describe a method to track nuclei movements by live cell imaging during myoblast differentiation and myotube formation and to perform a quantitative characterization of nuclei dynamics by extracting information from automatic tracking.

Nuclear positioning within cells is important for multiple cellular processes in development and regeneration. The most intriguing example of nuclear ...

Detecting and Characterizing Protein Self-Assembly In Vivo by Flow Cytometry

Protein self-assembly governs protein function and compartmentalizes cellular processes in space and time. Current methods to study it suffer from low-sensitivity, indirect read-outs, limited throughput, and/or population-level rather than single-cell resolution. We designed a flow cytometry-based single methodology that addresses all of these limitations: Distributed Amphifluoric FRET or DAmFRET. DAmFRET detects and quantifies protein self-assemblies by sensitized emission FRET in vivo, enables deployment across model systems-from yeast to human cells-and achieves sensitive, single-cell, high-throughput read-outs irrespective of protein localization or solubility.

This article describes a FRET-based flow cytometry protocol to quantify protein self-assembly in both S. cerevisiae and HEK293T cells.

Protein self-assembly governs protein function and compartmentalizes cellular processes in space and time. Current methods to study it suffer from ...

Oligomerization Dynamics of Cell Surface Receptors in Living Cells by Total Internal Reflection Fluorescence Microscopy Combined with Number and Brightness Analysis

Despite the importance and ubiquity of receptor oligomerization, few methods are applicable for detecting clustering events and measuring the degree of clustering. Here, we describe an imaging approach to determine the average oligomeric state of mEGFP-tagged-receptor homocomplexes in the membrane of living cells. The protocol is based on Total Internal Reflection Fluorescence (TIRF) microscopy combined with Number and Brightness (N&#38;B) analysis. N&#38;B is a method similar to fluorescence-correlation spectroscopy (FCS) and photon counting histogram (PCH), which are based on the statistical analysis of the fluctuations of the fluorescence intensity of fluorophores diffusing in and out of an illumination volume during an observation time. In particular, N&#38;B is a simplification of PCH to obtain information on the average number of proteins in oligomeric mixtures. The intensity fluctuation amplitudes are described by the molecular brightness of the fluorophore and the average number of fluorophores within the illumination volume. Thus, N&#38;B considers only the first and second moments of the amplitude distribution, namely, the mean intensity and the variance. This is, at the same time, the strength and the weakness of the method. Because only two moments are considered, N&#38;B cannot determine the molar fraction of unknown oligomers in a mixture, but it only estimates the average oligomerization state of the mixture. Nevertheless, it can be applied to relatively small time series (compared to other moment methods) of images of live cells on a pixel-by-pixel basis, simply by monitoring the time fluctuations of the fluorescence intensity. It reduces the effective time-per-pixel to a few microseconds, allowing acquisition in the time range of seconds to milliseconds, which is necessary for fast oligomerization kinetics. Finally, large cell areas as well as sub-cellular compartments can be explored.

We describe an imaging approach for the determination of the average oligomeric state of mEGFP-tagged-receptor oligomers induced by ligand binding in the plasma membrane of living cells. The protocol is based on Total Internal Reflection Fluorescence (TIRF) microscopy combined with Number and Brightness (N&#38;B) analysis.

Despite the importance and ubiquity of receptor oligomerization, few methods are applicable for detecting clustering events and measuring the degree of ...

Photodiode-Based Optical Imaging for Recording Network Dynamics with Single-Neuron Resolution in Non-Transgenic Invertebrates

The development of transgenic invertebrate preparations in which the activity of specifiable sets of neurons can be recorded and manipulated with light represents a revolutionary advance for studies of the neural basis of behavior. However, a downside of this development is its tendency to focus investigators on a very small number of &#8220;designer&#8221; organisms (e.g., C. elegans and Drosophila), potentially negatively impacting the pursuit of comparative studies across many species, which is needed for identifying general principles of network function. The present article illustrates how optical recording with voltage-sensitive dyes in the brains of non-transgenic gastropod species can be used to rapidly (i.e., within the time course of single experiments) reveal features of the functional organization of their neural networks with single-cell resolution. We outline in detail the dissection, staining, and recording methods used by our laboratory to obtain action potential traces from dozens to ~150 neurons during behaviorally relevant motor programs in the CNS of multiple gastropod species, including one new to neuroscience &#8211; the nudibranch Berghia stephanieae. Imaging is performed with absorbance voltage-sensitive dyes and a 464-element photodiode array that samples at 1,600 frames/second, fast enough to capture all action potentials generated by the recorded neurons. Multiple several-minute recordings can be obtained per preparation with little to no signal bleaching or phototoxicity. The raw optical data collected through the methods described can subsequently be analyzed through a variety of illustrated methods. Our optical recording approach can be readily used to probe network activity in a variety of non-transgenic species, making it well suited for comparative studies of how brains generate behavior.

This protocol presents a method for imaging neuronal population activity with single-cell resolution in non-transgenic invertebrate species using absorbance voltage-sensitive dyes and a photodiode array. This approach enables a rapid workflow, wherein imaging and analysis can be pursued over the course of a single day.

The development of transgenic invertebrate preparations in which the activity of specifiable sets of neurons can be recorded and manipulated with light ...